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CYTOLOGICAL AND MOLECULAR CHARACTERIZATION OF 
POPULATIONS OF THE ANOPHELES GAMBIAE GILES 
COMPLEX IN A COASTAL SAVANNA ZONE OF GHANA 
BY
JANET THERESA MIDEGA (B. Sc. HONS) Kenyafta University
A THESIS SUBMITTED TO THE AFRICAN REGIONAL POSTGRADUATE 
PROGRAMME IN INSECT SCIENCE (ARPPIS), UNIVERSITY OF GHANA, 
LEGON IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE 
AWARD OF MASTER OF PHILOSOPHY DEGREE IN ENTOMOLOGY
INSECT SCIENCE PROGRAMME*
UNIVERSITY OF GHANA
AUGUST 2001
♦JOINT INTERFACULTY INTERNATIONAL PROGRAMME FOR THE TRAINING 
OF ENTOMOLOGISTS IN WEST AFRICA 
COLLABORATING FACULTIES: AGRICULTURE AND SCIENCE
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DECLARATION
I hereby declare that with the exception of references to other peoples work which I have 
duly acknowledged all the experimental work described in Ihis thesis was carried out by 
me and this thesis, eilher in whole or in part, has not been presented elsewhere for 
another degree.
Janet T. Midega 
(Student)
Dr. Michael D. Wilson 
(Supervisor)
Dr. Daniel A. Boakye 
(Supervisor)
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DEDICATION
To My Parents, Ben and Julie, 
and To Professor K E  Senanu.
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ACKNOWLEDGEMENTS
I wish to express my sincere gratitude to all the individuals who through their support 
contributed to the successful completion of this dissertation.
I reserve my sincere gratitude to my supervisors, Dr. Michael D. Wilson and Dr. Daniel 
A. Boakye, both of the Parasitology Unit of the Noguchi Memorial Institute for Medical 
Research (NMIMR) for their direct involvement in this work. Their patience, expert 
guidance and treasured advice, concern, stimulating discussions and maximum co­
operation contributed greatly to the final result of this work. I am also graterul to Prof. 
W.Z. Coker of the Zoology Department for his excellent training in Medical 
Entomology, his contribution and advice throughout the course o f this research.
I also express my gratitude to the staff of NMIMR, who in various ways contributed 
towards the progress of this work. Special thanks to Mr. M A. Appawu who together 
with Dr. D.A. Boakye provided me with excellent training in cytotaxonomy, Mr. C. A  
Brown for his great contribution and brilliant ideas through out the laboratory work and 
data analysis. I want to express my gratitude and appreciation for his personal support, 
guidance and useful teaching. Special thanks to all the technicians particularly Messrs 
Sampson Otoo and Anthony Tetteh of Parasitology Unit for their substantial assistance 
during the fieldwork. I am equally grateful to Dr. K.A Koram and Mr. J.A Brandful.
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I would like to acknowledge the tremendous help, encouragement and friendly support 
given me by my colleagues at the parasitology laboratory, NMIMR. I am particularly 
indebted to Miss Nancy Duah, Naiki Puplampu, Helena Baidoo, Mrs. Anita Ghansah, 
Marian, Messrs Kofi Adasi, Kweku Boafo, Charles Brown, Neils Quashie and Bismark 
Sarfo. My utmost gratitude to Professor Ofori-Adjei, Director of the Institute, for the 
opportunity given me to work at the Institute.
I greatly appreciate the co-operation of Dr. V. Musewe, Head of capacity building, 
ICIPE, Kenya and the administration of the African Regional Postgraduate Programme 
in Insect Science (ARPPIS). I acknowledge the essential role played by Prof J.N 
Ayertey, coordinator, ARPPIS-West Africa in making this course a success. Special 
thanks to all my lecturers for helping me realize the potential in my work. My heartfelt 
thanks to all my classmates. I appreciate the companionship and useful discussions with 
Mr. Herve Bisseleua.
Finally, I express ray warmest obligation to the Midega family for their prayers while I 
was away working on this thesis. To the Senanu family, I can never thank you enough 
for your love and attention throughout the time I was in Ghana. May God richly bless 
you. To my friends, Lynda, Charity, Mabel, Miriam, Angie, Susan, Betty, Evelyn, Karis, 
Abanja, Too, Kitala, Isaac, Francis, Lepan and Tom, I express my appreciation for all 
the encouragement. Above all, I am most grateful to the almighty God for his provision.
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TABLE OF CONTENTS
DECLARATION...................................................................................................................1,1
DEDICATION...................................................................................................................... iv
ACKNOWLEDGEMENTS...................................................................................................v
TABLE OF CONTENTS......................................................................................................V1
LIST OF ILLUSTRATIONS................................................................................................«
LIST OF TABLES.................................................................................................................»
LIST OF PLATES................................................................................................................™
LIST OF APPENDICES................................................................................................... » »
LIST OF ABBREVIATIONS.............................................................................................xiv
ABSTRACT....................................................................................................................... xvii
CHAPTER ONE.................................................................................................................... 1
GENERAL INTRODUCTION..............................................................................................1
1.1 Introduction....................................................................................................... 1
1.2 Objectives.......................................................................................................... 9
CHAPTER 2......................................................................................................................... 10
LITERATURE REVIEW.....................................................................................................10
2.1 Malaria: The disease and symptomatology..........................................................10
2.2 Global Distribution.................................................................................................12
2.3 Social and Economic Impact of Malaria...............................................................14
2.4 The Life cycle and Transmission of human Plasmodium parasite.................... 15
2.5 The life cycle ofA nopheles vectors of malaria................................................... 20
2.6 Malaria control...................................................................................................... 25
2.6.1 Chemotherapy of malaria...............................................................................25
2.6.2 Vector Control.................................................................................................27
2.6.2.1 Current vector control interventions...................................................... 29
2.7 The Systematics of Anopheles gambiae complex................................................30
2.8 Distribution of Anopheles gambiae s.l..................................................................32
2.9 Methods used in the identification of Anopheles gambiae sibling.................... 34
species.......................................................................................................................... 34
2.9.1 Morphological identification..........................................................................35
2 9.2 Identification ofA nopheles gambiae based on reproductive...................... 36
incompatibility......................................................................................................... 36
2.9.3 Cuticular Hydrocarbon analysis.....................................................................36
2.9.4 Molecular methods for species identification and population studies........ 38
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2.9.4.1 Principles of PCR technology.................................................................39
2.9.5 PCR for the identification o f Anopheles gambiae species complex...........41
2.9.5.1 The ribosomal DNA method..................................................................41
2.9 5.2 Restriction Fragment Length Polymorphism (RFLP).......................... 42
2.9.5 3 Random Amplification of Polymorphic DNA (RAPD........................ 43
2.9.6. PCR for the studies of Anopheles gambiae populations............................ 44
2.9.6.1 Microsatellite DNA analysis...................................................................44
2.9.6.2 Applications of microsatellite analysis..................................................46
2.9.7 Cytotaxonomy.................................................................................................47
2.9.7.1 Polytene chromosomes and their value in cytogenetic studies........... 47
2.9.12 Chromosomal inversions........................................................................50
2.9.7.3 Inversion polymorphisms in the An. gambiae species......................... 52
CHAPTER THREE..............................................................................................................54
GENERAL MATERIALS AND METHODS....................................................................54
3.1 The Study Sites......................................................................................................54
3 .2 Field collection of mosquito larvae and pupae....................................................57
3 .3 Laboratory rearing of mosquitoes.........................................................................63
3.4 Morphological Identification of Female Anopheles Mosquitoes....................... 66
3 .4.1 Outline of criteria for the morphological features used for the identification 
of adult Anopheles gambiae s.l............................................................................... 66
3.5 Sample Preservation..............................................................................................67
3.6 Cytotaxonomy........................................................................................................68
3.6.1 Preparation of chromosomes for cytological studies................................... 68
3.6.2 Species identification by karyotyping...........................................................69
3.7 Molecular Studies..................................................................................................70
3.7.1 The extraction of genomic DNA...................................................................70
3 .7.2 PCR Identification of species of the Anopheles gambiae complex............ 70
3.7.3 Microsatellite DNA analysis..........................................................................73
3.7.4 Analysis of PCR products..............................................................................74
3.7.4.1 Agarose gel electrophoresis....................................................................74
3.7.4.2 Polyacrylamide gel electrophoresis.......................................................74
3.8 Data Analysis........................................................................................................ 76
3.8.1 Karyotype data................................................................................................76
3.8.2 Microsatellite data..........................................................................................76
CHAPTER FOUR............................................................................................................... 78
RESULTS............................................................................................................................ 78
4.1 Laboratory Rearing of Anopheles gambiae......................................................... 78
4.2 Karyotype analysis of Anopheles gambiae s.l......................................................78
4.3 PCR identification of Anopheles gambiae s.s......................................................84
4.4 Genetic structure of Anopheles gambiae populations........................................ 87
4.4.1 Allele and genotype frequency distributions............................................... 91
4.4.2 Population differentiation Index (Fst) ............................................................97
4.4.3 Estimate of heterozygote deficiency and excess (F1S) ................................. 98
4.4.4 Gene flow (Nm) ............................................................................................ 102
4.4.5 Phylogenetic relationship............................................................................ 104
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CHAPTER 5 ....................................................................................................................... 106
DISCUSSION AND CONCLUSION............................................................................... 106
REFERENCES...................................................................................................................113
APPENDICES.................................................................................................................... 139
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LIST OF ILLUSTRATIONS
Figure 1: World map showing the global distribution of malaria.
Figure 2: The life cycle of Plasmodium species.
Figure 3: The life cycle of Anopheles vectors of malaria.
Figure 4: The three chromosome pairs in the mitotic complement of Anopheles and 
the five elements seen in its polytene complement.
Figure 5: Diagrammatic representation of karyotypes expected from the eight 2R 
chromosome arrangements frequently observed in Anopheles gambiae. 
Figure 6: Map of the Greater Accra Region showing the study sites.
Figure 7: Chromosome complement o f An. gambiae showing the X chromosome 
(Xag) diagnostic for An. gambiae.
Figure 8: Standard Chromosome 2 R
Figure 9: Chromosome 2L showing the floating inversion 2La  with 180° rotation 
on section a.
Figure 10: Ethidium bromide stained 2%  agarose gel electrophoregram of genomic 
DNA extracted from An. gambiae species.
Figure 11: Ethidium bromide stained 2%  agarose gel electrophoregram of PCR 
products obtained from the amplification ofA n. gambiae DNA for 
species identification.
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Figure 12: Polyacrylamide gel electrophoregram of PCR products obtained from the 
amplification of An. gambiae s.s. microsatellite DNA with primer set 
AGXH7.
Figure 13: Polyacrylamide gel electrophoregram o f PCR products obtained from the 
amplification of An. gambiae s.s microsatellite DNA with primer set 
AG2H46.
Figure 14: Polyacrylamide gel electrophoregram of PCR products obtained from the 
amplification of An. gambiae s.s microsatellite DNA with primer set 
AG2H46
Figure 15: Allele frequency distribution at the AGXH7 locus
Figure 16: Allele frequency distribution at the AG2H26 locus
Figure 17: Allele frequency distribution at the AG2H46 locus
Figurel8: Estimates of F;s values indicating heterozygote deficiency and
heterozygote excess in the five^n. gambiae populations
Figure 19: A plot of gene flow between populations against distances between them.
Figure 20: Phenograms illustrating the genetic relationship between the Eve An.
gambiae populations.
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LIST OF TABLES
Table 1: DNA sequence details of the synthetic oligonucleotide primers used for 
the identification An. gambiae s.l. species and their melting temperatures.
Table 2: DNA Sequence details of the synthetic oligonucleotide primers used for the 
amplification o f An. gambiae s.s.
Table 3: Percent frequencies of karyotype 2La observed in An. gambiae s.s.
Table 4: Allele frequencies of most common alleles at all the three loci.
Table 5: Contingency X2 data for genotype frequencies among five An. gambiae 
s.s. populations
Table 6 : Estimates of differentiation (Frt), gene flow (Nm) and heterozygosity (Fb) 
at each locus
Table 7: Pairwise comparison of estimates differentiation Fa and gene flow (Nm) 
and genetic distances between An. gambiae populations
X I
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LIST OF PLATES
Plate 1: Sample collection site at Dodowa
Plate 2: Sample collection site at Adenta
Plate 3: Sample collection site at Madina
Plate 4: Sample collection site at East Legon
Plate 5: Sample collection site at Achimota
Plate 6: Polystyrene trays used in the laboratory rearing of Anopheles larvae
Plate 7: Wooden cages for holding emerging adult mosquitoes
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LIST OF APPENDICES
Appendix 1: Schedule of fixative and Stains.
Appendix II: Standard solutions
Appendix ID: An example input data format for the population genetics analysis 
software POPGENE Version 1.31.
Appendix IV: Standard polytene chromosome maps of Anopheles gambiae.
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LIST OF ABBREVIATIONS
AgN03 Silver Nitrate
bp base pair
dATP deoxyadenosine triphosphate
ddw distilled de-ionised water
dCTP deoxycytidine triphosphate
dGTP deoxyguanosine triphosphate
DNA deoxyribonucleic acid
dTTP deoxythymidine triphosphate
EDTA Disodium ethylene diamine tetraacetate. 2H2O
EthBr Ethidium Bromide
EtOH Ethanol
GPS Global positioning system.
H20 Water
KAc Potassium acetate
kb Kilobase
KOH Potassium hydroxide
M Molar
Mw Molecular weight
NaOH Sodium hydroxide
PCR Polymerase chain reaction
PH Hydrogen-ion exponent
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pH Hydrogen-ion exponent
RNA ribonucleic acid
RNase ribonuclease
rDNA ribosomal DNA
rpm revolution per minute
sddHzO sterile double distilled water
s.i Sensu lato
s.s Sensu stricto
TAE Tris - Acetate EDTA
TEMED N,N,N,’N ’- tetramethyl ethylene diamine
Tm Melting temperature
Tris 2  —amino-2-(hydroxymethyl)-l,3 propanediol
Hi microlitre
j*M micromolar
UPGMA Unweighted Pairgroup Method with Arithmetic Averaging
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ABSTRACT
The effective planning of vector control strategies, genetic manipulation of vector 
species and the management of insecticide resistance require information on the 
population structure and the assessment of gene flow between vector populations. 
Although the appetitive flight of mosquitoes is known to be less than 5 km, most studies 
assessing gene flow between An. gambiae populations have been conducted over 
geographical distances as far apart as 6000km. For vector control, operational purposes 
such as the use of insecticide treated materials within a particular country setting, the 
gene flow between populations that are some 10-100 Km is more important To assess 
the level of gene flow between populations over short distances, five populations of 
Anopheles mosquitoes from sites covering an area of about 25 km in diameter and 
located in the Coastal zone of Ghana were studied. One hundred and twenty-nine 
mosquitoes were identified by cytotaxonomy and PCR. A subset of these were fully 
kaiyotyped, but all were analyzed for three microsatellite DNA loci: AGXH7, AG2H26 
and AG2H46.
Karyotype analysis revealed the presence of the ‘Forest’ and ‘Savanna’ forms of An. 
gambiae s.s. characterized by the karyotypes 2R+, 2L+ and 2R+, 2La  respectively. The 
frequency of inversion a on the chromosomal arm 2L varied from 0% - 80% between 
populations that were furthest apart and differing in ecology from coastal forest to 
coastal savanna.
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A total of 30 polymorphic microsatellite alleles were observed. Number of alleles per 
locus ranged from nine to twelve (Mean = 10 per locus). Inter-population differentiation 
was highest between populations that were furthest apart (Frt = 0.0565) and lowest in 
closer populations (Fa = 0.0284). Accordingly, estimates of gene flow were higher in 
closer populations and lower in those further apart. Heterozygotes were in excess at the 
centrally located site (Fi„ = 0.0435) but deficient in others suggesting the presence of a 
possible hybrid zone in the study area.
The study established the presence of two distinct karyotypic populations of An. 
gambiae s.s in the coastal zone of Ghana and also revealed that levels of gene flow 
between populations decreased after 5km. The importance of karyotyping and 
microsatellite analysis in the characterization of vector populations is discussed.
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CHAPTER ONE
GENERAL INTRODUCTION 
1.1 Introduction
Mosquitoes are known vectors of parasites that cause tropical diseases such as malaria, 
lymphatic filariasis, yellow fever and arboviruses, e.g. dengue fever. Of these diseases, 
malaria is reported to be the most important worldwide (WHO, 1998) and still remains 
one of the world’s most prevalent vector bome disease and a leading cause of morbidity 
and mortality in the tropical countries. Malaria is caused by the infection of humans with 
either one or more of the protozoan parasites Plasmodium falciparum, P. ovale, P. 
malariae and P. vivax, transmitted through the bite of an infected female Anopheles 
mosquito (Service, 1993).
Over 400 species of Anopheles mosquitoes are known, most of which are members of 
species complexes, such as the^M. gambiae complex and An. funestus complex. Out of 
these, 80 species have been implicated in malaria transmission, but only 40 are important 
vectors (WHO, 1999). Members ofthe^w. gambiae complex, mainly An. gambiae sensu 
stricto and An. arabiensis are the most efficient vectors of malaria in sub-Saharan Africa, 
transmitting falciparum malaria which is the most common and the most lethal form of 
the disease (Coluzzi, 1984). Other known important vectors include An. pharoensis in 
Egypt, An. culicifacies in South West Asia, An. darlingi in North America and An. 
albimanus in Central America (Service, 1993).
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Previously widespread, the disease is now mainly confined to poorer tropical areas of 
Africa, Asia and Latin America and is endemic in a total of 101 countries, with pockets 
of transmission in 8 other countries (WHO, 1999). WHO, (1999) estimates that the 
disease represents 2.3% of the overall global disease burden and 9% in Africa, ranking 
third among major infectious disease threats after pneumococcal acute respiratory 
infections (3.5%) and tuberculosis (2 .8%). In Ghana, malaria is one of the major health 
problems. The disease is hyperendemic and accounts for over 43% of all out patient 
cases seen in health institutions and 44.1% of all clinic attendance. It is the most 
commonly reported disease responsible for 7-8% of all certified deaths and ranks 5* as 
the commonest cause of death in the 0-4 year age group (Centre for Health Statistics, 
1992).
Approximately, 41% of the world’s population is at risk and each year, there are an 
estimated 300 to 500 million clinical cases of malaria (more 95% of them in Africa), 
resulting in over 2 million deaths mainly of children under five years of age, especially 
in remote areas with poor access to health services. With acute disease, a child may die 
within 24 hours of infection. Pregnant women are four times more likely to suffer 
malaria attacks causing low weight babies and still births, endangering the health of the 
women and the prospects for the new bom (Lindsay et al., 2000). Outside tropical 
Africa, malaria deaths occur mainly among non-immune new comers to endemic areas, 
for example among agricultural workers, miners and settlers in newly colonized areas 
(WHO, 2000).
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Malaria poses as a serious obstacle to the socio economic development of tropical 
countries and interferes with industrial and agricultural development as it severely 
diminishes human work capacity, which consequently translates to food shortages and 
malnutrition. The direct and indirect costs of malaria are therefore very high and these, 
coupled with the burden of human suffering due to the disease call for its eradication. 
However, for economic and environmental reasons among others, it is not considered 
feasible to eradicate the disease from all parts of the world where it now occurs (Zahar, 
1984).
Malaria control therefore remains the only reliable option and an essential prerequisite 
for reducing the magnitude of this serious public health problem and for relieving the 
suffering of millions of people living in malaria endemic regions. Using current control 
methods it should be possible to reduce the severity and incidence of malaria to a level at 
which the disease ceases to be a public health problem The need to reduce malaria 
burden has recently led to two global initiatives; Multilateral Initiative on Malaria 
(MIM) and the Roll Back Malaria initiative (RBM). This need has also motivated 
research into the development of vaccines and the development of genetic control 
measures against the vector (Curtis, 1994; TDR 1998).
Previously, it was believed that the control and eradication of malaria would be easy. 
This belief was based on the assumption that the relationship between the parasite, the 
vector and the host was clearly understood, that effective therapeutic and 
chemoprophylactic agents were available and that insecticides held great promise for
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vector control (Beausoleil, 1984). However, decades later and after several pilot projects, 
reviews and revisions of strategies and various attempts at control, malaria still remains 
an insidious and ever-present scourge with serious public health, social, and economic 
consequences that constitute serious obstacles to development especially in rural areas 
(Rojas et al., 1992). Chief among the factors responsible for this situation are the 
climatic and ecological characteristics that favour vector survival and disease 
transmission (MARA, 1998) and the complexity of the vectorial system (Lanzaro et a l, 
1998; Lochouam and Fontenille, 1999), which consists of several sibling and incipient 
species.
Since the discovery of the European An. maculipennis complex (Hackett, 1937; Bates, 
1940), it has been repeatedly disco\ cred that human malaria vectors belong to cryptic or 
sibling species complexes, and that these, often sympatric species exhibit behavioural 
and ecological differences that influence their role as vectors (Coluzzi, 1970; Collins and 
Paskewitz, 1996). Thus, the failure of malariologists to recognize sibling species of 
Anopheles has meant in the past, failure to distinguish an epidemiologically important 
species from an unimportant one. ±vioreover, the studies of host preference, infection 
rate, resting habits, and biting cycle as well as the assessment of control measures were 
seriously misleading because the morphological species was in reality a mixture of two 
or mors spccics.
The current approaches to malaria control are chemotherapy, vector control and case 
management. Unfortunately these have also not achieved their expected levels of success
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due to limitations of development of resistance to anti-malarial drugs and 
chemoprophylactics by the parasites (Peters, 1970), insecticide resistance by the vector 
(Ranson et al., 2000) and disease resurgence in areas where the disease had been 
previously eradicated (Krogstard, 1996).
To date, the only malaria control strategy that has achieved some success were those 
aimed at the vector using insecticides (TDR, 1998). Insecticide application methods used 
include indoor insecticide spraying, aerial spraying, residual insecticide application on 
walls and surfaces, the use of insecticide treated materials such as curtains and bednets 
and the application of larvicides at mosquito breeding sites. Currently, the RBM 
initiative has adopted the promotion of the use of insecticide treated bednets (iTBNs) 
using pyrethroids in malaria endemic regions to reduce human-vector contact This has 
been shown to provide a remarkable degree of protection against malaria in Africa 
(Binka et al., 1996, Curtis et al., 1998). Recent studies by Abdulla et al. (2001) in 
Ifakara, Tanzania revealed that the use of ITBNs had a protective efficacy of 62% on the 
prevalence of parasitaemia. However, reports of reduced susceptibility o f An. gambiae to 
permethrin associated with the use of permethrin impregnated bednets and curtains in 
both East and West Africa (Vulule et al., 1994, 1996; Doannio et al., 1999) may reduce 
the effectiveness of ITBNs as a vector control tool. Insecticide resistance arising in a 
local vector population as a result of selection pressure due to insecticide use may spread 
to adjacent populations by gene flow, widening the distribution of the gene for 
resistance. For the effective management of insecticide resistance therefore, knowledge
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of the range of distribution of vector populations and gene flow patterns is a paramount 
requisite for any malaria control programme.
Apart from the problem of insecticide resistance, the diversity of the vectorial system 
has also contributed to the lack of success of most vector control operations (Coluzzi, 
1992). Formerly regarded as a single species, the afrotropical vectors of malaria are 
members of the An. gambiae Giles complex, which is now known to comprise of 6 
sibling species (Gillies and Coetzee, 1987) one unnamed species (Hunt etal., 1998) and 
several incipient species (Coluzzi et a l, 1979; Favia et al., 1997). These vectors 
generally cause high parasite inoculation rates and are also remarkably stable in a wide 
range of bio-ecological and seasonal conditions hence appear to be very flexible, both in 
exploiting new man-made environments and in their response to malaria control 
activities (Coluzzi, 1984).
The adaptability to environmental changes leading to marked contrasts in vector 
bionomics has led to the development of various levels of vectorial efficiency for 
populations ofA nopheles species in heterogenous environments within the same locality 
and has thus become an important factor in the determination of the epidemiology of 
malaria (Toure et al., 1994). Environmental heterogeneities have arisen mainly as a 
result of human activities which act as a means of constant evolutionary challenge as 
they provide a source of environmental change to which anthropophilic Anopheles have 
to respond by developing a highly dynamic vector-host relationship.
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The monitoring of processes such as vector movement and distribution requires an 
intimate knowledge of vcctcr behavioural attributes such as ho3t seeking and choice of 
resting and oviposition sites that may differ between sibling species yet are a prerequisite 
for the implementation of vector control (Coluzzi et al., 1979). In this respect, the 
identification of sibling species is a problem that needs to be solved before the 
bionomics of these species can be studied. The identification of sibling and incipient 
species and population characterisation is therefore necessary because vector control 
strategies must be specific to target the exact vector. A necessary condition for this is 
reference to taxonomic units; species, sub-species, forms and populations that have 
biological significance. Various methods exist in Anopheles studies that can be applied 
for all these taxonomic levels. These include the conventional morphological methods, 
cuticular hydrocarbon analysis, crossing experiments, biochemical methods, 
cytotaxonomy (Coluzzi, 1968; Coluzzi and Sabatini, 1967,1968 a; 1969) and molecular 
methods (Scott et al., 1993, Lanzaro et al., 1995).
To gain a better understanding of the population dynamics, gene flow and possible 
influence of ecological processes on the genetic variability in An. gambiae s.l. species 
among local populations within and between geographic regions, it is important to 
identify and characterize the vector populations, as this is important in the interpretation 
of studies on vector biology.
One of the methods most commonly used for An. gambiae species identification is 
cytotaxonomy which exploits the use of floating chromosomal inversions to delineate 
geographic distribution of An. gambiae s.s. populations (Coluzzi et al., 1979/
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Microsatellite DNA analysis has been widely applied in studies of An. gambiae 
population differentiation and assessment of gene flow between populations (Kamau et 
al., 1998).
The current malaria control activities in Ghana involve the use of chemotherapy and case 
management for disease control and no systematic use of insecticides for vector control. 
However, Ghana has adopted the Roll Back Malaria strategies and intends to promote 
large-scale use of ITBNs. However, there have been reports of pyrethroid resistance in 
malaria vectors in some parts of West Africa including the neighbouring Cote d’ Ivoire 
(Chandre et al., 1999 a, b, c) and in Accra (Adasi et al., 2001) in the absence of any 
organised vector control programmes. Although for the Accra study mosquitoes were 
collected from specific sites, pyrethroid sensitivity tests were carried out on pooled 
samples. It is therefore likely that pyrethroid resistance in mosquitoes is present at 
various levels at certain sites and not at others. This however, does not preclude the 
possibility that eventually pyrethroid resistance will spread to all mosquito populations 
in Accra through gene flow and human aided movement of the vectors.
The effective management of the existing pyrethroid resistance in Accra will require the 
characterization of the vector species and populations and also identification of the 
specific areas where resistance has developed. This will prevent the problem of 
misdirected vector control efforts, which have previously led to the failure of malaria 
control programmes based on vector control.
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1.2 Objectives
The main aim of this study was to identify and characterise Anopheles vector 
populations within an area covering approximately 25 kms across to determine the levels 
of population differentiation and gene flow between them. This was to be achieved by 
the use of karyotyping and microsatellite DNA analysis, to reveal intraspecific 
differences between An. gambiae s.s. populations.
Specific Objectives
The specific objectives were:
1. To collect and identify members of the An. gambiae Giles complex from selected sites
within a 25 km radius in the coastal savanna area of Ghana.
2. To characterize the^w. gambiae sensu stricto populations by use of karyotyping
3. To study the An. gambiae s.s populations using PCR- based microsatellite DNA 
analysis.
4. To determine the population structure of the ̂ 4n. gambiae s.s. populations.
5. To determine the levels of gene flow between the yin. gambiae s.s. populations.
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CHAPTER 2
LITERATURE REVIEW 
2.1 Malaria: The disease and symptomatology
Malaria is a febrile illness caused by protozoa of the class sporozoa, genus Plasmodium 
(Farr, 1988). The parasite is transmitted from human to human through the bites of 
infected female Anopheles mosquitoes. In endemic regions where transmission is high, 
people are continuously infected and some may develop immunity to the disease 
(Allison, 1984). Malaria is often described as a great imitator of other diseases, sharing 
various characteristics with other childhood illnesses like influenza, typhoid, brucellosis 
and urinary tract infections. Even more confusing, malaria may coexist with other 
diseases. Most of the morbidity due to malaria is caused by cerebral malaria and anaemia 
(Marsh, 1992). Malaria is diagnosed by the clinical symptoms and microscopic 
examination of blood films.
The symptoms first appear some 10 to 16 days after the infectious mosquito bite and 
coincide with the bursting of infected red blood cells. When many red blood cells are 
infected and break at the same time because of the parasites growth, malaria attacks can 
recur at regular time periods -  every two days for P. vivax malaria and P. ovale and 
every three days for P. malariae (Berendt et al., 1994). In P. vivax malaria, the patient 
may feel entirely well between attacks. Even without treatment, the paroxysms subside 
in a few weeks. However, a person with P. falciparum malaria is apt to feel very ill
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between attacks and without treatment, is in danger of dying. One reason why P. 
falciparum malaria is so virulent is that the parasite can infect red blood cells in all 
stages of development, and the number of parasites in the blood can reach extremely 
high levels (Bruce-Chwatt, 1985). In contrast, P. vivax parasites infect only young red 
blood cells which means the number of parasites in the blood does not reach the same 
high levels as seen in P. falciparum infection.
Malaria typically produces a string of recurrent attacks or paroxysms, each of which has 
three stages: chills, followed by fever, pain in the joints and then sweating 
(Cheesbrough, 1991). Along with chills, the patient is likely to have headache, nausea 
and repeated vomiting (Commey, 1989) and within an hour or two, the patient’s 
temperature may rise. One of the commonest causes of death in children and adults with 
falciparum malaria is cerebral malaria, which is due to parasitized red cells and fibrins 
blocking the capillaries and venules in the brain (Marsh, 1992). Cerebral oedema and 
peri-capillary haemorrhaging from these small blood vessels may occur, causing 
irreversible injury to the brain (Wairel et al, 1982). In adults, cerebral malaria often 
develops after several days of fever and other non-specific symptoms (Wanell, 1983; 
Molyneux et al., 1989), but in children, the history is usually less than two days. 
Cerebral malaria often starts with a generalized convulsion followed by persisting 
unconsciousness. It is frequently associated with major metabolic and physiologic 
changes, particularly hypoglycaemia, decreased cerebral perfusion/hypoxia and cytokine 
induction, resulting in tissue damage and generalized seizures, paralysis, speech and
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behaviour disorders and hearing impairment (MDM, 2001). Severe anaemia is often the 
attributable cause of death in areas with intense malaria transmission (Commey, 1989).
2.2 Global Distribution
It is estimated that over 40% of the world’s population lives in areas with malaria risk, 
with the great majority living in Sub-Saharan Africa (WHO, 2000). The prevalence and 
severity of malaria as well as the magnitude of associated social and economic effects 
vary widely in different geographical areas where the disease occurs. However, the 
worst effects of the disease are felt in Sub-Saharan Africa.
The geographical area affected by malaria has shrunk considerably over the past 50 
years, with successful eradication and cessation of transmission in large areas of North 
America, Southern Europe, and the former Soviet Union and some territories of Asia and 
South America (WHO, 1998). The disease is now confined to poorer tropical areas of 
Africa, Asia and Latin America because of the tropical climate, which is suitable for 
vector development and survival (Figure 1).
Malaria is endemic throughout Ghana (Ahmed, 1989) and records show that more cases 
of malaria occur in the middle forest zone (Ashanti, Brong Ahafo, Eastern and greater 
parts of Western Regions). This is followed by the coastal zone (Central, Greater Accra 
and parts of Western regions), which is a terrain of lagoons, creeks, etc and then by the 
northern savanna zone, which is usually drier than the rest of the country.
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2.3 Social and Economic Impact of Malaria
Recent estimates put the annual economic burden of malaria in Africa at about $11.7 
billion or 1 % of the gross domestic product. The cost of a case from society’s view point 
is $9.84 or 12 days equivalent of productivity and the cost of treatment per household 
ranges from $0.2-$ 15 each month in Africa. In 1998, it was reported that 39,000,000 
daily adjusted life years (DALYs) were lost as a result of illness due to malaria (WHO, 
1999).
The disease generally affects all age groups, but the most affected are children (Breman 
and Campbell, 1988) and pregnant women (Lindsay et al., 2000). Over 300-500 million 
people suffer from the disease annually and mortality due to malaria has been estimated 
to account for 2 million deaths per year, half of these being in children under five years 
of age in Africa (TDR Progress Report, 1997-98). In cases where the disease affects 
working adults such as labourers, loss in income is inclined due to lost labour hours and 
cost of treatment. Workers suffering a bout can be incapacitated for 5-20 days. Malaria 
stricken family spends an average of over one-quarter of its income on malaria 
treatment, and prevention. It is estimated that such a family can only harvest 40% of 
crops harvested by healthy families (WHO, 1990).
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2.4 The Life cycle and Transmission of Human Plasmodium Parasite
Four protozoan parasites, namely P. falciparum, P. ovale, P. malariae and P. vivax 
cause malaria in humans and they all have the similar life cycle (Figure 2).
During the infective bite of a female Anopheles mosquito numerous infective sporozoites 
are injected into the bloodstream, where they mat remain for about 30 minutes and then 
disappear from the blood stream. Most of the sporozoites are destroyed by phagocytes 
whilst some enter the liver parenchymal cells, develop and undergo asexual 
multiplication (pre-eiythrocytic schizogony). A large unpigmented schizont is formed 
containing several merozoites, which are released into the bloodstream to invade the 
erythrocytes. The merozoite attaches to an erythrocyte and is invaginated into the red 
cell through a parasitophorous vacuole, where it feeds and deposits a pigment called 
haemozoin as a by product (Aikawa, 1980). The ingested merozoite becomes a feeding 
trophozoite and in the early stages of an infection, the fully-grown trophozoite multiplies 
asexually to become a schizont (erythrocytic schizogony), producing a small number of 
merozoites (Aikawa and Seed, 1980).
Release of the merozoites from the erythrocytes brings on an attack of malaria, and the 
interval between attacks is the length of the schizogonic cycle. This may last several 
hours. The released merozoites repeat the cycle and invade other erythrocytes. After a 
number of cycles of schizogony, the trophozoites do not divide but become gametocytes, 
which develop no further in man but circulate in the bloodstream until a mosquito takes 
them up during a blood meal (Ketde, 1992).
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When in the Anopheles mosquito, microgametocytes exflagellate to become 
microgametes. The microgametes move away to find and fuse with macrogametes to 
form zygotes, which remain motionless for 18-24 hours then elongate to form ookinetes 
(Aikawa and Seed, 1980; Kettle, 1992). The ookinete penetrates the wall of the midgut 
and forms an oocyst and through sporogony, the oocyst develops into enlarged motile 
sporozoites, which burst through the ruptured wall of the oocyst and invade the body 
cavity of the mosquito. The sporozoites, which are now infective then migrate to the 
salivary glands of the mosquito and are injected into the wound when the mosquito takes 
the next blood meal.
The effective transmission, high incidence and mortality of malaria is determined by a 
number of factors which affect the distribution, transmission intensity, disease outcome, 
small-scale variation and seasonality (MARA, 1998).
The number and type of anopheline mosquitoes determine to a large extent the level of 
transmission in a given area. Malaria transmission is influenced by climate and 
geography and often coincides with the rainy season (Craig et al., 1999) when breeding 
sites are available, ensuring high numbers of Anopheles. The climatic conditions in 
tropical Africa favour intensive transmission of malaria parasites. Temperature affects 
the sporogonic cycle of the parasite and vector survival. Very low temperatures limit the 
vector by lengthening the gonotropic cycle and reducing the survival rate. Anopheles 
gambiae for example, only exists in frost-free regions (de Meillon, 1934) or where
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absolute minimum temperatures in winter remain above 5°C (Leeson, 1931). Rainfall 
provides mosquito-breeding sites and increases humidity, thus enhancing mosquito 
survival. However, the relationship between mosquito abundance and rainfall is not 
direct, as a specific amount of rain does not lead to a specific mosquito density. By 
examining rainfall patterns in known malaria and non-malarious areas, Craig et al. 
(1999) estimated that an average of 80mm per month for at least three to five months is a 
reasonable requirement for the availability of mosquito breeding sites to sustain malaria 
transmission.
Poor environmental sanitation, like poor drainage systems also increase mosquito- 
breeding sites. This, in addition to development projects in pursuit of economic 
development such as water schemes, construction of dams and bridges, oil drilling and 
mining activities, urban planning and development, logging activities inadvertently lead 
to an increase in mosquito breeding sites, thereby increasing their numbers, human- 
mosquito contact and therefore transmission (Okenu, 1999).
Historically, population movement has also contributed to the spread of the disease 
(Prothero, 1977). The unprecedented increase in mobility in the last few decades has led 
to the relationship between mobility and malaria. The movement of infected people from 
areas where malaria is still endemic to areas where the disease had been eradicated has 
led to the resurgence of the disease (Gubler, 1998). For example, in 1985,26 new active 
foci of malaria were introduced in Brazilian states outside the Amazon region (Marques, 
1986). As people move, they increase their risk for acquiring the disease through the
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ways in which they change the environment and through the technology they introduce, 
for example through deforestation for resettlement and irrigation systems (Service,
1991). Such environments create more favorable mosquito breeding habitats and at the 
same time, workers may have increased exposure to the vector. Furthermore, people can 
inadvertently transport infected mosquitoes to malaria free areas, reintroducing the 
disease (Martens and Hall, 2000). Population movement is also intensely implicated in 
the spread of drug resistance in malaria (Rajagopalan et a l,  1986), with people carrying 
resistant parasites moving from endemic to non-endemic disease areas and accelerating 
transmission of resistant strains.
Weak health systems, poor strategic development and inadequate funding of control 
programmes all play a role in the transmission of malaria. Recent rural to urban 
migration in search of economic and educational opportunities has created densely 
populated communities, especially in peri-urban areas and weakened health systems with 
the attendant increase in human activity and adverse effects on the environment, thus 
creating breeding sites. This promotes vector abundance leading to increased human- 
vector contact.
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Vporofotin
A ^
I i \ r r  s!>t*r ^  * • • / j
/ &- >
(  V - L . J  l i i f e d t r i  /
flrpiMoc?1«H
V1rt-n/oil<*s
, //   * '• * ,•'  * /*> — ~ ^ /  AJ
I‘ J /  *  ••. { ) " <
/  ̂ {/*!*-V*  \1  o#/’  ^, 112 n̂rin Ganrnorytes i
Srhizmils ( ^  | SniiJHl tT.vdirccytir lorin«
' *i• ’•B#•  /•*». ('  •'J»  T1 
— />;,
v ™  I V o p lit r o lm
A-Strut*! rryUiroryficslajB?
Figure 2: Schematic representation of the life cycle of Plasmodium species (After 
Gamharm, 1966)
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2.5 The Life Cycle of Anopheles Vectors of Malaria
The complete cycle from egg to adult takes a minimum of about 2 weeks or slightly less. 
Figure 3 shows the life cycle ofA nopheles mosquitoes.
The breeding habitats vary from large and usually permanent collections of water, such 
as fresh water swamps, marshes, rice fields and borrow pits to smaller collections of 
temporary water such as small pools, puddles, water filled car tracks, ditches, drains, 
gulleys, hoof prints, etc. The most common breeding sites are the shallow open sun lit 
pools (Service, 1993). Larvae also occur in wells and manmade container habitats such 
as clay pots, motor vehicle lyres, water storage jars and tin cans (Chinery, 1984). 
However Anopheles gambiae s.s prefer small and undisturbed temporary pools of water 
exposed to sunshine (Muirhead Thompson, 1945)
The female An. gambiae after mating and blood feeding lays some 50-200 small (1mm 
long) brown or blackish boat shaped eggs on the water surface Viable eggs hatch into 
larvae within 2-3 days in the tropics, but in cooler temperate regions they may not hatch 
until after 4 -7 days or longer (Service, 1980). The larvae, while on the water, lie parallel 
to the surface to allow water intake and surface feeding
At mean water temperatures of 25-28° C the larvae undergo four moults within 6-9 days 
to reach the pupal stage, which lasts 2-3 days depending on temperature. Thus, the 
minimum duration for one generation may be as long as 10-11. The pupae bear
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respiratory trumpets that are short and broad distally thus appearing conical. The most 
distinctive characteristic of Anopheles pupae is the presence of short peg like spines 
situated laterally near the distal margins of abdominal segments. The pupal skin splits 
dorsally and the adult emerges. Careful movements are required to ensure that the adult 
mosquito does not fell sideways and be trapped in the surface film. This danger is 
particularly acute when the adult is largely out of the pupal exuviae but the terminal 
appendages are still not free. Finally the legs become free and spread on the water 
surface giving stability. The newly emerged adult inflates its wings, and separates and 
grooms its head appendages before flying away (Kettle, 1992). When the progeny of any 
one egg batch emerge as adults the males emerge first The males become ready for 
mating within 24 hours after emergence such that by the time the females emerge, the 
males are competent for mating. Mating is often preceded or accompanied by swarming 
in which the males associate over a marker and fly in a particular manner. Most of the 
male mosquitoes usually die after mating. The females require a blood meal for ovarian 
development, followed by the maturation and oviposition of a batch of eggs (Gillies, 
1955).
The percentage of the eggs that goes to form the adults is unknown, but there is usually 
heavy mortality, especially among larvae due to predators, disease, drought, flood, etc. 
Larval loss due to predation is one of the factors that reduce the numbers of larvae that 
develop into adults. It is recognized that predation of larvae in established pools is an 
important factor in limiting their numbers. In some instances, Culex tigripes colonizes 
the same pools as An. gambiae, causing a dramatic reduction in larval density (Haddow,
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1942). In permanent wells in Tanzania, Christie (1958) concluded that predation 
pressure was so in tense that few larvae survived to pupae. Notonectidae were among the 
most important predators in these wells. It is possible that the same pressures exist in 
other types of permanent waters, thus limiting their productivity for An. gambiae. It may 
be noted that the vagility often displayed by An. gambiae larvae, in contrast to species 
such as An. funestus, would tend to increase their vulnerability to attack by predators 
(Service, 1980).
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MOSQUITO LIFE CYCLE
A dult Fem ale Feeds
Fenules Lay
Adults Eggs
Emerge
L arvae
D evelop
Figure 3: Schematic illustration of the life cycle of Anopheles vectors of malaria 
(After Service, 1980).
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Adult Anopheles usually rests with the body at an angle to the surface, with proboscis 
and abdomen in a straight line. In some species, they rest at almost right angles to the 
surface, whereas in others such as An. culicifacies the angles are much smaller (Kettle,
1992). "Most Anopheles mosquitoes have dark and pale scales on the wing veins arranged 
in blocks, forming a distinctive spotted pattern, which varies between species. Female 
mosquitoes have non-plumose antennae while males have plumose antennae. The palps 
of adult anopheline females are about as long as the proboscis and usually lie closely 
alongside it and may be marked, especially the apical half with broad and narrow rings 
of pale scales. In males the palps are distinctly swollen at the ends, and may also have 
apical rings of pale scales.
Most Anopheles are crepuscular or nocturnal in their activities, thus emergence from the 
pupae, mating, blood feeding and oviposition normally occur in the evenings, at night or 
early in the morning around sunrise. Some species such as Anopheles albimanus, a 
malaria vector in Central and South America, bite man mainly outdoors (exophagic) 
from about sunset to 2100 hours whereas in Africa species of the An. gambiae complex 
bite mainly after 2100 hours and mostly indoors (endophagic). Some species will rest in 
houses (endophilic) in between feeding whereas others will rest outside (exophilic) in a 
variety of natural shelters, such as amongst vegetation, in rodent burrows, cracks and 
crevices in trees, under bridges, in termite mounds, and other cracks in the ground. Most 
Anopheles species are not exclusively exophagic or endophagic, exophilic or endophilic, 
but exhibit a mixture of these extremes of behavior. Similarly, few Anopheles feed
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exclusively on either man or humans, most feed on both man and animals but the degree 
of anthropophilism and zoophilism varies according to species (Gillies et al., 2000).
2.6 Malaria Control
The strategy of malaria control is based on breaking the chain of transmission of the 
parasites between humans and mosquitoes. There are presently two approaches: control 
of parasites by chemotherapy and chemoprophylaxis and by breaking the contact 
between human host and vectors by controlling the vector by using insecticides, bednets 
and environmental management.
2.6.1 Chemotherapy of malaria
Chemotherapy in malaria control was started as early as the 1930’s with the discovery of 
the aetiology and mode of transmission of the disease (Hackett, 1937). When antivector 
measures failed to significantly reduce the effects of the disease in endemic areas despite 
some early successes, it became evident that there was still heavy reliance on 
chemotherapy. Quinine is the drug that was widely used at the time but had undesirous 
side effects. Thus the advent of new and effective therapeutic drugs particularly the 4- 
aminoquinolines in the 1940’s provided for the first time, reliable and acceptable means 
for treating malaria and for malaria chemoprophylaxis (Jeffery, 1984). Chloroquine
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became the antimalarial treatment of choice because it was safe, inexpensive and highly 
effective against susceptible malaria parasites (WHO, 1967; Djimde et ah, 2001). Thus, 
malaria chemotherapy assumed a major role in primary health care because of the 
rationale of preventing mortality and curbing morbidity and suffering of afflicted 
persons. However, the efficacy of chloroquine in malaria chemotherapy has been 
compromised with the development of resistance to the drug by malaria parasites. Of the 
four Plasmodium species infective to man that have developed drug resistance, only P. 
falciparum poses a very serious clinical problem. The first documented cases of P. 
falciparum resistance to chloroquine were in South America in the 1950’s (Peter, 1970), 
and confirmed in Thailand in 1959 (Harinasuta et al, 1962). Since then chloroquine 
resistance spread to almost all the malarious areas of the world with heterogenic 
distribution both in frequency and degree (Bjorkman and Philip-Howard, 1990). In 
Africa, resistant strains of P. falciparum were first recorded in Kenya and Sudan in 1978 
(WHO, 1986) and more recently in West Africa (Cheesbrough, 1991). In many parts of 
Africa, the drug is no longer used alone for therapy (Brasseur et al., 1998). 
Chemotherapy has also been complicated further by the development and spread of 
resistance to other antimalarials such as Fansidar (Watkins and Mosobo, 1993), which is 
known to be heavily influenced by a combination of factors; Human migration, heavy 
malaria transmission, severe infection, immune response to malaria and sustained and/or 
haphazard use of the drug. Currently, drug combination therapy is recommended for 
malaria control in areas where drug resistance is highly prevalent.
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2.6.2 Vector control
Malaria control with a vector control component has a long history in the African region. 
Prior to the Second World War, control was based mainly on antilarval measures, 
including source reduction, while pyrethrum spray as an adulticide was tried on a small 
scale in certain areas with variable results (Zahar, 1984). During the 1940’s when the 
organochlorine insecticide Dichlorodiphenyltrichloroethane (DDT) became available 
during the Second World War, malaria control by house spraying was initiated on a 
small scale in certain countries and on a larger scale in Madagascar, Mauritius, South 
Africa (Natal and Transvaal), Swaziland, and Zimbabwe, formerly Southern Rhodesia 
(Bruce-Chwatt, 1963). When the global malaria eradication programme was initiated by 
the WHO in 1955 the WHO African Region was excluded because of the problem of 
logistics requirement associated with vector control, which were beyond the scope o ft he 
vast majority of the African countries and also because of intense transmission of P. 
falciparum by An. gambiae s i ,  and An. funestus (WHO, 1999). To overcome this 
problem, a reorientation of control activities was undertaken in certain countries and 
islands, and eradication pilot projects were circumscribed in areas in several countries 
from the mid-1950’s to the early 1960’s (Bruce-Chwatt, 1963; NIAID, 2000). These 
projects, initiated by the WHO Malaria Eradication Programme between 1955 and 1969 
were aimed at complete interruption of malaria transmission as a pre-requisite for 
malaria eradication.
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The main approach was residual house spraying using DDT with or without 
chemoprophylaxis. In some areas, such as Cape Verde, Mayotte, Sao Tome and Principe 
there was a spectacular drop in parasite rates and malaria incidence, following the use of 
DDT (WHO, 1983). Malaria was successfully eradicated in large areas of South 
America, Southern Europe, the former Soviet Union and some territories of Asia and 
South America. The disease persisted in Latin America, most Asian countries and 
Africa. In the dry savanna of West Africa complete failure to control malaria 
transmission was encountered owing to a complex of technical, operational, 
administrative, logistic and financial factors (Haworth, 1981). In addition, population 
movements represented a constant threat of re-infection to the areas freed of malaria 
transmission (Prothero, 1961).
In 1968, DDT resistance in An. gambiae s.l. was reported in areas in West Africa where 
DDT was used in house spraying (Ivorra Cano and Bakri, unpublished report to WHO, 
1975). The uses of other pesticides in agriculture coupled with the extensive use of DDT 
in cotton production areas were believed to have contributed to the selection pressure for 
resistance (Zahar, 1984). A report of resistance to other classes of insecticides by 
mosquito vectors has further compromised the use of insecticide spraying in malaria 
vector control. The evolution of resistance to most insecticides by the vector prompted 
the need to develop new tools for vector control.
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==5.2.1 Current vector control interventions
Significant progress has been made since the 1980’s in research and development of new 
tools for use in malaria control activities. The use of insecticide treated bed nets (TTNs) 
and curtains with pyrethroids seems to be the most promising available method of 
controlling malaria in endemic tropical countries. Studies carried out in Senegal (Alonso 
et al., 1991) and China (Cheng et al., 1995) demonstrated the efficiency of ITNS for 
reducing infant mortality. These findings have been confirmed by subsequent large-scale 
multicenter studies in 6 countries across Africa including Ghana (Nevill et al., 1996; 
Binka et al., 1996; Lengeler et al., 1996).
However, there have been reports of the emergence of pyrethroid resistance in the 
Anopheles vectors (Curtis et al., 1998). Some cases of such resistance are known, 
notably in An. gambiae in West Africa where the kdr type of resistance has been selected 
probably due to the use of pyrethroids in cotton production (Zahar, 1984). Moreover, 
Doannio (1999) reports that despite the known repulsion effect of permethrin treated 
bednets on mosquitoes, the use of ITNs had no signficant impact on transmission. On the 
other hand, other studies have shown that the use of ITNS is effective in reducing 
morbidity and mortality due to malaria (Lengeler, 2000).
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2.7 The Systematics of Anopheles gambiae complex
Anopheles gambiae was thought to be one species until 1944 when studies by Ribbands 
and Miurhead Thomson (1944; 1945) in West Africa and Muirhead Thomson (1951) in 
East Africa provided the initial evidence for the specific distinctive nature of salt water 
breeding An. gambiae. The taxonomic status was still not understood but differences in 
behaviour were recognized. Cross mating studies by Muirhead Thomson (1951) later 
provided strong evidence of real differences by demonstrating hybrid male sterility 
between the two forms from East and West Africa.
In separate studies, hybrid male sterility between two fresh water-breeding populations 
was reported (Davidson, 1962). However, this was not considered to be evidence for 
interspecific differences between the two. Thus, as regards the fresh water gambiae, the 
familiar concept continued to prevail of a single opportunistic species adapted to a 
variety of environments. By 1964, three species had been recognized with Paterson 
(1964) presenting evidence that the three freshwater breeding species did not mate in 
nature. Holsten, (1952) had earlier suggested that the species consisted of 2 races. 
However on the basis of studies carried out by Davidson and Jackson (1962), Paterson
(1963) concluded that these authors were dealing with sibling species. Paterson et al.,
(1964) later provided direct evidence for the specific distinctiveness of three freshwater 
forms which he designated A  B and C that did not mate in nature. Later, another species 
breeding in heavily mineralized water in Uganda, originally documented by Haddow et
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al., (1947) was designated species D (Davidson and Hunt, 1973). This brought iheAn. 
gambiae complex to a total of six species.
Mattingly (1977) proposed the formal names gambiae Giles, arabiensis Patton and 
quadriannulatus Theobald for species A, B, and C respectively, and species D was 
named bwambae by White (1985). The two recognized salt-water breeding species are 
melas Theobald in West Africa and merus Donitz in East Africa (Paterson et al., 1964, 
Mahon etal., 1976). Recently, Huntef al. (1998) reported a new member of the conjff&V 
designated quadriannulatus species B from Ethiopia.
While the existence of the six formally named species of the An. gambiae complex is 
well established, there is the indication that further subdivision within species of the 
complex may exist. The evidence for cryptic taxa within An. gambiae s.s is the 
observation that the micromorphology of the second chromosome for different 
populations differ by inversions. These chromosomal forms, which are far from Hardy 
Weinberg equilibrium at certain times of the year in West Africa also carry different 
strain specific combinations of inversions and differ in vectorial capacity. This has been 
well studied in Mali from where three forms “Bamako”, “Mopti” and “Savanna” have 
been described (Toure et al., 1983). These forms appear to be fully or partially isolated 
reproductively in nature. In sympatric areas, hybrids between the “Savanna” and the two 
other forms have been observed at frequencies lower than expected. Also, no individuals 
carrying heterozygous complements of the “Mopti” and “Bamako” inversions are seen in 
nature, even though the two produce viable progeny under laboratory conditions
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(Coluzzi et al., 1985). Two other forms of ‘gambiae ’ namely “Forest” and “Bissau” have 
also been proposed (Biyan et ah, 1982).
2.8 Distribution of Anopheles gambiae s.L
Members of the An. gambiae complex have a wide geographical distribution and have 
been reported from most African countries and adjacent islands including Madagascar, 
as well as Saudi Arabia and Yemen (Coetzee et al., 2000). An. gambiae and An. 
arabiensis have the widest distributions. The two species show dissimilar distributional 
limits and asynchronous seasonal prevalence, probably due to divergent responses to 
climatic factors. An. gambiae predominates in forest and humid savanna zones whereas 
An. arabiensis is more successful in arid savannas and steppes, including those o f the 
south-western part of the Arabian Peninsula (Coetzee et al., 2000). Shifts in seasonal 
prevalence are observed showing an increase in the relative frequency of arabiensis 
during the dry season (Coluzzi et al., 1979).
The distribution of An. arabiensis is higher in the low rainfall zones, which are usually 
the drier savanna areas (Gillies and Coetzee 1987; Hunt et al., 1998). In areas where 
rainfall is less than 1000mm, An. arabiensis is recorded more often than An. gambiae 
and the reverse is true where rainfall is greater than 1000mm. An. quadriannulatus 
occurs sympatrically with An. arabiensis and less frequently with An. gambiae s.s. It 
appears to have a relict (narrow) distribution occurring in Southeast Africa, Ethiopia and
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Zanzibar (Hunt et a l, 1998). In Ethiopia, it occurs at localities with mean annual rainfall 
figures above 1000mm whereas in Southern Africa, the same species occurs in localities 
where annual rainfall is around 700mm (Coetzee et al., 2000).
Anopheles melas and An. merus are mainly salt and brackish water breeders on the West 
and East African Coast respectively, although An. melas has been found breeding in 
fresh water streams in the Gambia (Chineiy, 1984). Evidence points to a short dispersal 
range from preferred breeding sites by An. melas and the adults are usually not found at 
distances more than 3 km from the saline environment (Bryan, 1987). An. bwambae is 
restricted to the Semliki forest in the Rift valley near the Zaire border where it breeds in 
geothermal mineral springs (White, 1985).
The larval stages of the “Mopti” form occur mainly in man-made habitats, even during 
the dry season, while the “Savanna” and “Bamako” forms have a tendency to breed in 
more natural sites, exploiting rain dependent pools for larval development (Coluzzi et 
al., 1985). This affects their spatial and seasonal distribution, since the “Mopti” form 
will breed throughout the year and can therefore displace the other forms in irrigated 
areas, leading to changes in the patterns of malaria entomology (Toure et al., 1994). 
“Mopti” is the only form o f An. gambiae found during the dry season and could be more 
adapted than An. arabiensis, which could be competitively displaced (Toure et al, 
1998). “Mopti” is known from Mali, Guinea, Ivory Coast and Burkina Faso. Of the two 
other forms of "gambiae' namely “Forest” and “Bissau” that have been proposed (Bryan 
et al., 1982), “Forest” refers to forest breeding gambiae, occurring in rain forest areas
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and in humid or derived savannas. “Bissau” refers to a form ofA n. gambiae s.s recorded 
in the Gambia and Senegal where it is associated with the rice fields along the banks of 
river Gambia (Bryan et al., 1982). Unlike the six accepted species of the complex, these 
forms appear to display no post-mating reproductive isolation, with hybrids between 
forms actually having higher fecundity under laboratory conditions (Di Deco et al, 
1980).
The complexity surrounding the An. gambiae complex and presence of incipient species 
within An. gambiae s.s presents a need for accurate characterization of these strains, an 
exercise which is of utmost importance to malaria epidemiology studies.
2.9 Methods used in the identification of Anopheles gambiae sibling 
species
Although it has long been known that only some members of anopheline species 
complexes are important vectors of malaria, these sibling species have proven difficult, 
if not impossible to distinguish by traditional taxonomic measures. The similarity in the 
morphology of members of the Anopheles gambiae complex was a problem in their 
identification and characterization. The first indications of the presence of more than one 
species in An. gambiae came from crossmating experiments through which species A, B 
and C of the complex were recognized because of their mating incompatibility with 
known species (Davidson, 1964; Crampton, 1994). Since then, more techniques have 
been developed and applied in the differentiation of the cryptic species. These include:
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the detection of differential banding patterns of the polytene chromosomes from the 
ovarian nurse cells of the semi gravid adult females (Coluzzi and Sabatinni, 1967), 
analysis of iso-enzyme frequencies (Mahon et al., 1976; Miles 1978), analysis of 
cuticular hydrocarbons (Carlson and Service, 1979; Hamilton and Service, 1983) and the 
exploitation of species specific differences in the genome by the analysis of DNA 
sequences for distinguishing adults ofA nopheles gambiae complex (Gale and Crampton, 
1987; Collins etal., 1987; Besansky, 1999).
2.9.1 Morphological identification
The available morphological diagnostic characters although not reliable are of definite 
taxonomic value for the distinction of the saltwater and the fresh water sibling species of 
the Anopheles gambiae complex. The number of sensilla coeloconica, the value of the 
palpal index and the shape of the eggs can separate these two groups (Service, 1983). 
However, none of these characters appears to be completely discriminant as the 
morphological characters of adults are variable and overlap in many instances (Coluzzi, 
1964). No morphological features have been found as yet that can be reliably used for 
the distinction of freshwater species; An. gambiae and An. arabiensis; or between these 
species and An. quadriannulatus. The morphological divergences observed among the 
three fresh water species appears to be much less consistent and the morphological 
picture portrayed seems characterized by a wide intraspecific variability, presumably 
implying both polymorphism and polytipicism, which makes the evaluation of most of 
the taxonomic reliability of most of the characters studied problematic (Davidson, 1964).
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2.9.2 Identification of Anopheles gambiae based on reproductive 
incompatibility
Hybridization tests have been exploited to reveal various degrees of genetic 
incompatibility, typically expressed as hybrid male sterility, which was shown to exist 
between all the six taxa (Davidson, 1964). In experiments by Paterson, (1964) all crosses 
produced sterile male hybrids and the females from these hybrids were fertile but in 
certain crosses, few or no females were produced, meaning that the reproductive 
isolation is almost complete. However, there are practical limitations to this diagnostic 
approach, which depends on time-consuming laboratory activity, hence inappropriate for 
large-scale routine identification. Moreover, while species distinctness is generally 
indicated by hybrid sterility lack of it does not necessarily indicate or prove 
conspecificity.
2.9.3 Cuticular Hydrocarbon analysis
This method, fust described by Carlson and Service (1979) is based on the quantities of 
cuticular hydrocarbons of insects and has also been applied in the discrimination of 
Simulium blackflies (Carlson and Walsh, 1981) and in the Phlebotomine sand flies (Ryan 
et a l, 1986). Anyanwu et al. (2000) used this method in an attempt to separate the larval 
stages of members of the Anopheles gambiae complex. Overlaps in the hydrocarbon
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profiles were observed even after the use of discrminant analysis. This is not surprising 
because discrimination between the strains of An. gambiae is based on the quantities of 
various hydrocarbons and on the relative concentrations of the component chemicals, 
rather than mere presence or absence. However, Hamilton and Service (1983) observed 
that, although the fourth instars of An. gambiae and An. arabiensis had similar cuticular 
hydrocarbons, there were still differences in the relative level of some of these chemicals 
Although the biological implications of variations in cuticular hydrocarbons between 
very closely related species have not been fully studied, Phillips and Milligan, (1986) 
and Phillips et al. (1987) suggest that within such relationships species specific 
differences do exist which tend to highlight the effect of geographical variation and 
possible incipient speciation mechanisms. These chemicals, which serve mainly to 
prevent dessication and also assist in chemical communication, may additionally play a 
role in mate selection/recognition and population divergence. Hydrocarbon differences 
among sympatric populations of An. gambiae s.s. might also reflect a semiological 
function of the compounds enabling the insect to recognize potential mates in those 
locations where sibling species coexist (Phillips et al., 1987). In some insects, part of the 
mate recognition mechanism has been linked with the detection of specific hydrocarbon 
compounds and other components of the cuticular lipid layer e.g. fatty acids, alcohols, 
sterols, aldehydes, etc (Jallon, 1984; Pescke, 1987).
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2.9.4 Molecular methods for species identification and population 
studies
During the past twenty years, the use of molecular data in systematics has become veiy 
popular mainly because of the development of new biochemical tools such as gene 
cloning (Sambrook et al., 1989), DNA sequencing (Southern, 1975; Sanger etal., 1977), 
restriction enzyme analysis methods (McDonnell et al., 1977, Southern, 1979) and PCR 
(Mullis and Faloona, 1987). DNA analysis is highly suitable for population studies 
because it is the most direct analysis of genetic material possible and provides an easy to 
use method (Post et al., 1992). It is applicable to all developmental stages and both sexes 
and is quite sensitive, requiring only minute amount of material that is easily preserved.
Molecular identification has the advantage of being applicable to adult females 
independently of their gonotropic state, as well as to adult males and to pupae. A rDNA 
based method for the identification of 5 members of the An. gambiae complex exists 
(Scott et al., 1993). The rDNA-PCR method and other methods such as RAPD-PCR 
(Williams et al, 1990, Favia et al, 1994), RFLP (Beebe and Saul, 1995) and 
microsatellite DNA analysis (Lanzaro et al., 1995) have been applied to study An. 
gambiae s.l populations. All these methods are PCR based.
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2.9.4.1 Principles of PCR technology
The PCR is an in vitro method developed for the enzymatic synthesis of specific DNA 
sequences, using two oligonucleotide primers that hybridize to opposite strands and 
flank the region of interest in the target DNA (Saiki et al., 1985). The reaction utilizes 
the DNA polymerase enzyme, and occurs in a repetitive series of cycles resulting in the 
exponential accumulation of large amounts of the target DNA This process allows the 
enzymatic amplification of minute quantities of specific DNA sequences. The first 
experiments with PCR used the klenow fragment of Escherichia coli DNA polymerase 
(Saiki et al., 1985; Mullis and Faloona, 1987). However, the high temperatures used to 
denature the template DNA diminished the activity of the klenow fragment Therefore, 
more enzyme had to be added at the start of each cycle. This was both labour intensive 
and expensive. The technique was revolutionized by the discovery of a heat stable 
polymerase isolated from the thermophilic bacteria Thermus acquaticus and therefore 
called Taq polymerase (Saiki et al., 1985). The feet that Taq polymerase enzyme is 
added to the reaction once, at the beginning of the reaction has transformed the PCR into 
a simple and robust reaction, which is now automated by programmable thermal cycling 
devices. The reaction components (DNA template, oligonucleotide primers, Taq 
polymerase, deoxynucleotides and reaction buffer which contains salts) are all included 
in a reaction mixture and the amplification reaction carried out by simply cycling the 
temperature within the reaction tube (Saiki et al., 1988). The PCR reaction involves 
three steps of repeated cycles of heat denaturation of the DNA primer annealing to their
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complementary sequence at a lower temperature and extension of the annealed 
Oligonucleotide by the DNA polymerase.
The yield and specificity of PCR is dependent on several factors, which include 
concentration of the polymerase enzyme, primers, DNA template and salts, time 
allocated to each reaction for annealing and extension, and the total number of cycles. 
Because the primer extension products synthesized in one cycle can serve as a template 
in the next, the number of target DNA copies approximately doubles at every cycle. 
Thus, 20 cycles of PCR yields about one million-fold (220) amplifications
During PCR each primer becomes incorporated into the DNA template. Oligonucleotide 
primers are designed to amplify DNA from specific sequences of organisms. This 
increases the specificity of the PCR The annealing temperature of the primers is 
generally set as high as possible to promote specificity. However, it is recommended that 
annealing temperatures should be set at 5° C below the melting temperature (Tm) of the 
primer. The Tm is calculated using the formula of Thein and Wallace (1986):
Tm =[4(G+C)+2(A+T)]
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2.9.5 PCR for the identification of Anopheles gambiae species complex
2.9.5.1 The ribosomal DNA method
The PCR is currently widely used in the identification of member species of the An. 
gambiae complex. Scott et a l  (1993) developed the only method currently used for most 
studies. The method, which is based on species-specific nucleotide sequences in the 
ribosomal DNA (rDNA) intergenic spacer regions may be used to identify both species 
and inteispecies of hybrids regardless of life stage using either extracted DNA or 
fragments of a specimen. The non-coding rDNA spacer sequences can be highly variable 
in length and sequence between closely related species. Concerted evolution acting on 
rDNA assays maintains sequence homogeneity within species as it drives differentiation 
between species, a pattern that explains the utility of rDNA for species diagnostic assays 
(Collins and Paskewitz, 1996). In using this method for the identification of mosquitoes, 
intact portions of a specimen as small as an egg or the segment of one leg may be placed 
directly into the PCR mixture for amplification and analysis (Scott et al., 1993). The 
method uses a cocktail of five 20-base oligonucleotides to identify^/:, gambiae, An. 
arabiensis, An. quadriannulatus, and either An. melas in Western Africa or An. merus in 
Eastern and Southern Africa.
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2.9.5.2 Restriction Fragment Length Polymorphism (RFLP)
Restriction fragment length polymorphism (RFLP) provides a means of discriminating 
between similar DNA samples at a species or strain level. Extracted DNA is digested 
using restriction endonucleases, which cleave at specific recognition sites. The resulting 
restriction fragments are separated according to size by electrophoresis on agarose gels 
revealing diagnostic polymorphisms with distinct DNA fragments. However, in most 
cases, the digestion products appear as a smear with fragments ranging upto 
approximately 25kb in size. To gain useful information on these gels, the DNA must 
first be transferred to a nylon or nitrocellulose membrane by Southern blotting 
(Sambrook et al., 1989) and selective fragments visualized by hybridization with a 
suitable labeled DNA probe (Mills, 1994). Favia et al. (1997) designed a polymerase 
chain reaction (PCR-RFLP) assay, which unambiguously separates the An. gambiae 
Mopti form from Savanna and Bamako. This method, based on the presence of a 
restriction site length polymorphism has been tested on previously karyotyped sympatric 
specimens from Mali and Burkina Faso and it is already applied to verify the distribution 
of other molecular markers, as for example the pyrethroid resistance gene (kdr) among 
these chromosomal forms (Chandre et al., 1999). It should however be noted that this 
technique has so far, not differentiated between any chromosomal forms (Favia and 
Louis, 1999).
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2.9.5.3 Random Amplification of Polymorphic DNA (RAPD)
A further approach extensively used to distiguinsh members of cryptic species has been 
the use of Random amplified polymorphic DNA (RAPD) markets (Williams et a l, 
1990) The analysis of random amplified polymorphic DNA (RAPDs) provides a novel 
and effective method for distinguishing Anopheles species and other organisms 
according to the banding patterns of their DNA as well as providing a new means of 
obtaining genetic markers (Hedrick, 1992). RAPD markers have been used to identify 
An. gambiae from An. arabiensis species. The technique is fast, technically easy, and 
requires little material. Most importantly, no previous nucleotide sequence information 
is needed for the construction of primers. Many markers can be readily identified for a 
variety of taxonomic levels and, in comparison with DNA sequencing, the effort and 
cost are modest so that many individuals can be assayed. However, questions have been 
raised concerning the validity of the method and these may both hinder the general use 
of the technique for identification purposes and cause concern for routine identifications 
(Rollinson and Stothard, 1994). One potential problem associated with this technique is 
that RAPD-PCR protocols based on short (10-12mer) primers are often difficult to 
reproduce in different laboratories with different equipment and reagents, or with DNA 
extracted and stored by different methods (Black, 1993). Since reproducibility is a 
fundamental element of the monitoring of vector distribution, RAPD markers do not 
represent the approach of choice.
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2.9.6. PCR for the studies of Anopheles gambiae populations
2.9.6.1 Microsatellite DNA analysis
Microsatellites are described as DNA fragments consisting of simple short sequences 
usually of 2 -6  nucleotides (nt), tandemly repeated in more or less uniform tracts upto 
approximately 102nt long (Tautz, 1993; Chambers and Me Avoy, 2000). Microsatellites 
are found in the genomes of just about every known organism and organelle. In most 
eukaryotic organisms, microsatellites are dispersed throughout the genome (Hamada et 
a l, 1982) and can occur as frequently as every lOkb (Tautz, 1989). Polymorphism in 
microsatellites leads to an increased probability of finding heterozygous individuals. 
Microsatellite loci are also frequently hypervariable, (i.e. they possess several alleles at 
relatively high frequency) improving their versatility and hence are ideal tools for 
molecular characterization of individuals and studies of intraspecific variation (Tautz, 
1993; Lanzaro et al, 1995).
They are usually polymorphic in length due to variation in the number of repeats within 
a given locus as a result of uneven cross over (as in mini satellite refer to Jeffery et al., 
1985), slippage of the DNA polymerase during replication or unequal sister chromatid 
exchange (Tautz, 1989). The high polymorphism of microsatellite loci results from high 
mutation rates, estimated to range fromlO ~2 -10' 5 locus/gamete/generation (Dallas, 1992; 
Edwards et a l, 1992; Weber and Wong, 1993) with most estimates being between the
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10' 3 and 10"4 Replication slippage events are considered to be the main process of 
producing insertion/ deletion mutations of one or infrequently several repeat units 
(Levinson and Gutman, 1987). Other evidence suggests that forces that shape allele 
composition include unbiased mutation rate (Garza et al., 1995) and/or selection acting 
on allele size (Epplen et al., 1993). However, if such forces strongly affect allele 
composition at these loci, inter-population differentiation will be underestimated and 
gene flow will be overestimated to an unknown extent (Lehmann et al., 1996).
Microsatellite loci are more common in some organisms than in others and screening 
may produce a few useful loci in some species, particularly insects (Cooper, 1995). 
Repeats such as (dG-dT)n have been found in all eukaryotic organisms, except yeast In 
An. gambiae, conservative estimates suggest the presence of about 10,000 (dG-dT)n 
microsatellites in the haploid genome. In Aedes aegypti, the search for simple sequence 
repeats has not been fruitful. For example, approximately 104 recombinant cosmids were 
screened with a (dG-dT)is probe, and microsatellites were detected at a frequency of 
approximately 1 in 300 colonies that were screened (S.E Brown, unpublished data). 
Since the average size was >30 kb, it appears that (dG-dT)„ microsatellites are relatively 
infrequent in theAe. aegypti genome.
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2.9.6.2 Applications of microsatellite analysis
Exploitation of microsatellite loci has provided biologists with a set of molecular tools 
with unsurpassed versatility. The key to this versatility lies in the high levels of 
variability, which are characteristically found at such loci, coupled with the speed and 
reliability with which this information can be accessed in the laboratory (Chambers and 
Me Avoy, 2000). Their applications range from estimation of the spatial relationships 
between chromosome segments to the elucidation of temporal relationships between 
origins of species and genera. Microsatellite loci have also been described as ideal 
markers for measuring population level phenomena such as population structure due to 
their high polymorphism, codominance, abundant presence throughout the genome and 
relative ease in scoring (Bowcock et al., 1994; Buchanan et al., 1994; Scribner et al., 
1994; Eustop et al, 1995; Lanzaro et al., 1995).
Other applications include use in the analysis of laboratory and agricultural organisms 
and human genome diversity projects to map single gene traits rapidly (Heame et al.,
1992), DNA profiling (Gill et al., 1994) and also in forensic work (Chambers et al, 
1997). Microsatellites are almost perfect tools for application in determining the pattern 
of relationships between individuals because they are capable of highly discriminating 
biparentally inherited co-dominant markers. In the analysis of An. gambiae populations, 
microsatellite DNA analysis has been used to generate a genetic map of the mosquito 
(Zheng et al., 1993) genetic differentiation of populations (Lanzaro et al., 1995) and in 
the study of population structure (Donnelly et al., 1999).
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2.9.7 Cytotaxonomy
2.9.7.1 Polytene chromosomes and their value in cytogenetic studies
The repeated replication of chromatids without the occurrence of any mitosis-like 
events (endoreplication) leads to the production of thousands of sister chromatids within 
the same nucleus. This results in the formation of extended axial parts and contracted 
chromomeric loops of different sizes. The outcome are polytene chromosomes, 
appearing as bundles of extended interphase chromatids, characterized by peculiar 
banding patterns and by an axial length approximately 1 0 0 -2 0 0  times the length of the 
same chromosomes during mitotic metaphase (Della Torre, 1997). Such chromosomes 
can be easily observed under a compound microscope and are useful for cytological 
studies.
The pattern of cross bands and allied features is what makes polytene chromosomes 
useful for cytological studies. Each chromosome bears a unique pattern of light and dark 
bands, puffs (sometimes called balbianni rings) and constrictions that are readable. This 
permits the characterization and identification of species or groups as the patterns of 
bands and puffs are characteristic for every group or species. Moreover, the course by 
which closely related species have evolved away from each other may be traced by 
seeing which chromosome mutations (i.e translocations, deletions, duplications, 
inversions) are shared between species and which of these changes are distinctive. 
Inversions, in particular have proved to be important in the study of chromosomal
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relationships within sibling species complex for example in the Simulium damnosum 
complex (Dunbar 1969; Dunbar and Vajime 1972; Vajime and Dunbar, 1975; Boakye,
1993), the Anopheles maculipennis complex (Kitzmiller et a l, 1967) and the An. 
gambiae complex (Coluzzi and Sabatini 1967; 1969 and White, 1973).
In most mosquito genera, the polytene chromosomes are either not suitable for 
cytological manipulation or only partially replicated. Only those from Anophelines give 
excellent preparations, although in some species it is invariably difficult to obtain good 
polytene preparations. Polytene chromosomes of An. gambiae can be obtained from the 
salivary glands of fourth instar larvae and from the nurse cells of half gravid adult 
females (Coluzzi 1968). The species can be characterized by one pair of heteromorphic 
sex chromosomes (X and Y) and two pairs of autosomes that are numbered 2 and 3 
(Figure 4). However, since the heterochromatic region of the X chromosome and the 
whole Y chromosome are under-replicated, the typical anopheline polytene complement 
is substantially constituted by the euchromatic region of the X chromosome and by the 
two metacentric or submetacentric autosomes. This means a total configuration made of 
five polytenic arms (X, 2R, 2L, 3R, 3L) where X represents the sex chromosome, 2R, 2L 
and 3R, 3L the long and short arms of chromosome 2 and 3 respectiwely.
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MITOTIC
COMPLEMENT
3L
POLYTENE COMPLEMENT
Figure 4: The three chromosome pairs in the mitotic complement of Anopheles and the 
five elements seen in its polytene complement (a=autosomes; c=centromeres, L and R = 
short and long arms of chromosome 2 and 3) (From White et al., 1975).
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2.9.7.2 Chromosomal inversions
A chromosomal inversion is a form of mutation that constitutes an intra-chromosomal 
reversal of a block of genes. For example, a hypothetical gene arrangement ABCDEF 
may become ABEDCF after re-arrangement The inverted genotype persists only if it is 
intrinsically viable and if so, it may or may not be cross-fertile with the standard 
genotype (Coluzzi et al., 1985). A new inversion is an important evolutionary step ofthe 
sort that frequently accompanies dipteran speciation. When standard inverted gene 
arrangements interbreed freely in a population, a situation of balanced inversion 
polymorphism prevails in which a proportion of individuals are heterozygotes. To 
achieve maximum synapsis in somatic cells of the heterozygote, the homologous 
chromosomes form a loop configuration. Such loops are easily discerned in cytogenetic 
polytene slide preparations and have important homeostatic function in suppressing the 
process of crossing over, which normally gives rise to novel gene combinations during 
meiosis. Therefore, while inversion heterozyogtes exhibit heterotic vigour, which may 
enhance individual fitness, they reduce random genetic variability, which limits 
population adaptability.
It was the analysis of inversion polymorphism that led Coluzzi (1982) to propose that the 
key behavioral and adaptive differences between incipient and sibling species are 
integral rather than incidental to the speciation process. This idea was actually put 
forward in the context of chromosomal inversions bearing (and protecting from 
recombination), the determinants of these species-specific adaptations. Inversions also
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act as chromosomal mechanisms that preserve gene associations or supergenes arising in
j _ _____  ,  • « . _ ___i  ;  _ j . ___ j  i _  ____________1 . : ___11_.  — j  .  _  „  „  „i
tcr/iporarj’ i5uiuiC3 Suujwi/I. u> iiUdu anu  LIUJU ui gw ugiu|/m vuu^ c**ivj vii CwuJugiM*nj
marginal zones (Coluzzi, 1984; Petrarca and Bier, 1992; Coluzzi, 1992). Thus, 
inversions might express specific adaptive values for the environmental conditions in 
which they originated and alternative arrangements in a vector population might express 
ecotypic divergences that can be of interest in malaria transmission. Understanding the 
basis of chromosomal differences at the molecular level might give some insight into the 
speciation process and how human activity impinges on this process. In terms of 
malariology, some of these behavioural distinctions can make the difference between a 
good and a poor vector, and an understanding of their genetic basis may enable the 
manipulation of relevant behaviours to control disease transmission (Coluzzi et al, 
1985).
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2.9.1.5 Inversion polymorphisms in the An. gambiae species
Polytene chromosome studies on An. gambiae complex show that the rearrangements of 
the banding pattern are all based on paracentric inversions (Coluzzi, 1984). These occur 
in the homozygous state in different species (i.e the fixed inversions used for species 
identification) or constitute intraspecific chromosomal polymorphisms (floating 
inversions) for studying population variation. Inversion polymorphisms have been 
recorded in all the six species of the An. gambiae complex except An.merus and have 
been found to be particularly frequent in An. gambiae and An. arabiensis, the two taxa 
showing the widest distribution and the closest association with man-made breeding 
places (Favia and Louis, 1999).
Anopheles gambiae karyotypes are recorded according to the nomenclature of Coluzzi et 
al., 1978 in which standard arrangements for each karyotype are indicated by a + sign 
followed by a letter referring to the chromosome section and inverted arrangements are 
indicated by a letter alone, for example, 2L + a indicating the standard arrangement of 
inversion a on the short arm of chromosome 2 . 2 L+a/a indicates the heterokaryotype for 
the same inversion. Figure 5 shows the various karyotypes expected from eight 2R 
chromosome arrangements.
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Figure 5: Diagrammatic representation of karyotypes expected from the eight 2R 
Chromosome arrangements frequently observed in An. gambiae (From: Toure et al., 
1983)
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CHAPTER THREE
GENERAL MATERIALS AND METHODS 
3.1 The Study Sites
The An. gambiae larvae and pupae used for this study were collected from five breeding sites 
located within Greater Accra Region, in the southern part of Ghana. One study site, Dodowa is 
located in the coastal forest region, and the other four study sites, namely Adenta, Madina, 
Achimota and Legon are located in the coastal savanna region, which is characterized by dry 
climatic conditions receiving the least amounts of rainfall in Ghana.
The region has two rainfall peaks from April to June and from September to October, with the 
mean annual rainfall ranging between 740 and 890mm. The highest mean monthly temperature 
(about 30°C) occurs between March and April and the lowest (about 26°C) in August Relative 
humidity is high throughout the year, ranging between 65 and 75% in the afternoon.
The vegetation consists mainly of grass with isolated patches of scrub and sparse trees. The 
vegetation in Dodowa is described as coastal forest and is located between the coastal savanna 
and the beginning of the secondary forest areas. Very little of the original forest however 
remains and most of what is left is a mixture of few large trees with grass covering most of the 
area. The zone is very humid, with persistent rainfall (Dickson and Benneh, 1977).
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Small shallow pools of stagnant rainwater characterized the Dodowa site. In Adenta, Madina 
and East Legon, the samples were collected from open gutters exposed to sunlight, and 
containing stagnant water and these contained a mixture of both Anopheles and Culex larvae. 
The Achimota site was a stretched pool of water in the open and exposed to sunlight, flowing 
from a leaking water supply pipe. Photographs of the sample collection sites are shown in Plates 
1-5.
The global positioning system (GPS) was used to determine the geographical coordinates of the 
sites as being: Dodowa 05°85239N, 00°. 07028W; Adenta 05° 70339 N, 00° 15131W; Madina 
05° 67444N, 00° 15131W, East Legon 05° 64653 N, 00° 1446Wand Achimota 05°63436N, 00° 
23 993W. The selection of the study sites was based on distances apart. The furthest distance 
was between Dodowa and Achimota (25kms), followed by Dodowa and Adenta, 16kms, 
Achimota and East Legon 11 km. The shortest distance was 4Rms between Adenta and Madina 
(Fig 6 ). Sample collection was carried out in the dry season during September and October 
2000.
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C/3 
—I
c
o
<
tn
m
co
Figure 6  : Sample collection sites
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3.2 Field collection of mosquito larvae and pupae
The breeding sites ofA nopheles mosquitoes were identified by random sampling of small pools 
of stagnant water in the open and in gutters, exposed to sunlight. Small shallow pools of 
stagnant rainwater characterized the Dodowa and Madina sites. In Adenta and East Legon, the 
samples were collected from gutters containing stagnant water and these contained a mixture of 
both Anopheles and Culex larvae. The Achimota site was a stretched pool of water, flowing 
from a broken water supply pipe. From all these sites, Anopheles larvae were identified by their 
characteristic horizontal position on the surface of the water. These were carefully collected into 
small plastic containers by scooping gently to avoid injuring the larvae. The containers were 
loosely capped to avoid suffocation and immediately transported to the insectary for rearing to 
adults.
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Plate 1: Sample collection site at Dodowa. Shallow pool of stagnant rainwater 
exposed to sunlight.
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Plate 2: Sample collection site at Adenta. An open gutter exposed to sunlight, 
and containing stagnant water This site contained a mixture of both Anopheles 
and Culex larvae.
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Plate 3: Sample collection site at Madina. Shallow pool of stagnant water flowing from 
a residential washing area and exposed to sunlight.
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Plate 4: Sample collection site at East Leeon.
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Plate 5: Sample collection site at Achimota. A narrow stretched pool of water 
flowing from a leaking water supply pipe and exposed to sunlight
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3.3 Laboratory rearing of mosquitoes
The larvae and pupae were transferred into plastic labelled rearing trays containing pond 
water (Plate 6 ). In cases of where larvae of Culex species were present they were 
identified by their angular position on the water surface and were removed. The level of 
water in the rearing trays was maintained at a depth of approximately 2cm. Rearing 
conditions were maintained at air conditions of27-30°C and 76±2% relative humidity. A 
12h:12h light and dark cycle was maintained. The larvae were fed on finely ground 
nutrafm goldfish food (Rolf Hagen, USA). Each morning the larvae that hatched to 
pupae were collected into shallow plastic cups using rubber pippettes, and then placed in 
appropriately labelled cages for adult emergence (Plate 7). The adults were fed on 10% 
sugar solution within 24 hours of emergence.
After 3 days, the males and females were assumed to have mated. Most of the males 
dropped dead at this stage and were picked out of the cage. Any live ones, which were 
probably yet to mate were left in the cage, presuming that some of the females had not 
mated. The mated females were then starved for 12-14  hours prior to blood feeding by 
removing the sucrose solution from each of the cages. This was followed by aspiration 
of the females into small paper cups (plate 7) and blood fed, then returned into the cage 
for ovarian development to the half gravid stage. The development to the half gravid 
stage took about 12-15 hours. The rate of development differed for each individual 
female hence the development was regularly monitored after 12  hours by aspirating into 
tubes and checking each individual female abdomen for the half gravid appearance. The 
half-gravid females were killed by brief refrigeration at 4° C.
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Plate 6 : Larval rearing trays used in the laboratory rearing of Anopheles larvae and 
pupae
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Paper cups
Adult cage
Sugar 
— solution 
soaked in 
cotton 
wool
Plate 7: Adult cages for holding emerging adult mosquitoes
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3.4 Morphological Identification of Female Anopheles Mosquitoes
The identification of adult female An. gambiae was made using morphological 
characters outlined in 3.4.1 below.
3.4.1 Outline of criteria for the morphological features used for the 
identification of adult Anopheles gambiae s.l.
Palps: Smooth, with 3 pale bands: The apical band broad and enveloping the whole of 
the 5th and apex of the 4th segment, median band narrow, less than half width of apical, 
involving apex of third and extreme base of 4th segments; basal band as median, usually 
confined to apex of 2nd segment.
Thorax: Ground colour variable, usually light brown or grayish; mediannally clothed 
with cream or yellow scales, pointed and moderately broad; laterally scales much 
broader and often rounded apically; anterior pronotal lobes with scales.
Pharynx: With 12-20 pairs of rods and cones, latter with deep roots and long flattish 
pointed filaments, which are somewhat fimbriated, crest not bifid in posterior view; rods 
with short filaments, post pharyngeal ridges with very long hair like processes.
Legs: Femora, tibiae and first tarsal segment speckled to a variable degree, sometimes 
with speckles fused to form short lines, rarely in the form of complete rings; tibiae rather 
narrowly pale apically; on fore and mid legs apices of first and fourth tarsi and apices
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and bases of tarsi 2 and 3 narrowly but distinctly pale, 5**1 all dark; hind tarsi similar but 
basal pale bands on 2 and 3 absent or reduced 5th tarsus usually all dark.
Wing length. 2.8-4.4 mm
Wings: Pale markings, yellowish or cream. Wing field largely pale and with pale costal 
spots fairly long. The illustration is that of an average type of wing but there is much 
variation in the relative sizes of the pale and dark markings. Some of this variation may 
be indicative of inter-specific differences. Fringe spots present opposite veins 3-6; in 
addition a pale spot present between 6  and 5 .2 and between 6  and the base of the wing.
Abdomen: Light brown, mainly clothed with hairs but 8 th tergite usually with some 
scales, which may extend onto 7th tergite. Cerci scaly. Stemites conspicuously marked 
with symmetrical grey patches.
3.5 Sample Preservation
Each identified An. gambiae was divided into two parts by separating the abdomen with 
the half gravid ovaries from the head and thorax. The ovaries were immediately fixed in 
freshly prepared modified Camoys solution (three parts absolute ethanol and one part of 
glacial acetic acid) for cytotaxonomy (Hunt, 1973) and stored at -20°C. The head and 
thorax were preserved dry on silica gel until ready to use for DNA extraction.
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3.6 Cytotaxonomy
3.6.1 Preparation of chromosomes for cytological studies
The various stains and solutions used for the cytological studies were prepared as 
outlined in Appendix I.
Each of the fixed abdomens was placed onto a clean (dust-free and grease-free) 
microscope slide and pierced laterally with a dissection needle. The first three 
abdominal segments were pulled off directly into a drop of 50% propionic acid. The 
ovaries were then extracted simultaneously and isolated and then transferred into a drop 
of 50% propionic acid on a fresh microscope slide. The ovaries were left in the solution 
until they had swollen to approximately twice their original size and then divided 
laterally to separate the follicles. A drop of 2% Aceto-lactic orcein stain was added to 
propionic acid with the follicles and stirred with the needles to ensure homogenous 
staining. The tissue was left in the stain for 1 -5 minutes after which excess fluid was 
drawn off using absorbent paper. This was followed by washing of the tissue with 
several changes of propionic acid to give a pale pink colour. A grease free and dust free 
cover slip was placed on the tissues and carefully squashed by gently tapping on the 
cover slip with the back of a pen. The preparation was then examined under a compound 
microscope.
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3.6.2 Species identification by karyotyping
Using a phase-contrast microscope (Olympus B201, Japan), inversions on the specific 
band sequences on the whole chromosome complement were examined. The inversions 
on X chromosome were used for the identification of An. gambiae species (Coluzzi et 
al., 1977). Full karyotyping was carried out and inversions were scored according to the 
nomenclature of Coluzzi et al. (1988). Standard arrangements for each karyotype were 
indicated by a “+” sign followed by a letter referring to the chromosome section. A letter 
alone indicated inverted arrangements. For example, 2L + “a”, indicates the standard 
arrangement of inversion “a” on the short arm of chromosome 2. Photographs of 
chromosome prepartations were taken using a camera attachment to the microscope with 
green filter and Ilford Pan F film (Ilford Imaging, UK).
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3.7 Molecular Studies
The various buffers and solutions used were prepared as outlined in Appendix II.
3.7.1 The extraction of genomic DNA
DNA extraction was done using the method by Collins et al. (1987) with slight modifications. 
Briefly, each specimen (head and thorax) was placed in a 1.5 ml eppendorf tube and homogenized 
with a sterile glass rod in 100(il Bender buffer (0.1 M NaCl, 0.2M Sucrose, 0.1M Tris—HC1, 
0.05M EDTA pH 8.0 and 0.5% SDS). This was then incubated at 65°C for 30 minutes followed 
by the addition of 15 fxl of pre-chilled 8M KAc and mixed well by tapping the tube, then left on 
ice for 30 minutes. It was then centrifuged for 5 minutes at 14000 rpm and the supernatant 
transferred into a fresh tube. To the supernatant was added two volumes of absolute ethanol, 
mixed well by tube inversion and then incubated at -40°C for 2 hours. This was followed by 
centrifugation at 14000 rpm to pellet the DNA The supernatant was discarded and the pellet was 
left to dry by evaporation after which it was re-dissolved in 25 |jL of sddH20, and left on ice for 1 
hour. The DNA was stored at -20°C until ready for use.
3.7.2 PCR Identification of species of the Anopheles gambiae complex
Five sets of primers designed from the DNA sequences of the intergenic spacer region of 
An. gambiae complex ribosomal DNA (rDNA) were used in PCR for the identification 
(Scott et al., 1993). The sequence details of these primers abbreviated UN, GA, ME, AR 
and QD and the expected sizes of the PCR products are given in Table 1. The UN primer 
anneals to the same position on the rDNA sequences of all five species, GA anneals
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specifically to An. gambiae s.s., ME anneals to both An. merus and melas, AR to An. 
arabiensis and QD to An. quadriannulatus.
The PCR reaction mix of 25 |il contained IX PCR buffer supplied by the manufacturer 
(Sigma, USA), 200|iM of each of the 4 oligonucleotide triphosphates (dNTPs), 10(iM of 
each oligonucleotide primers and 0.125 units of Taq Polymerase enzyme (Sigma, USA). 
0.5 (j.1 of the genomic DNA was used as template for the amplification reaction. Sterile 
double distilled water was used to make up the volume to 25(il. The reaction mix was spun 
down briefly at 14,000rpm and overlaid with mineral oil to avoid evaporation and 
refluxing during thermo cycling. The amplification was carried out using a PTC 100 
thermal cycler, MJ Research Inc., USA
The cycling parameters for the reaotion were as follows: 93° C for 3min (initial 
denaturation), followed by 35 cycles of 93°C for 30s, 50°C for 30s, 72°C for 60s 
(annealing) and a single extension cycle of 93° C for 30s, 50° C for 30 s and 72°C for 10 
min. For each reaction a positive control with PCR products of Anopheles of the same 
primer set and a negative control that contained no DNA template were included.
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Table 1: DNA sequence details of the synthetic oligonucleotide primers used for the 
identification An. gambiae s.l. species and their melting temperatures.
Name of Sequences (5’ —3’) Melting Expected 
Primer Temperature Amplified 
Tm (°C) DNA 
size (bp)
UN GTG TGC CCC TTC CTC GAT GT 58.3 468
GA CTG GTT TGG TCG GCA CGT TT 59.3 390
ME TGA CCA ACC CAC TCC CTT GA 57.2 464
AR AAG TGT CCT TCT CCA TCC TA 47.4 315
QD CAG ACC AAG ATG GTT AGT AT 42.7 153
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3.7.3 Microsatellite DNA analysis
Three sets of oligonucleotide microsatellite primers; AGXH7, AG2H26 and AG2H46 (Lanzaro et 
al., 1995) were used in PCR to amplify microsatellite DNA sequences in the An. gambiae s.s. All 
the three primer sets each consist of a foward primer and a reverse primer. The primer set 
AGXH7 is designed to amplify a microsatellite locus on the X chromosome, whereas AG2H26 
and AG2H46 are designed to amplify microsatellite loci on chromosome 2. The sequence details 
of these primers and the expected sizes of the PCR products are given in Table 2.
The amplification of the microsatellite sequences was carried out using the method ofZheng et al. 
(1995). The reaction mix of 20 microlitres (|il) contained IX PCR buffer supplied by the 
manufacturer (Sigma, USA), 100nM of each of the 4 oligonucleotide triphosphates (dNTPs) 
(Pharmacia, Sweden), IOjiM of each oligonucleotide primers and 0.125 units of Taq Polymerase 
enzyme (Sigma, USA). 0.5|il of the genomic DNA was used as template for the amplification 
reaction. Three primer sets AGXH7, AG2H26 and AG2H46 were used respectively in each 
reaction. Sterile double distilled water was used to make up the volume to 20|il. The reaction mix 
was spun down briefly at 14000 rpm and overlaid with 1 0 ja 1 mineral oil to avoid evaporation and 
refluxing during thermocycling.
The cycling parameters for the reaction were as follows: 95°C for 5 min (initial denaturation), 
followed by 35 cycles of 95°C for 30s denaturation, 55°C for 30s annealing, 72°C for 2 min 
(extension) and a final cycle of 95° C for 30° s, 55° C for 30 s and 72°C for 5 mins. The annealing 
temperature used for each reaction depended on the (Tm) of each primer set. For each reaction a
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negative control that contained no DNA template were included. PCR amplification was carried 
out using a PTC 100 thermal cycler (M.J. Research Inc., USA).
3.7.4 Analysis of PCR products
3.7.4.1 Agarose gel electrophoresis
Following the PCR, the products were electrophoresed on a 2% agarose gel stained with 
0.5|ig/ml EtBr to detect the presence of amplified DNA fragments. Eight microlitres of 
each sample was added to ljil of orange G (5X) gel loading dye for the electrophoresis.
The gel was prepared and electrophoresed in IX TAE buffer using a mini gel system 
(BIORAD USA) at 100 volts for one hour and the gel photographed over a UV 
transilluminator (UPC, USA) at short wavelength using a Polaroid camera and film type 
667 (Polaroid, USA).
3.7.4.2. Polyacrylamide gel electrophoresis
The microsatellite DNA strands were electrophoresed in a 7 % polyacrylamide gel. The 
ingredients used in the preparation of the polyacrylamide gel are shown in the Appendix 
ILFifteen microlitres of the PCR product from an individual mosquito was mixed with 
10|iL bromophenol blue dye and loaded into each well. Prior to loading the samples onto 
the gel, the wells were flushed with sddFfeO to remove any urea from the wells. The gel 
was run at 13 mA for 7 hours, after which the glass plates were separated and the gel 
carefully transferred into a plastic tray containing 5% EtBr for 5-10minutes. The gel was 
visualized and photographed as described in section 3.7.4.1 above.
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Table 2: DNA Sequence of the synthetic oligonucleotide primers used for the 
amplification of An. gambiae s. s. Each primer is given with its annealing temperature 
and the size of the expected PCR product for An. gambiae s.s. (F) and (R) denote 
forward and reverse primers respectively.
Name of Sequences 5’ - 3 ’ Annealing Expected 
Primer Temperature Amplified 
(°C) DNA size(bp)
AGXH7 CACGATGGTTTTCGGTGTGG (F) 61 99
ATTTGAGCTCTCCCGGGTG (R)
AG2H46 CGCCCATAGACAACGAAAGG (F) 63 139
TTGTACAGCTGCAGAACGAGC (R)
AG2H26 CCGGCAACACACAAACAATCGG (F) 6 6 154
GGTTCCCTGTTACTTCCTGCC(R)
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3.8 Data Analysis
3.8.1 Karyotype data
The frequencies of the different karyotypes observed were tested for the Hardy - 
Weinberg equilibrium using the Chi square test for observed and expected frequencies.
3.8.2 Microsatellite data
Each primer set was expected to give specific allele (band) sizes. The banding profile of 
the alleles of each sample as visualized on the gel was scored forthe absence or presence 
of alleles (bands), depending on the DNA fragment sizes. The different sizes were 
denoted A, B, C, D, E, F and G. Distinct single bands were scored as homozygotes (e.g. 
AA) whereas bands which appeared either in pairs and in fours were scored as 
heterozygotes (e.g. AB). Missing bands were denoted by two dots (forthe details of 
results, see Appendix HI). The alphabetic diploid and co-dominant data was analyzed to 
determine the genetic variation among and within populations using population genetics 
analysis software, POPGENE, Version 1.31 for Microsoft Windows™ (Yeh etal., 1999). 
This was used to compute the relevant statistics for single locus in single populations 
and multiple populations; multilocus in single populations and multilocus in multiple 
populations. The output statistics included allele frequency, genotypic frequency, genetic 
distances, differentiation indices (Fst and Fi„) and gene flow (Nm).
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Using the algorithm described by Levene (1949) with a null hypothesis of random union 
of gametes and an alternative hypothesis of heterozygote deficiency, Chi square (x2) tests 
were performed for each locus to test for departure from the Hardy-Weinberg 
equilibrium at P<0.05. Population differentiation and gene flow were determined by 
computing Weir and Cockerham’s (1984) estimates of F statistics (F;sand Fst). The Fis is 
an estimate of heterozygote deficiency or excess. This was calculated to determine 
heterozygote deficiency (Positive F;svalue) or excess (Negative F;s value). FJf is an 
estimate of inter-population genetic differentiation and was considered to be low if the 
differentiation index (Fst) was below 0.05 and high when Fa was greater than 0.05 
(Wright, 1978). Positive Fjs values indicate heterozygote deficiency while negative 
values indicate heterozygote excess (Lehmann e ta l, 1997). The derived F statistics were 
used to estimate gene flow (Not), i.e. the number of migrants per population per 
generation using the formula N/m = (1- Fst)/4Fst (Slatkin and Barton, 1989). Gene flow 
was considered low when Nm ranges values are less than 3. Gene flow is high when Nm 
ranges from 3-30 and very large when Nm is greater than 30 (Donelly et a l, 1999). 
Estimates of Nei’s (1972) genetic distances were also calculated. Based on these 
distances, phylogenetic trees indicating the genetic relationship between the populations 
at each locus were constructed using Unweighted Pair Group Mean Average method 
(UPGMA)
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CHAPTER FOUR
RESULTS 
4.1 Laboratory Rearing of Anopheles gambiae
About 80% of the larvae and pupae that were reared in the insectary completed their 
growth and emerged as adults. Development from the larval to pupal stage lasted 3-5 
days depending on the larval stage from the field whereas adult emergence from pupae 
lasted 1-2 days. Fourth stage larvae pupated overnight (10-12 hours from time of 
collection). It was observed that the first batch of mosquitoes that emerged had more 
males than the subsequent batches, which had more females. The development of blood 
fed females to the half gravid stage was very successful and lasted 12-15 hours, 
depending on temperature. Morphological identification revealed that all the specimens 
that were studied 'were Anopheles gambiae s.l.
4.2 Karyotype analysis of Anopheles gambiae s.l.
Out of the 129 samples dissected, only 35 gave readable polytene chromosome 
preparations and all were An. gambiae s.s. as identified by the presence of the diagnostic 
homozygous inversion Xag on the X chromosome (Figure 7). No hybrids or 
heterozygous inversions on the whole polytene complement were found. No inversions 
were recorded on the chromosomal arm 2R; hence this karyotype was standard 2R+ in
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all the specimens (Figure 8 ). However, a homozygous floating inversion “ a ” on the 
chromosomal arm 2L was recorded (Figure 9), hence 2L was polymorphic with the 
alternative arrangements + a and a. The inversion 2La was absent in the 10 specimens 
from Dodowa and in the 10 Adenta samples. In the EastLegon samples, 8 out of 10 were 
standard for 2R and 2L (2R+, 2L+) whilst the remaining 2 samples possessed the 2R+, 
2La chromosomal arrangement. 4 out of 5 samples from Achimota had the chromosome 
2L inversion (2La), which was absent in only one sample from this site. Table 3 shows 
the percent frequencies of the 2La karyotypes observed at the different sites. A 
significant deviation from the Hardy-Weinberg equilibrium was observed only in the 
Achimota samples (P<0.05).
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> * «
‘ S ' - ?
* Z  <
* •  s i *  * ^  *
*  - -
~a
s S jrV * *  1 *  * * •
. Xag1.
Figure 7: Chromosome complement of Anopheles gambiae showing 
the X chromosome (labelled) and the breakpoints of the diagnostic 
inversion (Xag)
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Figure 8 : The right aim of chromosome 2 (2R) of An. gambiae s.s. showing the 
standard arrangement.
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Figure 9: The left arm of chromosome 2 (2L) showing the floating inversion 2La 
which is a 180° rotation on section a.
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4.3 PCR identification of Anopheles gambiae s.s.
The DNA extraction from the 129 An. gambiae specimens was successful in all cases 
resulting in single bands (Figurel 0). PCR amplifications revealed that all the specimens 
were An.gambiae s. s., confirmed by the size of the amplified DNA fragment which is 
390 bp (Figure 11). The numbers of mosquitoes identified per site were 30 for Dodowa, 
30 Adenta, 22 Madina, 22 East Legon and 25 Achimota.
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1 2  3 4 M
< -------  1 0 0 bp
Figure 10: Ethidium bromide stained 1% agarose gel electrophoresis of DNA extracted 
from Anopheles gambiae mosquitoes. Lanes 1-4 = genomic DNA; lane M = lOObp 
molecular weight marker.
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M 1 2 3 4 5
<----------  390bp
Figure 11: Ethidium bromide stained 2%  agarose gel electrophoresis of PCR products 
obtained from the amplification of Anopheles gambiae DNA for species identification. 
Lanes 1= Dodowa; lane 2 = Adenta; lane 3 = Madina; lane 4 = East Legon; lane 5 
Achimota M= 100 bp Molecular weight marker.
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4.4 Genetic structure of Anopheles gambiae populations
Microsatellite DNA analysis was done using the primer sets shown in Table 2. A total 
of 129 specimens were analysed. Out of these, 80 (62%) were successfully amplified on 
first attempt Of the remaining 49 (38%), only 10 (20 %) were successful on re­
amplification. The failure of some samples to show any bands even after re­
amplification was taken to be due to the possession of null alleles.
Following the convention of Excoffier et al. (1992), DNA bands that differed in sizes 
were taken to be distinct alleles. All the three microsatellite loci studied were 
polymorphic and a total of 30 alleles were recorded which were not uniformly 
distributed in all the populations. The distribution was as follows: 12 alleles at locus 
AGXH7 on the X chromosome and 9 alleles each at the two loci AG2H26 and AG2H46 
on chromosome 2. At locus AGXH7, 8 8  out of the overall total of 129 mosquitoes were 
heterozygous for the 99bp allele whilst at locus AG2H26, 74 were heterozygous for the 
98bp allele. At locus AG2H46 homozygotes were found to be dominant forming 76.7% 
of the 129 specimens examined.
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Figure 12: Polyacrylamide gel electrophoresis of PCR products obtained from the 
amplification of An. gambiae s.s. microsatellite DNA with primer AGXH7. Lanes 
land 3 = Homozygous single bands (AA), Lane 2,5 and 8  = heterozygous double 
band (AB), Lane 4 and 7 = Heterozygous set of four bands (AB), lanes 6,9 = No 
bands (..), lane M = 100 bp Molecular weight marker.
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Figure 13:Polyacrylamide gel electrophoresis of PCR products obtained from the 
amplification of An.gambiae s.s microsatellite DNA with primer AG2H26. Lane 
3 = Homozygous single band (BB), lane 6  = homozygous single band (DD), lanes 
1,2,8 ,9 = heterozygous double bands (BC), lanes 5 ,7 = heterozygous double bands 
(CD), lane 4 = heterozygous double bands (AC), lane 10 = No bands (..), lane M =
100 bp Molecular weight marker.
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1 2 3  4 M 5 6 7 8 9  10 M
<---------250bp
■4---------154bp
*--------- lOObp
Figure 14: Polyacrylamide gel electrophoresis of PCR products obtained from the 
amplification of An. gambiae s. s. microsatellite DNA with primer AG2H46. Lane 1 = 
Heterozygous four bands (DE), lanes 2,8,9 & 10 = heterozygous double bands (EF), 
lanes 3,4 & 5 = Homozygous single bands (DD, EE) lanes 6,7 = No bands (..) M = 100 
bp Molecular weight marker.
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4.4.1 Allele and genotype frequency distributions
Table 4 and Figures 15-17 show the distribution and frequencies of all the common 
alleles that were found. Eight alleles were the most common; however alleles 1 and 2 
were the most dominant and were present in all the five populations but at different 
frequencies. The mean allele frequency in the pooled populations was 7.33.
The genotype frequencies for all the loci deviated from the Hardy-Weinberg equilibrium 
(HWE) at the 0.05 level of significance when pooled population data were analysed 
(Table 5) When each of the three loci and five populations were tested individually the 
Dodowa, Madina and Achimota populations were in equilibrium at the AGXH7 locus 
(P>0.05), whereas Adenta and East Legon populations significantly deviated from it 
(P<0.05). At the AG2H26 locus only the Achimota population was in HWE (P>0.05), 
whilst Dodowa, Adenta, Madina and East Legon populations were not (P<0.05). At the 
AG2H46 locus, Adenta, East Legon and Achimota populations were in HWE (P>0.05) 
whereas Dodowa and Madina were not (P<0.05).
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Table 4: Allele frequencies of most common alleles at all the three loci.
Locus Sample No of Frequency of the most common
size alleles alleles
1 2 3 4 5 6 7 8
AGXH7 129 12 0.30 0.25 0.18 0.03 0.05 0.05 0.03 0.03
AG2H26 129 9 0.34 0.29 0.13 0.03 0.16 0.00 0.03 0.03
AG2H46 129 9 0.28 0.31 0.08 0.00 0.17 0.06 0.08 0.08
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Alleles
□ 1 
B 2
□ 3
0.8 □ 4 ■ 5
O □ 6 
©cS3 0.6 ”  7 
<CDT □ 8
0.2
0
Dodowa Adenta Madina East Legon Achimota
Populations
Figure 15: Allele frequency distribution at the AGXH7 locus
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Alleles
□□  1 2
□  3
□  4
□□  65 
□ 8
Dodowa Adenta Madina East Legon Achimota
Populations
Figure 16: Allele frequency distribution at lie AG2H26 locus
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4.4.2 Population differentiation Index (Fst)
The differentiation index i.e. F.,/ indicating the genetic relatedness of mosquitoes within 
each population was zero in all cases for all the loci. However, when pooled loci data 
were analysed for all the populations together, considerable differentiation between loci 
was observed (Table 6). The mean value over all loci was 0.0984. Estimates of F« 
were highest for the AGXH7 locus (Fst = 0.1282) followed by AG2H46 (F,t =  0.0993) 
and was lowest at locus AG2H26 (F,, = 0.0664). F,t was significant for all the loci, being 
greater than 0.05 (see section 3.8.2) but most significant at locus AGXH7 (F^ = 0.1282) 
indicating high differentiation at this locus compared to the other two.
Pairwise comparison of genetic differentiation between populations revealed that the 
highest Fst was between the furthest site Dodowa and the rest of the four sites (Fst 
=0.0565). This was followed by a differentiation index of 0.0511 between East Legon 
and Achimota, separated by a distance o f approximately 8 Km. The lower F« values 
were found between the closer populations being 0.0344 for Madina and Achimota (5km 
apart) and 0.0284 for Madina and East Legon (2km apart).
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4.4.3 Estimate of heterozygote deficiency and excess (Fj*)
The heterozygote deficiency indicated by a positive F« value was obtained in all 
populations except the Madina population, which had a negative value. The Fis values 
were 0.0204 at Dodowa, 0.1457 at Adenta, -0.0435 at Madina, followed by an increase 
to 0.4910 in East Legon and 0.5219 in Achimota (Figure 18).
However, when the data were analysed according to loci, the two loci on chromosome 2, 
Locus AGXH7 and AG2H46 indicated heterozygote excess values o f -0 .0 0 9 8  and -  
0.0654 respectively. Only locus AG2H26 showed a heterozygote deficiency o f 0.1427. 
The mean estimate of heterozygosity (Fjs) over all loci was an excess value o f -0 .065 5 
(Table 6).
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Table 6: Estimates o f differentiation (Fst), gene flow (Nm) and 
heterozygosity (FjS) at each Loeus
Locus Fst F|. Nm
AGXH7 0.1282 -0.1427 1.7004
AG2H26 0.0664 0.0098 3.5152
AG2H46 0.0993 -0.0654 2.2680
Mean 0.0984 -0.0655 2.2900
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Table 7:
Pairwise comparison of estimates differentiation F,( and gene flow (Nm), and genetic 
distances between populations based on pooled loci data.
Populations F* Nm Genetic distances
Dodowa vs Adenta 0.0565 4.1773 0.2999
Dodowa vs Madina 0.0565 4.1773 0.2999
Dodowa vs E ast Legon 0.0565 4.1773 0.2999
Dodowa vs Achimota 0.0565 4.1773 0.2999
Adenta vs Madina 0.0284 8.5672 0.1957
Adenta vs East Legon 0.0284 8.5672 0.1957
Adenta vs Achimota 0.0284 8.5672 0.1957
Madina vs East Legon 0.0344 7.0163 0.1461
Madina vs Achimota 0.0344 7.0163 0.1461
East Legon vs Achimota 0.0511 4.6433 0.1410
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Populations
□  Dodowa
□ Adenta
□ Madina
□  EastLegon 
■  Achimota
0 6
0 5  
0 4  
0 3  
0.2 
0 1
0 
- 0.1
Figure 18: Estimates of Fb values showing deficiency and excess 
of heterozygotes.
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4.4.4 Gene flow (Nm)
Fairly large estimates of gene flow (Not) were derived from F^. Details of results 
obtained are given in Table 7 and Figure 20 Accordingly, estimates o f gene flow (Not) 
were highest between the closest populations, Adenta and Madina (Not = 8.5672); 
followed by Madina and Achimota (Not =  7.0163), East Legon and Achimota Not = 
4.6433. The lowest estimate of gene flow was between the furthest populations, Dodowa 
and Achimota (Not =4.1773).
A plot o f the estimates of gene flow against distances between populations indicates the 
increase in level of gene flow between populations with a decrease in the physical 
distances between the populations (Figure 20). Linearization o f the plot which is an 
inverse function (Y=a/x + b) resulted in the determination of the two unknowns, a and b 
(a = 15, b = 3.3), where a = the point at which the linear function intersects with the 
straight line and b is the gradient o f the straight line. The equation o f the inverse function 
Y=15/x +3.3 was thus derived, where Y = gene flow and x = the distance between 
populations.
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Distance between populations (Km)
Figure 19: A plot o f gene flow between populations against distances between them.
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4.4.5 Phylogenetic relationship
To illustrate the phylogenetic relationship among the five An. gambiae populations and 
the three loci, phenograms were constructed for each chromosome separately and for 
the pooled ensemble using the Unweighted Pair Group Method with Arithmetic 
averaging o f Fst values, calculated with the POPGENE software. It was observed that 
the loci on chromosome 2  provided a phenogram that was most similar to the pooled 
loci and different from that of the locus on the X chromosome (Figure 20).
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---------- Dodowa
----------A:himota
------------------------------Madina
—  Merita
—  East Legon
I-----------------1
0.005
Locus AGXH7
---------------------------------------D odow a
____________  _________________________ E a s t  Legon
----------------------------------------------------------------------------------— ----------------- - Ad e n ta
----------------------------------------------------------------- Madina
----------------------------------------------------------------- Achim ota
I---------------1
0.01
Locus AG2H26
--------------------------------------  D od o w a
____________  _________________________ E a s tL e g a n
----------------------------------------------------------------------------------------- ----------------Ad e n ta
----------------------------------------------------------------- Madina
----------------------------------------------------------------- Achim ota
I---------------1
0 01
All Loci combined (AGXH7, AG2H26 & AG2H46).
Figure 20: Phenograms based on UPGMA to illustrate the phylogenetic relationships 
among the An. gambiae s.s chromosomal forms and populations.
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CHAPTER 5
DISCUSSION AND CONCLUSION
Anopheles gambiae is the principal vector of human malaria in sub-Saharan Africa. A 
better understanding o f  the species and its distribution is needed for the planning of 
effective malaria control strategies and the management of insecticide resistance. This 
study was set to characterize An. gambiae species from five sites at various distances up 
to 25 km within the coastal savanna and coastal forest zones of Ghana by karyotyping 
and microsatellite t)N A  analysis. The main aim was to establish the karyotypes in the 
zone and assess the levels of gene flow between the populations.
Anopheles gambiae s.l. was identified as the dominant Anopheles mosquito species. 
Further identification o f these species as An. gambiae s.s by both karyotyping and PCR 
revealed An. gambiae s.s. as the major sibling species of the An. gambiae complex in 
this ecological zone. No other species o f the An. gambiae complex were encountered 
in the study area. There have been no previous studies on species identification to 
determine which species of the An. gambiae complex occur in Adenta, Madina, East 
Legon and Achimota. However, Chinery (1984) in a study carried out in Accra 
(unidentified sites) suggested that An. arabiensis had displaced An. gambiae s.s. as the 
predominant species. The presence o f only An. gambiae s.s. at these sites in Accra as 
found by the present study clearly contradicts the observation by Chinery (1984). In a 
more recent study Appawu et al. (2001) have reported An. gambiae s.s. as the most
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important vectors of malaria in forest and savanna areas o f Ghana in terms o f its 
distribution and transmission. An. melas has also been reported in studies carried out 
along the coast (Appawu et al., 2001). But this was not encountered in the present 
study. This is not surprising since An. melas breeds in salt water and the study sites 
were far from the areas with salt pans and lagoons.
The finding that An. gambiae s..s. was the major sibling species in the forest area agrees 
with the general observation by Coluzzi et al. (1979) that it is the most prevalent 
species in the forest belts of West Africa. Although no An. arabiensis was encountered 
Coluzzi et al. (1979) have reported that An. arabiensis penetrates areas after 
deforestation and urbanization, but this species was not observed in the study area, 
especially Dodowa where considerable deforestation and urbanization have taken place.
Cytological analysis was also used in this study to identify the different karyotypes that 
occur in the study area and to seek evidence for distinctiveness of populations and gene 
flow. Karyotype analysis revealed the existence of the ‘Forest’ and ‘Savanna’ forms of 
An. gambiae s.s. in the study area. Only he ‘Forest’ form which is characterized by the 
standard arrangements (2R+, 2L+) on chromosome 2 was found at Dodowa and 
Adenta. At these two sites, all the 20 specimens studied were the forest form o f An. 
gambiae s.s. These findings are also in agreement with Appawu et al. (1985) who 
previously recorded the occurrence o f the forest form with inversions 2R+ and 2L+ in 
An. gambiae s.s. populations, from Dodowa, in the coastal forest zone of Ghana. The 
‘Forest’ karyotype was also found at East Legon and Achimota, occurring together with
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the ‘Savanna’ form, which is characterized by the chromosome 2R standard 
arrangement (2R+) together with inversion a on chromosomal arm 2L (2La). At these 
two sites, the ‘Savanna’ form was recorded in 2 out of 10 specimens studied at East 
Legon and in 4 out of 5 specimens studied at Achimota. This karyotype (2R+, 2La), 
which may be further differentiated by one or more 2R inversions (Toure et al., 1983; 
Coluzzi et al., 1985) is characteristic of the coastal savanna forms, which may be 
further differentiated by one or more 2R inversions. However, such further 
differentiation of the savanna form was not found in all the specimens that were 
studied
The observation that two distinct karyotypic populations were found in two different 
ecological zones is indicative of ecophenotypic plasticity of An. gambiae species 
(Powell et al., 1999), a factor influencing their efficiency as vectors o f Plasmodium. 
Moreover, the clinal geographical changes in karyotype frequencies of the ‘Savanna’ 
form from 0%to 80% from Dodowa to Achimota correlated with the change in 
vegetation from coastal forest to coastal savanna. Dodowa represents a degraded forest 
area whereas Adenta is a peri-urban area followed by Madina, East Legon and 
Achimota, which are urbanized areas. This trend is indicative o f the ecological 
adaptability/intergradations of karyotypes. The forest form of An. gambiae s.s has been 
found to be monomorphic for the standard 2R arrangement in other studies, whereas the 
savanna samples form becomes progressively more differentiated by inverted 
arrangements which increase in frequency eventually becoming fixed in drier savannas 
with frequencies ranging from 0% to 100% (Coluzzi etal., 1985).
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The agreement between the observed and expected karyotypic frequencies according to 
the Hardy-Weinberg law (HWE) suggests that each population is panmictic (random 
mating). The evidence for gene flow with respect to use o f karyotypes has been 
reported in other studies (Bryan et at., 1982, Toure et al., 1983, Robert et a l,  1990). 
Moreover, kaiyotype frequencies have been described as poor indicators of gene flow 
because their frequencies are apparently strongly affected by natural selection (Lanzaro 
et al, 1998). Hence the importance o f using other methods such as DNA analysis in 
addition to karyotyping for population studies
Lehmann et al. (1997; 1999) have reported that Fa values lower than 0.008 indicate 
high levels of population differentiation, and therefore an evidence for a division o f the 
gene pool. In the present study, it was observed that inter-population differentiation was 
highest (Fst= 0.0565) between the furthest population i.e. Dodowa and the four others 
Adenta, Madina, East Legon and Achimota. Dodowa is separated from the other 
populations by approximately 25 km. When East Legon and Achimota populations 
(8km apart) are compared, a lower differentiation index is reocrded (Fst =0.051), and 
this decreases further (Fst = 0.0284) when Adenta and East Legon, which are 2.5 km 
apart, are compared. It is therefore clear from the Fst values obtained from this study 
that differentiation between these populations is relatively high and increases with 
distances between them. These estimates of differentiation correspond to effective 
migration indices (N/w) that also increase with a reduction in distance (Table 7, Figure 
19). This observation is similar to previous studies investigating gene flow across the
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Rift valley in Kenya covering distances as wide apart as 400-500km and found that 
increasing distance contributed to differentiation across the Rift valley whereas closer 
populations were only marginally differentiated (Besansky et al., 1997; Kamau et al., 
1998; Lehmann etal., 1996).
The interpopulation differences observed is further supported by the estimates of 
heterozygote deficiency and excess (F;s) values, which indicate non-random mating 
between all populations except at Madina which geographically, is the centrally located 
point with respect to the study sites. The evidence for random mating in Madina but not 
at other populations suggests that it may be a hybrid zone where genetic exchange 
occurs between it and two adjacent populations, East Legon and Adenta. This evidence 
is further supported by the levels of gene flow, which are highest amongst these three 
populations compared to those that are further apart (Dodowa, Achimota).
The distribution of alleles at the three different loci was not uniform in all the five 
populations and this is indicative of the different levels of association at the different 
loci. The high differentiation index of Fst = 0.1282 recorded for the AGHX7 locus 
suggests that the populations are not related. On the other hand, the low levels of 
differentiation and heterozygote deficiency observed at the AG2H26 and AG2H46 loci 
may not necessarily be as a result of random mating but could also be due to natural 
selection resulting from ecological adaptation hence localizing their existence at certain 
localities. Another possible reason could be the presence of null alleles. In this study,
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the highest number of null alleles and homoyzgotes was observed at locus 2H46. In a 
study carried out on various loci including locus 2H46 in An. gambiae and An. 
arabiensis in East Africa, Lehmann et al. (1996b) inferred that locus 2H46 was under 
mutational constraint in An. gambiae. Null alleles, that is alleles that are not amplified 
as a result o f mutations at the primer binding sites can result in underestimation of 
heterozygosity because they are detected only in the homozygous state (Donnelly et al., 
1999).
At the multi-population level, genotype frequencies at all the three loci showed 
significant deviation (P<0.05) from the HWE due to lack of heterozygotes. Deviation 
from HWE is a common finding in studies that utilize microsatellite loci. This deficit is 
usually attributed to null alleles (Callen et al., 1993; Lehmann et al., 1996a, Garcia de 
Leon et al., 1997), selection (Garcia de Leon et al., 1997) or grouping o f gene pools (so 
called Wahlund effect) (Gibbs et al., 1997). It is often apparent that pooled populations 
show more evidence of departure from HWE consistent with a Wahlund effect from 
pooling subpopulations with unequal allele frequencies (Lehmann et al., 1996; Lanzaro 
et a l, 1998). But in this case, there were still substantial departures between local 
populations as well. Thus any heterozygote deficit at the loci is unlikely to have 
resulted from pooling samples from different populations with unequal allele 
frequencies (Wahlund effect) but could be due to the excess o f homozygotes which can 
be attributed to the presence o f  null alleles as a result of nucleotide substitutions in the 
primer annealing sites. It is likely that the presence of null alleles is the cause o f the 
heterozygote deficit observed in this study.
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Whereas the use of karyotypes to reveal details of species population differentiation is 
limited because of selection due to environmental adaptation, microsatellite DNA 
analysis because of their high polymorphism reveals a higher level of inter- and intra­
population variation supporting their description as powerful markers for measuring 
differentiation (Bowcock et al., 1994; Eustop et al., 1995). Taken together, the two data 
sets provided by karyotype and microsatellite DNA analysis in this study both support 
the view that physical distances separating An. gambiae populations, coupled with 
ecological adaptations may greatly influence the genetic structure o f these malaria 
vector populations. The results obtained indicate population subdivision and an inverse 
relationship between the distance and population genetic relatedness. The higher levels 
of gene flow (Nm>3) between An. gambiae s.s. populations may be attributable to 
human-mediated and other means of transport, as appetitive flights in anopheline 
species are usually 1 -2  km and this is not readily reconciled with the apparently high 
levels of gene flow observed.
In this study, the use of karyotyping and microsatellite DNA analysis has provided an 
insight into the cytotaxonomic status and level of differentiation of An. gambiae s.s. 
populations providing evidence for the existence of two distinct populations and 
confirming these two methods as useful tools in population studies. However, since no 
specimens from Madina were karyotyped, it was not possible to determine the presence 
of heterokaryotypes at this site as was indicated by microsatellite analysis. In view of 
the possible involvement of different members of the Anopheles gambiae complex in
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malaria transmission, there is a need to conduct more intensive studies covering a larger 
geographical area to define in more detail the species composition, distribution and 
temporal variation of the different chromosomal forms of An. gambiae s.s. in Ghana to 
determine their respective roles in malaria transmission. In effect, this will provide 
reliable information that can be used in the implementation o f disease control 
interventions and the possible management o f insecticide resistance.
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APPENDICES 
Appendix I 
Schedule of fixative and Stains
Carnoys Fixative
3 parts absolute alcohol 
1 part glacial acetic acid
Camoys fixative should be freshly prepared prior to use.
Lacto-acetic orcein stain
Stock solution 
lg orcein
25cc glacial acetic acid 
25cc 85% Lactic acid
This can be kept in a well stoppered bottle for long periods of time.
For orcein staining, 1 part of the stock solution is diluted with 3 parts 50% acetic acid.
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APPENDIX II 
STANDARD SOLUTIONS
The following standard solutions were prepared using sterile double distilled water (sddw). 
Where appropriate, the solutions were autoclaved at 121 lb/sq in. for 15 minutes in an Eyela 
Autoclave (Rikikakki Tokyo).
DNA Extraction
Bender buffer
0.1 M  NaCl, 0.2 M sucrose. 0.1 M  Tris-HCl pH 7.5, 0.05 M EDTA pH 9.1, 0.5% 
SDS. Stored at 4° C
0.5 M EDTA (pH8.0)
186.1 g/1 in water, ph adjusted with NaOH pellets and stored at room 
temperature.
EtBr(10mg/ml)
lg  of EtBr was completely dissolved in 100ml sddw and stored in the dark at room 
temperature.
KAc (5M K 8M Acetate)
60ml of 5 M KAc and 11.5 ml glacial acetic acid in 28.5 ml distilled water.
TE (pH 8.0), lOMm Tris-HCl (pH), ImM EDTA (pH8.0). Stored at room 
temperature.
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Solutions for Electrophoresis 
Agarose Gels
1 OX TAE buffer
242 g Tris base, 57.1 ml glacial acetic acid, 100ml 0.5MEDTA, pH adjusted to 7.7 
(with glacial acetic acid) and the volume made to 1000 ml with sddw.
0.5 M  EDTA (pH 8.0)
186g of EDTA, dissolved in 800ml ddw, pH adjusted with NaOH pellets, the 
volume made to 1000 ml with sddw and stored at room temperature.
Urea Polyacrylamide Gels
1 OX TBE electrophoresis buffer:
108g/l Tris base, 55g/l boric acid, 9.3 g/1 NaaEDTA. Use distilled water added to make 
up volume toll. Stored at room temperature on the bench. Diluted to IX working 
solution for electrophoresis.
40% Acrylamide
38g acrylamide, 2g bis-acrylamide in 100ml sddH20. Filtered and stored in the 
dark at 4°C.
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The table below gives the required amount of ingredients necessary to prepare
the solution for the 7% polyacrylamide running (separation) gel and the plug.
INGREDIENTS 5% POLYACRYLAMIDE GEL 
SOLUTION
Water (ml) 30.00
Urea (g) 8.25
40%Aciylamide (ml) 11.66
10X TBE Buffer (ml) 5 50
Ten microlitres ammonium persulphate (APS) and 5^1 of TEMED were added to 
2.4ml of the gel to plug the plates to avoid leakage.
Sixty microlitres of APS and 30jj,1 of TEMED were added to the rest of the solution 
(48ml) to form the separating gel, before pouring.
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The table below gives the required amount of ingredients necessary to prepare the stacking gel for a
5% polyacrylamide gel.
The solution for the 5% polyacrylamide running (separation) gel and the plug.
INGREDIENTS 4.5% POLYACRYLAMIDE GEL 
SOLUTION
Water (ml) 31.00
Urea (g) 8.25
3 0%Acrylamide/0.8 %Bis-acrylamide (ml) 7.50
10X TBE Buffer (ml) 5.50
Total 50.4 ml
3.5 ml of this solution were taken per gel, 30(j.l of APS and 10pl of TEMED were added 
and used as stacking gel. The gel was stained in EthBr.
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Gel Loading Buffers
6x Bromophenol blue
0.25% bromophenol blue was added to 40% sucrose in water 
and stored at 4° C.
Bromophenol blue xylene cyanol: 1 volume of bromophenol blue xylene 
cyanol and 4 volumes of cyanide
5X orange G
20%w/v Ficoll, 25Mm EDTA, 2.5mM EDTA. 2.5%(w/v) orange G. Stored 
at room temperature.
DNA Molecular weight size markers
The 100 bp DNA molecular weight size marker obtained from Sigma were diluted according to 
the manufacturers recommendations and used. For the 100 bp ladder, the first band size is 100 
bp, the subsequent ones are 200,300.............lOOObp.
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APPENDIX III
An example input data format for the population genetics analysis software POPGENE
VERSION 1 31
/*Diploid data of 5 populations each with 3 loci */ 
Number of populations =5 
Number of loci = 3 
locus name :
AGXH7 AG2H26 AG2H46
Name = DODOWA
AB..... .............. ..................................
AB..... .............. ..........................ABCCEE..
AB..... ..............AB..EFGG BBCC. ..
ABCC... AB....... ............ ABCCEE..
ABCC... AA..... ............ AB.......
....... ..................AB EF AB........
AB..EFGH AB..EF.. .......
ABCD...   AB..EE..
....... ..................AB..EEGH AB..EE..
ABCC.... ABCD .. AB. EE .
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APPENDIX IV
Standard Polytene Chromosome maps o f Anopheles gambiae
2L
' • ••{ /.. . .. 1r1  X-lliy s -. ' •  [  r i '  ,  «•  I  I '2 f t  " » i* « »; «
n lWl t f  <nV' ,v
i *
Chromosome 2
Chromosome 3
. c
' , V .
X
X Chromosome
153