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RESISTANCE MECHANISMS AND SUSCEPTIBILITY TO 
ORGANOPHOSPHATES, CARBAMATES AND PYRETHROIDS IN 
ANOPHELES GAMBIAE S.L. GILES (DIPTERA: CULICIDAE) IN 
HOHOE DISTRICT, GHANA.
BY
CHARO SAMUEL KAHINDI (B.Sc. Hons.)
University of Nairobi, Kenya.
A thesis submitted in partial fulfillment o f the requirements for the award o f  
Master of Philosophy degree in Entomology o f the University o f Ghana, Legon
Insect Science Programme* 
University o f Ghana, Legon
August 2005
*Joint Interfaculty International Programme for the training of entomologists in West 
Africa. Collaborating Departments: Zoology (Faculty of Science) &  Crop science 
(School of Agriculture and Consumer Sciences).
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DECLARATION
I certify that all material in this thesis which is not my own work has been duly 
acknowledged and that no material has previously been submitted and approved for the 
award of a degree by this or any other university.
Samuel K. Charo 
(Student)
(Supervisor)
Prof Michael D. Wilson 
(Supervisor)
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DEDICATION
To my entire Family and Friends, 
Your concern, love and support 
Kept me focussed.
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ACKNOWLEDGEMENTS
I extend my sincere gratitude and appreciation to all those who made this work 
possible. Special thanks go to my supervisors Dr. Daniel A. Boakye and Prof. 
Michael D. Wilson both o f the Parasitology Department o f  the Noguchi Memorial 
Institute for Medical Research (N.M.I.M.R) for their commitment, guidance, useful 
suggestions and encouragement throughout the period o f this study. Their invaluable 
comments helped to improve the quality o f  this work. I greatly appreciate the crucial 
role played by Rev. (Dr.) Winfred S.K. Gbewonyo o f the Department o f 
Biochemistry towards the success o f the biochemical aspect o f this work.
My sincere gratitude also goes to the Director N.M.I.M.R Prof. David Ofori-Adjei, 
for allowing me to carry out my work at the Institute. Special thanks to Mr. Maxwell 
Appawu of the Parasitology Department, N.M.I.M.R for allowing me to use his 
laboratory for the biochemical assays. I am greatly indebted to Dr. Charles Brown, 
Beverly Egyr, Yvonne Aryetey and Richard Larbi for their assistance in the 
molecular aspect of the work, to Philip Doku, Tony-Tetteh Kumah and Isiae 
Sibomana for their assistance in field collection and laboratory rearing o f  mosquitoes. 
A word o f  appreciation goes to all staff o f the Parasitology and Transport 
Departments, N.M.I.M.R. Special thanks to the village chiefs o f  Adabraka, Atabu 
Newtown, Kledzo and Likpe-Bakua, to the Director and staff o f  Hohoe District 
Hospital Onchocerciasis Research lab. for allowing me to work in their areas.
I appreciate the crucial role o f  all lecturers o f African Regional Postgraduate 
Programme in Insect Science (ARPPIS) for the excellent training and encouragement.
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My special thanks to Prof. John N. Ayertey for his invaluable contribution to the 
success o f this training. To my classmates, Abdullahi, Badi, Jacinter, Olivia and 
Zakka, I say it has been nice knowing you all. To Kipruto and Sospeter o f the 
Economic Management Programme, to Mamuye and Bismark o f N.M.I.M.R, you all 
made my social life wonderful.
I extend special thanks to my special friends o f Legon hall senior common room, 
Prof. E. Laing, Mr. E. A. Amartey and Mr. P. Yarkwah, I really enjoyed your 
company and fatherly advice. For all those who participated to the success o f  this 
work, whose names I have not mentioned, I thank you all. Finally, I thank the 
German Education Exchage Services (DAAD) and the Ghana Malaria Control 
Program through the N.M.I.M.R for sponsoring the work.
God Bless you all
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TABLE OF CONTENTS
TITLE PAGE.......................................................................................................................................... 1
DECLARATION................................................................................................................................... 11
DEDICATION...................................................................................................................................... 111
ACKNOWLEDGEMENTS.................................................................................................................. lv
TABLE OF CONTENTS......................... .............................................................................................'n
LIST OF FIGURES............................................................................................................................... “
LIST OF TABLES................................................................................................................................. x
LIST OF ABBREVIATIONS............................................................................................................... xi
LIST OF APPENDICES...................................................................................................................... xii
ABSTRACT........................................................................................................................................xiii
CHAPTER ONE................................................................................................................................... 1
GENERAL INTRODUCTION........................................................................................................... 1
1.0 Introduction......................................................................................................................................1
1.2 Rationale and Objectives............................................................................................................. 4
1.2.1 General objective....................................................................................................................5
1.2.2 Specific objectives...................................................................................................................5
CHAPTER TWO..................................................................................................................................6
LITERATURE REVIEW....................................................................................................................6
2.1 Malaria: Disease and Symptoms..................................................................................................6
2.2 Life cycle and transmission of malaria parasites.........................................................................7
2.3 Global distribution of Malaria....................................................................................................11
2.4 Socioeconomic burden and risk of malaria................................................................................13
2.5 Biology and Life Cycle of the Vectors; the Genus Anopheles..................................................15
2.5.1 Anopheles gambiae Giles complex.....................................................................................19
2.6.0 Malaria control........................................................................................................................20
2.6.1 Historical background of malaria control...........................................................................20
2.6.2.Mortality control.................................................................................................................21
2.6.3 .Transmission control..........................................................................................  22
2.6.3.1 Environmental management.......................................................................  23
2.6.3.2 Intradomicile Application of Residual Insecticides....................................................24
2.6.3.3. Insecticide Treated Materials......................................................................  26
2.6.3.4. Personal protection..........................................................................  2 7
2.6.3.5 Transgenic Mosquitoes.............................................................  2R
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2.6.4 Multilateral Malaria Research and Control programs........................................................
...... 30
2.7 Insecticide Resistance........................................................................................................
_ 30
2.7.1 Mechanisms of Insecticide Resistance...............................................................................
2.7.1.1 Reduced penetration.................................................................................................... 31
2.7.1.2.Metabolic resistance.................................................................................................... J3 1*
2.7.1.2.1. Glutathione S-transferases.................................................................................. 32
2.7.1.2.2 Mixed Function Oxidases (MFO)........................................................................ 32
2.7.1.2.3. Esterases and Hydrolases.................................................................................... 34
2.7.1.3..Target-Site Mechanisms............................................................................................. 35
2.7.1.3.1 Knock-down resistance........................................................................................ 36
2.7.1.3.2 Insensitive acetylcholinesterase........................................................................... 38
2.7.1.3.3 GABA Receptor Changes.................................................................................... 49
2.8 Management of Insecticide Resistance..................................................................................40
CHAPTER THREE........................................................................................................................... 42
MATERIALS AND METHODS....................................................................................................... 42
3.1 Study area..................................................................................................................................42
3.2 Field collection of mosquitoes...................................................................................................44
3.2.1 Adult mosquito collection..................................................................................................44
3.2.2 Larval mosquito collection.................................................................................................44
3.2.3 Laboratory rearing of mosquitoes...........................................................................................49
3.2.4 Morphological Identification of Anopheles gambiae s.l. Mosquitoes....................................51
3.2.3 Preservation of mosquito samples...........................................................................................51
3.3.Bioassay experiments.................................................................................................................52
3.4 Molecular Studies......................................................................................................................55
3.4.1 DNA Extraction..................................................................................................................5 5
3.4.2 PCR Identification of Anopheles gambiae complex...........................................................56
3.4.4 Identification of the molecular forms of Anopheles gambiae s .s .......................................57
3.4.5 PCR detection of the kdr alleles in Anopheles gambiae complex......................................58
3.5 Biochemical assays....................................................................................................................6 1
3.5.1 Sample Preparation.............................................................................................................61
3.5.2.Calibration curves.................................................................................................................
3.5.2.1 Protein......................................................................................................................  6 3
3.5.2.2 Oxidase..................................................................................................  6 3
3.5.2.3 Non- specific esterases.....................................................................  6 4
3.5.3 Enzyme activity assays...................................................................  6 4
3.5.3.1 Protein assay................................................................  ^
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3.5.3.2 Oxidase assay..............................................................................................................
3.5.3.3 Acetylcholinesterase assay..........................................................................................
3.5.3.4 Glutathione S-transferases........................................................................................... ^6
3.5.3.5 Non-specific esterases................................................................................................. ^
3.6 Data Analysis......................................................................................................................... ^
CHAPTER FOUR.............................................................................................................................. 68
RESULTS............................................................................................................................................ 68
4.1 Bioassays...................................................................................................................................68
4.1.1 Permethrin (0.75%)........................................................................................................... 69
4.1.2 Deltamethrin (0.05%)........................................................................................................ 69
4.1.3 DDT (4%).......................................................................................................................... 69
4.1.4 Malathion (5%).................................................................................................................. 70
4.1.5 Propoxur (0.1%)................................................................................................................70
4.1.6 Resistance ratios................................................................................................................ 72
4.2 Molecular studies.......................................................................................................................75
4.2.1 Species and molecular forms identification........................................................................75
4.2.2 Distribution of the Kdr allele in An. gambiae s.s. population...........................................75
4.3 Biochemical assays....................................................................................................................81
4.3.1 Calibration curves...............................................................................................................81
4.3.2 Total protein content...........................................................................................................83
4.3.3 Oxidase...............................................................................................................................83
4.3.4 Acetylcholinesterase...........................................................................................................86
4.3.5 Glutathione-S-Transferase..................................................................................................88
4.3.6 Non-specific esterase assay................................................................................................90
4.3.7 Level of increase in enzyme activity..................................................................................9 5
CHAPTER FIVE................................................................................................................................ ..
DISCUSSION.....................................................................................................................................97
REFERENCES...................................................................................................................................
APPENDICES................................................................................................................................... ..
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LIST OF FIGURES
Figure 2.1 Global distribution of Malaria
Figure 2.2 The Life cycle stages o f  malaria parasites
Figure 2.3 Life cycle of stages o f  Anopheles mosquitoes
Figure 2.4 Differences between the anophelines and other common mosquitoes
Figure 3.1 Typical adult collection house at Adabraka
Figure 3.2 Typical adult collection house at Atabu Newtown
Figure 3.3 Typical larval collection site at Atabu Newtown
Figure 3.4 Typical larval collection site at Adabraka
Figure 3.5 Laboratory mosquito rearing system
Figure 3.6 Bioassay experiment set-up
Figure 3.7 A flat bottomed microtiter plate used for biochemical assays
Figure 4.1 Mean percent mortalities recorded for pyrethroids and DDT
Figure 4.2 Mean percent mortalities recorded for Malathion and Propoxur.
Figure 4.3 An electrophoregram o f identification o f  Anopheles gambiae s.l.
Figure 4.4 An electrophoregram of Anopheles gambiae s.s. molecular forms
Figure 4.5 An electrophoregram o f kdr Alleles
Figure 4.6 Calibration curves
Figure 4.7 Distribution patterns o f Oxidase activity
Figure 4.8 Distribution patterns o f AcCHE activity
Figure 4.9 Distribution patterns o f  GST activity
Figure 4.10 Distribution patterns o f a-esterase activity
Figure 4.11 Distribution patterns o f /3-esterase activity
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LIST OF TABLES
Table 1 DNA sequences of synthetic oligonucleotide primers used for the PCR- 
based method o f identification o f Anopheles gambiae s.l., their melting 
points and the diagnostic band sizes o f the amplified products (Scott et al., 
1993).
Table 2 DNA sequences of synthetic kdr oligonucleotide primers.
Table 3 Knockdown times for Permethrin, Deltamethrin and DDT in the four wild 
populations and the susceptible Kisumu strain.
Table 4 Molecular identification o f Anopheles gambiae s.s samples from four field 
populations into M and S forms.
Table 5 Distribution of kdr alleles in the ‘M ’ and ‘S’ forms o f  Anopheles gambiae 
s.s. in the four wild populations.
Table 6 Mean activity of Oxidase, Acetylcholinesterase and Glutathione-S- 
transferase in the four wild populations and the susceptible Kisumu s t r a in
Table 7 Mean activity o f Non-specific esterases in the four wild populations and 
the susceptible Kisumu strain.
Table 8 Level o f increase in enzyme activity.
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LIST OF ABBREVIATIONS
bp base pairs
ddw distilled de-ionised water
dCPT deoxycytidine triphosphate
dGTP deoxyguanosine triphosphate
dATP deoxyadenosine triphosphate
DNA deoxyribonucleic acid
dNTP deoxyribonucleotide phosphate
dTTP deoxythymidine triaphosphate
EDTA disodium ethylene diamine tetracetate.2H20
EtBr ethidium bromide
EtOH ethanol
H20 water
M molar (moles per liter)
mM millimolar
juM micromolar
ml milliliter
fil microliter
Mw molecular weight
NaOH sodium hydroxide
KAc potassium acetate
pH -log.o [H+]
rpm revolution per minute
RNase ribonuclease
s.l. sensu lato
s.s. senso stricto
Tm melting temperature
g gram
mg milligram
Mg microgram
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LIST OF APPENDICES
Appendix I Standard solutions
Appendix II W.H.O. susceptibility results 
Appendix II Biochemical assays results
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ABSTRACT
Malaria is a threat to more than 40% of the world's population accounting for more 
than 300 million acute cases and between 1.1 and 2.7 million deaths annually. Over 
90% of the malaria cases are in sub-Saharan Africa, constituting 10% of the total 
disease burden. Currently activities support the extensive use of insecticide treated 
materials for malaria control. However these efforts are threatened by the evolution of 
resistance in the main malaria vectors, Anopheles gambiae s.l. towards the commonly 
used insecticides for mosquito control. The study was done to determine the 
susceptibility status and the underlying mechanisms of resistance in An. gambiae s.l 
against commonly used insecticides for control in Hohoe District, Volta region of 
Ghana. Mosquitoes were sampled in four villages; Adabraka, Atabu Newtown, 
Kledzo and Likpe-Bakwa. Adult mosquitoes were tested against the WHO diagnostic 
concentrations of 0.75% permethrin, 0.05% deltamethrin, 4% DDT, 5% malathion 
and 0.1% propoxur. The susceptible Kisumu strain of An. gambiae s.l was used as the 
reference.
Results obtained revealed high levels of resistance to DDT; 6-51% mortality rates in 
the four villages sampled. Susceptibility to Permethrin was considerably low with 32­
82% mortality rates. Mortalities were very high with 0.05% Deltamethrin; 91-97%. 
All field mosquitoes tested were fully susceptible to Malathion with 100% mortality 
rates across the 4 villages. Susceptibility to Propoxur was similarly higher; 94-95% 
mortality rates.
Median Knockdown times in field populations variously increased compared with the 
susceptible Kisumu strain; 4-6, 1.5-2 and 3 fold with Permethrin, Deltamethrin and
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DDT respectively. PCR identification revealed that all the mosquitoes tested were An. 
gambiae sensu stricto with 64% ‘Savanna’ and 36% ‘Mopti’ forms. The kdr mutation 
occurred at a frequency of 23.7%, 21.7%, 22.7% and 31% in An. gambiae 
populations of Adabraka, Atabu Newtown, Kledzo and Likpe respectively. Over 70% 
of the kdr mutation occurred in the ‘Savanna’ form. Biochemical mechanisms of 
resistance were investigated by the CDC microplate assays protocol. There was a 
significant elevated activity of mixed function oxidases in An. gambiae populations in 
Likpe as compared to the susceptible Kisumu strain. Activity of Acetylcholine 
esterase enzyme was significantly elevated in Adabraka population while Glutathione 
S-transferases showed no significant increase in activity in all the four wild 
population. There was a significant elevated activity of both α  and β-nonspecific 
esterases in Adabraka and Likpe populations. The high frequency of kdr and elevated 
activity of several detoxification enzymes indicate the occurrence of multi resistance 
in An. gambiae populations in Hohoe district.
Insecticide resistance has been a problem in all insect groups that are vectors of 
diseases. Regular testing is therefore vital for mosquito control operations because 
resistant populations of mosquitoes reduce the effectiveness of control procedures. 
The main defence against resistance is close surveillance of the susceptibility of 
vector populations so as to detect changes in their susceptibility status at an early 
stage and to implement resistance management strategies.
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CHAPTER ONE
GENERAL INTRODUCTION 
1.1 Introduction
Malaria is a life-threatening parasitic disease affecting more than 40% o f the worlds 
population and out o f the more than 300 million acute cases each year, between 1.1 and 2.7 
million people die (RBM, 2002; WHO, 2000). Over 90% o f malaria cases are in sub-Saharan 
Africa, where malaria accounts for 10% o f the total disease burden. Children under five and 
pregnant women are most at risk (TDR/WHO, 2002; RBM/WHO, 2000). Malaria constitutes 
nearly 25% o f all childhood mortality in Africa (WHO, 2000).
Malaria is caused by protozoan parasites o f  the genus Plasmodium  and is transmitted 
amongst humans by mosquitoes of the genus Anopheles. There are 120 Plasmodium species, 
of which four; P. falciparum, P. vivax, P. malariae and P. ovale are known to infect humans. 
All the four species share a common basic life cycle although there are considerable 
differences in their pathogenecity, epidemiology, appearance, development, and host-parasite 
relationships. It is P. falciparum, however, that causes nearly all o f the mortality in cases o f  
malaria infection. Mosquitoes of the genus Anopheles are always the vectors and out o f the 
more than 380 species of anopheline mosquitoes, only 60 can transmit malaria. Only female 
mosquitoes are involved in transmission as the males do not require a blood meal.
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The World Health Organization suggests that there are three essential elements o f malaria 
control (WHO, 1995). First is the selective application o f vector control by reduction o f  the 
numbers o f vector mosquitoes and reducing human-mosquito contact. Secondly, the early 
diagnosis followed by effective and prompt treatment o f malaria cases in all areas where 
people are at risk, and the third element is early detection or forecasting o f epidemics and 
rapid application of control measure.
Since malaria is transmitted by the female Anopheles mosquito, a major strategy o f  control is 
to attack the vector with insecticides. Extended use, however, has led and continues to lead to 
the emergence of insecticide resistance in mosquitoes (Hemingway and Ranson, 2000; 
Philips, 2001). There has recently been an increase in antimalarial activities with the Roll 
Back Malaria initiative and Global Fund for Health, which support extensive use o f 
pyrethroid-impregnated bed nets for mosquito control campaigns in Africa and other malaria- 
endemic regions (Curtis et al, 1998; Akogbeto and Yakoubou, 1999). However these 
activities are greatly threatened by the evolution o f resistance among malaria vectors towards 
the commonly used insecticides to control them (Chandre et al., 1999a,b; Hougard et al., 
2003; Corbel et al., 2003; Curtis et al., 2003). Furthermore, insecticide resistance is assumed 
to increase the likelihood of mosquito-borne disease transmission by increasing the vector 
population size and allowing mosquitoes to live longer in the presence o f insecticide 
(McCarroll and Hemingway 2002).
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Mosquitoes can develop resistance through several different mechanisms (Hemingway and 
Ranson, 2000). Physiological resistance is one way mosquitoes can become immune to 
insecticides. For example, reduced penetration through the cuticle has been noted in several 
mosquito species. This gives the detoxification mechanisms in the mosquito more time to 
deal with uptake o f the toxicant hence the greater its chances for survival (Hemingway and 
Ranson, 2000). Insecticide resistance can also result from reduced sensitivity o f the target 
sites that normally bind to the insecticides. For example, mutations in sodium channels (the 
target of DDT and pyrethroids) and in acetylcholinesterase (the target o f organophosphates 
and carbamates) have been well documented in many insect species including mosquitoes 
(Hemingway et al., 2004). Metabolic resistance occurs when detoxification enzymes are used 
to break down the insecticide into compounds such as amino acids and sugars, which can be 
metabolized by the mosquito.
Insecticide resistance has been a problem in all insect groups that serve as vectors o f 
emerging diseases (Hemingway and Ranson, 2000). Although mechanisms by which 
insecticides become less effective are similar across all vector taxa, each resistance problem 
is potentially unique and may involve a complex pattern of resistance foci. Several strategies 
and recommendations have been proposed for the management o f  insecticide resistance in 
field populations. The most important aspect o f  the management o f resistance is to either 
avoid or delay the onset o f resistance by using the available insecticides judiciously, for 
example as mixtures, in rotation or in mosaics. The practice o f  using an insecticide until 
resistance becomes a limiting factor is rapidly eroding the number of suitable insecticides for 
vector control. Thus, the main defense against resistance is close surveillance o f the 
susceptibility o f vector populations.
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1.2 Rationale and Objectives
Regular resistance testing is vital for mosquito control operations because resistant 
populations of mosquitoes reduce the effectiveness of control strategies. The resistant 
phenotype is relatively easy to monitor with direct insecticide bioassays. However, in many 
cases the actual molecular and biochemical mechanisms responsible for the resistant 
phenotypes are still unknown. Currently, the mechanisms that regulate insecticide resistance 
are poorly understood. Anopheles gambiae s.l. has multiple resistance mechanisms that have 
been field-selected in both East and West Africa through exposure to DDT and pyrethroids 
(Hemingway et al., 2002). It is not clear how much the current large-scale pyrethroid 
resistance of mosquitoes in West Africa will affect the extensive use o f  pyrethroid 
impregnated bed nets for mosquito control campaigns in Africa, and what will replace the 
pyrethroid-treated nets if  selection o f multi-resistance mechanisms results in widespread 
failure of this strategy (Curtis et al., 1998).
In Ghana several studies on susceptibility/resistance on An. gambiae complex to insecticides 
have been conducted, however they have been focused mainly in the Greater Accra Region 
and its environs (Adasi et al., 2000; Adeniran, 2002; Otieno, 2004) and in the Western 
Region (Kristan et al., 2003). It is therefore imperative that more studies on 
susceptibility/resistance in malaria vectors towards the commonly used insecticides be 
carried out in different ecological zones o f  Ghana in order to obtain broader baseline 
information so as to enable the development malaria vector control programs on national 
scale and to implement resistance management strategies. Furthermore due to the complex 
interplay o f factors conferring resistance in mosquitoes, there is need for an in-depth study on 
the underlying mechanisms of resistance especially the biochemical mechanisms in An.
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gambiae complex, in addition to the knockdown resistance mechanism o f which was the 
focuss of most o f the earlier studies in the country.
The present study was therefore carried out to gather information on susceptibility profiles 
and the underlying mechanisms of resistance against the commonly used insecticides in An. 
gambiae s.l. populations from Hohoe district in the Volta Region, an area in Ghana where no 
such information exist.
1.2.1 General objective
The general objective o f the study was to determine the susceptibility status and the 
mechanisms of resistance in Anopheles gambiae s.l Giles to the commonly used insecticides 
in the Hohoe area.
1.2.2 Specific objectives
1. To determine the susceptibility status of adult Anopheles gambiae s.l to Permethrin, 
Deltamethrin, DDT, Malathion and Propoxur using WHO adult bioassay methods.
2. To identify Anopheles gambiae s.l mosquitoes to species and also the molecular 
forms o f Anopheles gambiae s.s.
3. To characterize the kdr alleles in populations of Anopheles gambiae s.l using a PCR 
based method.
4. To determine the operative biochemical mechanisms o f resistance by conducting 
enzyme assays for Oxidase, Acetylcholinesterase, Non-specific esterases and 
Glutathione S-transferase.
5. To relate data on bioassays with those o f  kdr and enzyme assays.
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CHAPTER TWO
LITERATURE REVIEW
2.1 Malaria: Disease and Symptoms
The term malaria is Roman in origin, although the disease was not known by its present 
name until the mid-eighteenth century. Before then it was referred to variously as ague, 
intermittent fever, swamp fever, Roman fever, and death fever. Previously, it was thought 
that "miasma" (bad air or gas from swamps - "mal air ia") caused the disease. Malaria has 
been known since time immemorial, but it was centuries before the true causes were 
understood. Hippocrates was the first to describe the manifestations o f the disease, and relate 
them to the time of year and to where the patients lived before this, supernatural were 
blamed.
There are four species of malaria parasites in humans; Plasmodium falciparum, P. vivax, P. 
malariae, and P. ovale (Phillips, 2001). Out o f the four, two are most common; Plasmodium 
falciparum, which is found globally but is commonest in Africa, is the most aggressive 
species, often killing by coma or anaemia (Miller et al., 2002). Plasmodium vivax, which 
ranges widely throughout Asia, Africa, the Middle East, Oceania and the Americas, can 
cause recurring and debilitating infection, but rarely kills.
The different malaria parasites produce fevers o f  different frequency, depending on how long 
it takes to complete shizogony in erythrocytes. Generally the patient will complain o f 
headache, fever and aches and pains all over the body. However, fever is not always present 
and rigours may or may not be present. At its peak, a person's fever can soar to 41°C (106°F). 
Several hours later, the fever drops and chills set in. Two to four days later, the cycle repeats.
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Diarrhoea and abdominal pain are sometimes present. Spleen and liver are often palpable on 
clinical examination. This may be misdiagnosed as influenza in non-endemic areas and 
unless treated promptly, the clinical picture can deteriorate rapidly. A patient with severe and 
complicated malaria will often present with impaired consciousness, weakness, and jaundice.
Other complications are cerebral malaria, generalised convulsions, normocytic anaemia, 
renal failure, hypoglycaemia, fluid, electrolyte and acid-base disturbances, pulmonary 
oedema, circulatory collapse, shock, disseminated intravascular coagulation, hyperpyrexia, 
hyperparasitaemia, and malarial haemoglobulinurea. These features may occur singly or in 
combinations. Severe and complicated malaria is usually caused by delay in treating an 
uncomplicated attack o f P. falciparum. Cerebral malaria is the most dreaded form o f disease 
and is unique to P. falciparum. Red blood cells infected by the parasite are sticky and can 
gum up the capillaries of the brain (Marsh, 1992; Miller et al., 2002). The victim enters a 
coma and even if  death does not occur, brain damage can be the result. Death can strike in as 
little as 24 hours from first symptoms. Anemia is another threat due to the parasite's cyclical 
attacks and rupture o f red blood cells.
2.2 Life cycle and transmission of malaria parasites
The Plasmodium parasites have a life cycle which is split between a vertebrate host and an 
insect vector (Figure 1). Briefly, a biting female Anopheles mosquito transfers about 10% of 
its sporozoite load on each occasion, in her saliva, into the circulating blood o f  the host and 
within 30 to 45 minutes have entered hepatocytes. Growth and division in the liver for the 
human malaria parasites take from approximately 6 to 15 days depending on the species,
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approximately 6, 10 and 15 days for P falciparum, P  vivax, and P  ovale and P. malariae, 
respectively. At the end o f the pre-erythrocytic cycle, thousands o f  merozoites are released 
into the blood flowing through the sinusoids and, within 15 to 20 seconds, attach to and 
invade erythrocytes.
Recognition and attachment are via a receptor-ligand interaction. In P. vivax and P. ovale, 
some of the sporozoites appear to develop for about 24 hours before becoming dormant as a 
hypnozoite stage. This form can remain as such for months and even years until reactivated 
to complete the liver cycle, releasing merozoites into the blood to precipitate a relapse 
infection.
The asexual erythrocytic cycle produces more merozoites that are released with the 
destruction of the red blood cells after 48 or 72 hours for the human malaria parasites, 
depending on the species, and which then immediately invade additional erythrocytes. 
Consequently, they complete schizogony together at the end of the asexual cycle, releasing 
pyrogenic materials which induce the characteristic fever spike and clinical symptoms. The 
morbidity and mortality associated with malaria are derived solely from the erythrocytic 
stages (Miller et al., 2002). The asexual cycle usually continues until controlled by the 
immune response or chemotherapy or until the patient dies (in the case o f  P. falciparum).
After invading red blood cells, eventually some merozoites differentiate into sexual forms 
(gametocytes) which have no further activity within the human host and, following ingestion 
by another female mosquito, will mature to male and female gametes in the blood meal.
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After fertilization, the resulting zygote matures within 24 hours to the motile ookinete, which 
burrows through the midgut wall to encyst on the basal lamina, the extracellular matrix layer 
separating the haemocoel from the midgut. Within the developing oocysts, there are many 
mitotic divisions resulting in oocysts full o f sporozoites. Rupture o f the oocysts releases the 
sporozoites, which migrate through the haemocoel to the salivary glands to complete the 
cycle approximately 7 to 18 days after gametocyte ingestion, depending on host-parasite 
combination and external environmental conditions.
Not all Anopheles mosquitoes are vectors for Plasmodium parasites and these refractory 
mosquitoes posses substances toxic to Plasmodium within their cells (Beier, 1998). A higher 
trypsin-like activity has also been found in the midgut o f resistant mosquito species, possibly 
inhibiting ookinete development. Sporogony within the mosquito is governed by 
environmental temperature as anopheline mosquitoes are poikilotherms. The female 
anopheline mosquitoes' ability or competence to transmit malaria is governed by a complex 
interaction o f environment, behavioural and biological features, including vector density, 
blood meal preference, feeding and resting habits, flight range, longevity, humidity and 
temperature.
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Figure 2.1: The Life cycle stages of malaria parasites (Phillips, 2001).
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2.3 Global distribution of Malaria.
Malaria is generally endemic in the tropics, with extensions into the subtropics. Previously 
extremely widespread, malaria is now mainly confined to Africa, Asia and Latin America 
(Figure 2.2). Malaria affects the lives o f almost all people living in the area o f Africa defined 
by the southern fringes of the Sahara Desert in the north, and a latitude of about 28° in the 
south (WHO/UNICEF, 2003). Most people at risk o f  the disease live in areas o f  relatively 
stable malaria transmission where infection is common and occurs with sufficient frequency 
that some level o f immunity develops. A smaller proportion o f people live in areas where risk 
o f malaria is more seasonal and less predictable, because o f either altitude or rainfall patterns. 
People living in the peripheral areas north or south o f the main endemic area or bordering 
highland areas are vulnerable to highly seasonal transmission and to malaria epidemics.
Malaria is responsible globally, for 500 million cases o f  clinical disease and presents a public 
health problem for 2.4 billion people, representing over 40% o f the world’s population in 
over 90 countries. Almost 10% o f the world’s population will suffer a clinical attack o f 
malaria each year (Phillips, 2001). About 90% o f all malaria deaths in the world today occur 
in Africa south of the Sahara. This is because the majority o f  infections in Africa are caused 
by P. falciparum, the most dangerous o f the four human malaria parasites. It is also because 
the most effective malaria vector, An. gambiae, is the most widespread in Africa and the 
most difficult to control (Greenwood and Mutabingwa, 2002).
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2.4 Socio-economic burden and risk of malaria
“Where malaria prospers most, human societies have prospered least” (Sachs and Malaney, 
2002). Malaria's cost to human and social well-being is enormous. It is a major cause o f 
poverty and poverty exacerbates the malaria situation (UNICEF, 2000). So too is the 
economic loss, which in Africa alone is estimated at more than $2 billion annually (WHO, 
2000). The disease has slowed economic growth in African countries by 1.3% per year, the 
compounded effects o f which are a gross domestic product level now up to 32% lower than it 
would have been had disease been eradicated from Africa in 1960 (RBM, 2002). Malaria is 
most intractable for countries in the poorest continent, Africa (Gallup and Sachs, 2001) and 
poor countries predominate in the same regions as malaria (Miller et al., 2002). Almost all o f 
the rich countries are outside the bounds of intensive malaria.
In all malaria-endemic countries in Africa, 25-40%  of all outpatient clinic visits are for 
malaria (with most diagnosis made clinically). In these same countries, between 20% and 
50% o f all hospital admissions are a consequence of malaria. With high case-fatality rates 
due to late presentation, inadequate management, and unavailability or stock-outs of effective 
drugs, malaria is also a major contributor to deaths among hospital inpatients. Malaria is 
responsible for a high proportion of public health expenditure on curative treatment, and 
substantial reductions in malaria incidence would free up available health resources and 
facilities and health workers’ time, to tackle other health problems (WHO/UNICEF, 2003).
In areas of stable malaria transmission, children and pregnant women are the population 
groups at highest risk of morbidity and mortality. Most children experience their first malaria
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infections during the first year or two of life, when they have not yet acquired adequate 
clinical immunity, which makes these early years particularly dangerous (Greenwood and 
Mutabingwa, 2002). Ninety percent o f all malaria deaths in Africa occur in children aged 
below 5 years. At least 20% of all deaths in children in this age group is due to the disease. 
Children who survive malaria may suffer long-term consequences o f the infection. Repeated 
episodes of fever and illness reduce appetite and restrict play, social interaction and 
educational opportunities, thereby contributing to poor development. An estimated 2% o f 
children who recover from malaria infections affecting the brain (cerebral malaria) suffer 
from learning impairments and disabilities due to brain damage including epilepsy and 
spasticity.
Adult women in areas o f stable transmission have a high level o f  immunity, but the normal 
weakening of the immune system especially during the first pregnancy makes infection more 
likely and the anaemia associated with pregnancy gives the parasite an advantage. Pregnant 
women are four times as likely to get the disease, and half as likely to survive cerebral 
malaria. If they do, their foetus may not as the extreme fevers often cause spontaneous 
abortion and stillbirths.
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2.5 Biology and Life Cycle of the Vectors; the Genus Anopheles
Anopheles mosquitoes belong to the order Diptera, Suborder Nematocera, Family Culicidae 
and Subfamily Anophelinae. They have a worldwide distribution, occurring in both tropical 
and temperate regions. O f the over 500 known species of Anopheles, only about 60 are able 
to transmit malaria. Important malaria vectors include An. culicifacies in South West Asia, 
An. darlingi in North America and An. albimanus in Central America (Service, 1993). In 
Africa, they include An. pharoencis in Egypt, An. gambiae complex and An. funestis in West 
and East Africa (Nchinda, 1998).
Like all mosquitoes, anophelines go through four stages in their life cycle: egg, larva, pupa, 
and adult (Figure 2.3). The first three stages are aquatic and last 5-14 days, depending on the 
species and the ambient temperature. Eggs are generally laid directly on water or damp soil, 
often in tiny bodies o f water such as those formed by flooded hoof prints or tyre tracks. The 
female anopheline deposits eggs singly on the water surface and the boat-shaped eggs have 
some grooves on each side to keep them afloat. Eggs hatch in approximately 2 to 3 days after 
oviposition, although some larvae can remain quiescent in unhatched eggs on damp soil for 
up to 2 weeks, thus the population can survive periods o f  erratic rains (Robert and Collins, 
1996).
Larva hangs suspended by surface tension, breathing through air tubes and feeding on micro­
organisms by filtering the water. Larval development is rapid and can be completed in less 
than one week in very warm conditions and ample food. Larvae develop through 4 stages, or 
instars, after which they metamorphose into pupae. At the end o f each instar, the larvae
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moult, shedding their exoskeleton to allow for further growth. Pupation typically takes place 
in full sunlight. The coma-shaped pupa is active and does not feed but has to come to the 
water surface to breathe. Pupal stage is short, approximately 1 to 2 days and adult emerges 
typically at sunset.
Both male and female adult require at least 24 hours to reach sexual maturity and mating is 
associated with a swarming behaviour. A swarm consists mainly o f males, and females fly 
into it to mate, which typically takes place after sundown. Mated females seek blood meals 
only at night. Egg development takes about 2 days during the warm season, but it can be 
longer in cooler months. Oviposition like blood-feeding also occurs at night. Consequently, 
the gravid female generally lays her eggs the second night after she has blood fed and after 
oviposition searches for another blood meal. This repeated feeding and oviposition has major 
implications for transmission o f  malaria parasites which have a required developmental cycle 
in the mosquito (Robert and Collins, 1996).
Since anopheline mosquitoes typically breed in stagnant, unpolluted surface waters they are 
mainly associated with rural settings. Anopheles mosquitoes can be distinguished from other 
mosquitoes by the palps, which are as long as the proboscis, and by the presence o f discrete 
blocks of black and white scales on the wings. Adult Anopheles can also be identified by 
their typical resting position as males and females rest with their abdomens sticking up in the 
air rather than parallel to the surface on which they are resting (figure 2.4).
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Female Anopheles 
Mosquito
ZZM
9 JPQc
Eggs (laid singly)
Pupa
Larva
Figure 2.3: Life cycle stages of Anopheles mosquitoes
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A N O P H E L I N E S C U L IC IM E S
A N O PH ELES AEDES CULEX
a f * 4 ^ \
* ^
m  f m  f m f
z
Q
1gth—/i
LOUS
------------------------------
Figure 2.4: Differences between the anopheline mosquito and other common mosquitoes 
(UNICEF, 2000).
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2.5.1 Anopheles gambiae Giles complex
Anopheles gambiae complex, the major malaria vector in sub-Saharan Africa, is probably the 
most important vector o f a human pathogen in the world. The complex consists o f seven 
species with varying abilities in malaria transmission (Coluzzi et al., 2002). These are An. 
gambiae sensu stricto, An. arabiensis, An. merus, An. melas, An. quadriannulatus and An. 
bwambae. The seventh species is An. quadriannulatus B from Ethiopia. Two species o f  the 
complex; An. gambiae s.s and An. arabiensis axe the most widely distributed and the most 
efficient malaria vectors in West Africa (Coetzee et al., 2000; Coluzzi et al., 2002).
Anopheles gambiae s.s, the nominal taxon, is the most anthropophilic member o f  the 
complex and the main malaria vector in sub-Saharan Africa (Pates et al., 2001). Anopheles 
gambiae s.s, is split into five chromosomal forms characterized by the presence or absence of 
paracentric inversions on the second chromosome (Lehmann et al., 2003; Gentile et al., 
2002). These are the ‘Savanna’, ‘Mopti’, ‘Forest’, ‘Bissau’ and the ‘Bamako’ forms. 
Anopheles merus, An. melas and An. bwambae are very minor malaria vectors while An. 
quadriannulatus is zoophilic and o f  no importance as far as malaria transmission in humans 
is concerned (Coetzee et al., 2000).
Anopheles gambiae complex has an extreme preference for living around human habitations 
and is highly anthropophilic. Typical oviposition sites are small temporary bodies o f water 
exposed to full sunlight, such as small puddles produced by rain. The number o f  available 
larval habitats increases during rainy seasons which makes the annual abundance o f this 
mosquito correlates highly with rainfall.
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2.6.0 Malaria control
The control o f malaria today is mainly focussed on the mosquito vector and on the parasite.
2.6.1 Historical background of malaria control
Although people were unaware of the origin o f  malaria and the mode o f transmission, 
protective measures against the mosquito have been used for many hundreds o f years. The 
association with stagnant waters (breeding grounds for Anopheles) led the Romans to begin 
drainage programs, the first intervention against malaria. The inhabitants o f swampy regions 
in Egypt were recorded as sleeping in tower-like structures out o f the reach o f mosquitoes, 
whereas others slept under nets as early as 450 B.C. Ways o f dealing with malaria followed 
its proliferation, from administering the herb qinghaosu as a treatment in China more than 
2000 years ago to employing the bark of the "fever bark tree" (Cinchona) in the seventeenth 
century and possibly much earlier, in South America, to the use of bednets as a preventive 
method, going back several millennia. The first recorded treatment dates back to 1600, where 
the bitter bark of the cinchona tree in Peru was used by the native Peruvian Indians.
Systematic control o f  malaria started after the discovery o f malaria parasite by Laveran in 
1889 (for which he received the Nobel Prize for medicine in 1907), and the demonstration by 
Ross in 1897 that the mosquito was the vector o f malaria. These discoveries and the 
invention of DDT during the World War II, made the idea o f  global eradication of malaria 
seem possible. Subsequently, widespread systematic control measures such as spraying with 
DDT, coating marshes with paraffin (to block Anopheles mosquito larvae spiracles), draining 
stagnant water, and the widespread use o f nets and cheap, effective drugs such as chloroquine 
were implemented with impressive results.
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Despite initial success, there was a complete failure to eradicate malaria in many countries 
due to a number o f  factors (Shiff, 2002). Although technical difficulties such as vector and 
parasite drug resistance have played a part, the main failure to reduce the disease is probably 
due to social and political factors preventing efficient application o f  control measures. The 
hope of global eradication of malaria was finally abandoned in 1969 when it was recognised 
that this was unlikely ever to be achieved.
2.6.2. Mortality control
The major impact o f malaria in any community is that o f  the death o f individuals and to 
prevent people dying from the disease, appropriate treatment is necessary (Shiff, 2002). This 
strategy involves detecting presumptive cases, determining which cases are parasite positive 
and administering effective treatment. Mortality control is the main thrust o f the current 
“Global Malaria Control Strategy”. The main problem is that chemotherapy alone is not a 
means of controlling malaria and is not sustainable in the long term.
Malaria is diagnosed by the clinical symptoms and microscopic examination o f  the blood. It 
can normally be cured by antimalarial drugs. Antimalarial drugs form an important element 
in control programs in treating cases to remove a source o f infection for feeding mosquitoes 
(Shiff, 2002). Current drugs to treat malaria such as doxycycline, proguanil and primaquine 
attack the liver stage, thus preventing the release o f  parasites into the bloodstream. Others, 
such as chloroquine, quinine, sulfadoxine pyrimethamine (Fansidar) and mefloquine, kill the 
parasite within the red blood cells.
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Antimalarials are also used prophylactically (Shiff, 2002). Chemoprophylaxis for malaria is 
recommended to non-immune persons entering areas endemic for malaria to reduce but not 
totally eliminate the risk of infection. Chemoprophylaxis is also recommended for high-risk 
groups, such as young children and pregnant women and recent immigrants from malaria- 
free areas (Collins et al, 2000).
The problem o f malaria has been exacerbated in recent years by the development and rapid 
spread of resistance in P. falciparum  to the more commonly used and affordable antimalarial 
drugs (Collins et al., 2000; UNICEF 2000). Drug resistance in malaria, has emerged as one 
o f the greatest challenges facing malaria control today (Boland, 2002). It leads to greater 
parasite longevity in the host, which, in turn, prolongs the period o f infectiveness. 
Chloroquine resistance, which first appeared in East Africa in the late 1970s, has now spread 
throughout most o f sub-Saharan Africa, and resistance to pyrimethamine-sulfadoxine 
(Fansidar) has followed rapidly (Figure 2.2). Mefloquine resistance has emerged in South­
east Asia. One of the major implications o f  the diversity o f  resistance is to make it more 
critical that public health measures to control malaria be region-specific.
2.6.3 Transmission control
This strategy recognizes that malaria is an important cause o f morbidity as well as mortality 
and it involves all efforts aimed at controlling the vector o f the disease (Shiff, 2002). This 
approach is effective in most epidemiological conditions and is an effective control strategy 
for a sustained attack on the malaria problem. It is adaptable to the use o f  insecticide-treated 
mosquito nets as well as indoor spraying o f insecticides. Effective transmission control will
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reduce the incidence o f infection and re-infection in the community. When Grassi and his 
Italian colleagues demonstrated that anopheline mosquitoes were the vectors o f  malaria to 
humans, the concept o f malaria control became synonymous with mosquito control and 
mosquitoes became the main target o f control efforts (Robert and Collins, 1996; Shiff, 2002). 
In the absence of methods to kill adult mosquitoes, the strategy was to reduce breeding sites.
2.6.3.1 Environmental management
Environmental management is one o f the earliest malaria control measures practiced even 
before mosquitoes were implicated in malaria transmission. The value o f environmental 
management is well recognized as a form o f source reduction and involves measures to 
reduce breeding sites and overall populations o f vector species. These include the filling o f 
ditches, covering water containers, flushing irrigation channels, clearing ponds o f weed 
growth, which allows the introduction into ponds o f fish which eat mosquito eggs and larvae. 
There have been some situations where source reduction was effective. In the United States, 
major modifications o f  mosquito habitat through the Tennessee Valley Authority malaria 
control program, habitat degradation, deforestation, flooding, and other effects o f 
development restricted the habitat o f  the malaria mosquito Anopheles quadrimaculatus and 
led to the local decline o f  malaria (Shiff, 2002).
However several major environmental changes due to human activities in the quest o f 
development run counter to the source reduction efforts. Mining, logging and land clearance 
for Agriculture are three such operations which can have a rapid impact on the tropical 
environment. Extensive borrow pits which hold water are dug alongside new roads
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constructed for access to these kinds of developments and the new roads often obstruct 
existing drainage, causing water to accumulate and thus new mosquito breeding sites emerge 
(Phillips, 2001).
Moreover, on the whole, anopheline mosquitoes are opportunistic breeders that favour open 
sunlit pools or small streams and rivulets. In most cases it is impractical to suggest source 
reduction as an effective control effort for anophelines. Since anophelines are opportunists, 
their populations expand during rainy spells and they breed in such a variety o f situations that 
any attempts to limit the extent o f suitable habitat will not be very successful. Importantly, it 
is not the number o f  mosquitoes that is critical in the cycle but, rather, the length o f  mosquito 
survival which contributes to the efficient transmission o f malaria (Roberts and Collins, 
1996; Shiff, 2002).
2.6.3.2 Intradomicile application of residual insecticides
Intradomicile application of residual insecticides, also referred to as indoor spraying, has 
been the mainstay of malaria control operations since the early parts of the last century. The 
rationale, in short, is indoor spraying with a persistent insecticide that remains active on the 
sprayed surface for weeks or even months to kill or at least repel the adult female mosquito 
(Shiff, 2002). The motivation for this method is based on the feeding and resting habits o f 
most malaria vectors (Curtis and Towson, 1998). The majority of important malaria vectors 
feed late at night, with peak biting activities between the hours o f 20:00 and 05:00 nightly. 
They are also highly anthropophilic and endophagic. The term endophily refers to the
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preference of a female mosquito to rest indoors during the period between the end o f  feeding 
and the onset o f the search for an oviposition site.
Residual house spraying is likely to be effective only if  the mosquito species concerned is 
endophilic or at least partially endophilic, because the mosquito needs to rest on the 
insecticide-treated walls for a sufficient time if  it is to pick up a lethal dose. Naturally 
endophilic species include An. gambiae s.s. and An. funestus in Africa, An. culicifacies in 
India, and An. minimus in East and Southeast Asia, (Pates and Curtis, 2005). Endophilic 
behavior varies among species and is affected by insecticidal irritancy. The spraying o f the 
walls and ceilings o f houses with residual insecticides such as DDT reduces the survival 
prospects o f indoor resting Anopheles mosquitoes sufficiently to greatly reduce the chance o f 
malaria transmission.
However, behavioral resistance in vectors in some countries has arisen in response to 
prolonged spraying programs (Pates and Curtis, 2005). Exophilic behavior has evolved in 
certain populations exposed to prolonged spraying programs. This can have an impact on a 
control effort and may result from an immediate response to the irritant insecticides (DDT or 
pyrethroids), or it may be a genetic trait evolved under selection from the presence o f 
insecticides in houses. Insecticide irritancy can be demonstrated by a strong stimulation to 
take off and fly, a high proportion o f mosquitoes exiting from a treated house, or both (Pates 
and Curtis, 2005).
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2.6.3.3 Insecticide treated materials
The use o f  ITNs is a new and somewhat revolutionary tool for effective vector control. 
Bednets are an effective method to reduce malaria transmission as they stop more humans 
being infected or infectious humans from transmitting the parasite. The use o f bednets is 
especially important if  the room does not have ceiling and insect screens which cover doors 
and windows to stop mosquitoes from getting in. Before the development o f  insecticide 
treated nets (ITNs) as a new technology in the mid-1980s, people in many countries were 
already using nets, mainly to protect themselves against biting insects and for cultural 
reasons.
It was only recently appreciated that a net treated with insecticide offers much greater 
protection against malaria. Insecticide treated bednets locate a deposit o f  a quick-acting 
insecticide of low human toxicity between a sleeper and host-seeking mosquitoes. Thus a 
chemical barrier is added to the often incomplete physical barrier provided by the net (Miller 
et al., 1991; Hodjati et a l 2003). Not only does the net act as a barrier to prevent mosquitoes 
biting, but also the insecticide repels, inhibits or kills any mosquitoes attracted to feed 
(Mbogo et al., 1996; Mathenge et al., 2001). Thus ITNs provide protection both to 
individuals sleeping under them and to other community members. The effect is so 
significant that use o f ITNs is considered to be one o f the most effective prevention measures 
for malaria (WHO/UNICEF, 2003). The use o f  insecticide treated bed nets (ITNs) for both 
individual and collective protection against malaria has shown potential, reducing childhood 
malaria morbidity by 50% and global mortality by 20-30% in The Gambia, Ghana, and 
Kenya (Alonso et al., 1991; Choi et al., 1995; Binka et al., 1996).
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The insecticides o f choice for bed net impregnation are pyrethroids because o f their high 
efficacy, rapid rate o f  knockdown, strong mosquito excito-repellent properties and low 
mammalian toxicity (Hougard et al., 2003; Corbel et al., 2002; Curtis et al., 2003; Zaim et 
al., 2000). However, the increasing resistance o f malaria vectors to pyrethroids threatens to 
reduce the potency o f this important method o f vector control (Curtis et al., 2003; Corbel et 
al., 2002). Furthermore pyrethroid-treated nets have been reported to be involved in the 
selection for the kdr resistance allele (Fanello et al., 1999).
The World Health Organization (WHO) recommends the large-scale use o f  ITNs to control 
malaria transmission because they offer a good cost-efficiency ratio based on active 
community involvement (Diabate et al., 2002). To be an effective control intervention for the 
malaria vectors, a high coverage is required for ITN to act, through a ‘mass effect’. It is 
therefore easier to expand ITN coverage in areas where there is already a culture o f 
mosquito-net usage and the most suitable areas to be targeted will be those where at least 
20% of households already have at least one net each (Manga, 2002). Once such areas have 
been identified, a local plan for improving coverage and for ensuring that at least 80% o f the 
nets in use are (re)impregnated with insecticide is to be developed.
2.6.3.4 Personal protection
The number o f bites can also be reduced by wearing long sleeved shirts and trousers to 
reduce the amount o f  exposed skin, the use o f insect repellents on clothes and exposed skin 
(especially those containing DEET) and spraying bedrooms with aerosols to kill any 
mosquitoes before sleeping.
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2.6.3.5 Transgenic mosquitoes
Consideration of the potential use of genetically modified organisms (GMOs) is driven by 
the realization o f the enormous human cost o f diseases like malaria and o f  the inadequacy o f 
present control measures (Alphey et al., 2002). GMOs could be used in either o f two ways 
for malaria control. The idea is to generate transgenic mosquitoes that express antiparasitic 
genes in their midgut epithelium, thus rendering them inefficient vectors for the disease. 
Because mosquitoes are obligatory vectors for malaria transmission, the spread o f malaria 
could be curtailed by rendering them incapable o f transmitting parasites (Beier, 1998; Alphey 
et al., 2002; Ito et al., 2002). These strategies target the malaria parasite, rather than the 
mosquito itself, for reduction. An alternative use of genetic engineering for malaria control 
takes a more traditional approach. This involves targeting the mosquito population per se for 
reduction through sterile male release.
However, the release of large numbers o f insects presents other specific challenges: for 
example, the need to release only male mosquitoes so as not to increase the number or nature 
of mosquito bites per person per night. Plasmodium-refractory mosquitoes are being rapidly 
developed for malaria control but will only succeed if  they can successfully compete for 
mates when released into the wild (Okanda et al., 2002). Precopulatory behavioural traits 
maintain genetic population structure in wild mosquito populations and mating barriers have 
foiled previous attempts to control malaria vectors through sterile male release.
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2.6.4 Multilateral malaria research and control programs
Multilateral programs are those activities involving two or more nations that are channeled 
through an international or regional agency (Alilio et al., 2004). Examples of these programs 
include the Multilateral Initiative on Malaria (MIM), the Roll Back Malaria (RBM) 
Partnership, the Global Fund for HIV, Tuberculosis and Malaria (Global Fund), the 
Medicines for Malaria Venture (MMV), and the Malaria Vaccine Initiative (MVI). These 
programs have gained prominence due to their great potential for facilitating important 
discoveries and coordination o f large-scale control actions, which cannot be achieved by a 
single African country working alone (Shiff, 2000).
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2.7 Insecticide Resistance
Resistance is defined as the acquired ability o f an insect population to tolerate doses o f 
insecticide which will kill the majority of individuals in a normal population o f  the same 
species (WHO, 1992). Several years o f  intensive use o f  organic insecticides to control 
arthropod pests and disease vectors has led to the selection of pesticide resistance in some 
species. Resistance to insecticides among mosquitoes that act as vectors o f disease emerged 
more than 25 years ago in Africa, America and Europe (Weill et al., 2003).
Factors that induce resistance are numerous and the mechanism adopted by organism 
depends on the prevailing pressure and on the mode o f action o f  the insecticide in use. 
Intoxication o f arthropod by a pesticide encompasses different levels o f pharmacokinetic 
interaction: penetration o f  barrier tissue, distribution, storage, metabolism in internal tissue, 
and molecular interaction with the ultimate target site. Many chemicals are being used 
against arthropods and there are hundreds o f  examples of resistance, and a number o f 
resistance mechanisms have been identified.
2.7.1 Mechanisms of insecticide resistance
Three main mechanisms of resistance to insecticides occur: reduction o f insecticide 
penetration, increased degradation and modification of the insecticide target (Hemingway et 
al., 2004).
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2.7.1.1 Reduced penetration
Here the composition of the insect's exoskeleton becomes modified in ways that inhibit 
insecticide penetration (Matsumura, 1983). Decreased penetration o f insecticides would 
allow ample time for detoxifying enzymes to metabolize the chemical and therefore would be 
less effective. Reduced cuticular penetration alone usually confers only a low level o f 
resistance, however in combination with other mechanisms, it can potentially result in large 
non-additive increase in resistance (Oppemoorth, 1985). Plapp and Hoyer (1968) reported 
reduced penetration of dieldrin and DDT in a strain o f housefly.
2.7.1.2 Metabolic resistance
Enzyme detoxification, by modifying or increasing endogenous enzymes within the insect, is 
major mechanism of resistance (Chareonviriyaphap et al., 2000). In metabolic resistance the 
metabolic pathways o f the insect become modified in ways that detoxify the insecticide, or 
disallow metabolism of the applied compound into its toxic forms. Metabolic resistance to 
insecticides is mediated by qualitative and quantitative changes in proteins that can often be 
difficult to define precisely at the biochemical level. Three broad enzyme classes are 
involved in insecticide detoxification, the mixed function oxidases (MFO), esterases and 
glutathione S-transferases. Their involvement in resistance is commonly identified by 
increases in the characteristic metabolites they produce. All three classes exist in multiple 
forms within each species and it is often not known whether increased activity arises from 
qualitative or quantitative changes in these enzyme complex. Increased synthesis o f these 
enzymes seem to result from gene amplification.
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1.7.1.2.1 Glutathione S-transferases
jlutathione S-transferases (GSTs) are soluble dimeric proteins that are ubiquitous in nature. 
They are involved in the metabolism, detoxification and excretion o f  a large number of 
aidogenous and exogenous compounds from the cell (Ortelli et al., 2003). GSTs catalyses 
he conjugation o f glutathione with compounds having a reactive electrophilic centre, leading 
o the formulation o f a water-soluble, less reactive product. Although there are many 
examples of increased metabolism o f insecticide or model substrates by glutathione S- 
ransferases of resistant insects, few are characterized at the molecular level.
3STs have no direct role in the metabolism of pyrethroid insecticides but they play a very 
mportant role in conferring resistance to this insecticide class by detoxifying lipid 
Deroxidation products induced by pyrethroids (Hemingway et al., 2004). GSTs may also 
Drotect against pyrethroid toxicity in insects by sequestering the insecticide. Insect GSTs are 
}f particular interest because o f their potential to cause resistance to all the major families of 
insecticide. Metabolism mediated by these enzymes has been implicated in DDT and 
Drganophosphate resistance. Increased levels of DDT-dehydrochlorinase have been reported 
in different species resistant to DDT. Biochemical studies on partially purified GST fractions 
from DDT-resistant and susceptible A. gambiae have indicated that resistance was associated 
with both qualitative and quantitative changes in multiple GST enzymes (Ortelli et al., 2003)
2.7.1.2.2 Mixed function oxidases (MFO)
MFO enzymes are o f a great significant both in mammals and arthropods in giving protection 
to a variety of insecticides, particularly to some chlorinated hydrocarbons, to many
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organophosphates and carbamates and to some pyrethroids. Monooxygenases are a chain o f 
enzymes, with the rate limiting enzyme usually being cytochrome P450. Alterations in this 
rate-limiting enzyme can dictate levels o f  resistance to pyrethroids, organophosphates, and 
carbamate insecticides using this metabolic mechanism (Chareonviriyaphap et al., 2003)
An increase in MFO activity is one o f the most versatile mechanisms o f resistance in insects. 
Insect P-450 enzymes also activate certain types o f  insecticides, for instance the conversion 
of phosphorothioates (P=S) to phosphate (P=0). This can result in an increase in potency for 
inhibition of acetylcholinesterase by 3 or 4 orders of magnitude. Monooxygenases can 
contribute to Malathion resistance in two ways, by either increasing the rate o f metabolism to 
non-toxic products, or decreasing the rate at which the insecticidal malaoxon is produced 
from the malathion parent compound (Karunaratne and Hemingway, 2001).
Elevated monooxygenase activity is associated with pyrethroid resistance in An. stephensi, 
An. subpictus, An. gambiae and C. quinquefasciatus (Hemingway and Ranson, 2000). 
Elevated levels o f mixed function oxidases were found to be responsible for the 
detoxification o f pyrethroids in resistant Anopheles funestus Giles from northern 
Kwazulu/Natal in South Africa and the Beluluane region o f  southern Mozambique (Brooke 
et al., 2001) and were further implicated to be conferring cross-resistance to the carbamate 
insecticide propoxur. Similarly, monooxygenases have been responsible for degradation o f 
pyrethroids in Anopheles pseudopunctipennis (Ocampo et al., 2000).
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2.7.1.2.3. Esterases and Hydrolases
Esterases are the most significant enzymes for insecticide detoxification in insects. 
Organophosphate, carbamate and pyrethroids contain carboxylester and phosphotriester 
bonds that are subject to attack by esterase enzymes. These esterases can often be separated 
into isozymes with different substrate specificities (Chareonviriyaphap et al., 2000). 
Polymorphism is a notable characteristic of insect esterases. Esterases are widely distributed 
in many insect tissues such as gut, cuticle and fat body and multiple forms o f  esterases are 
present in the soluble, cytosolic fraction o f insect. Few o f the multiple forms o f  esterase 
isozymes that exist in insects participate in insecticide metabolism, where each isozyme 
probably has a certain range of substrates. Unlike the monooxygenase reaction, esterases do 
not utilize high energy co-factors.
Elevated esterase activity has been linked to pyrethroid, organophosphate and carbamate 
resistance patterns in a variety of insects. Pyrethroids are insecticidal esters derived from 
primary alcohols and are thus susceptible to hydrolysis by esterases. Chareonviriyaphap et 
al., (2000), have reported highly elevated esterase levels in Anopheles albimanus resistant to 
deltamethrin in Guatemala and they further suggest this may limit pyrethroid use against An. 
albimanus population in parts o f Central America.
Different types o f esterases (A l, B l, A2, B2) have been recognized in organophospates 
insecticide resistant populations of Culex pipiens complex throughout the world and 
overproduction o f  nonspecific esterases is a common mechanism o f resistance. For esterase 
B l, resistance to OP insecticides has been shown to be due to sequestration o f insecticide and
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overproduction o f all esterase B is due to gene amplification. Enzymatic assays suggested 
that sequestration rather than metabolism is the primary mode o f operation o f these esterases 
on malathion.
The basis o f malathion resistance in the adults o f  An. arabiansis from Sudan was a 
carboxylesterase. Malathion resistance due to an increase in degradation at the carboxylester 
linkage is a common detoxification pathway that has been implicated in An. culicifacies; 
An.stephensi Liston. Esterase dependent cross- resistance between Organophosphates, 
carbamate and pyrethroids has been detected in several insect species. An. gambiae from 
Kenya has been reported to have demonstrated elevated oxidase and esterase levels in 
permethrin-resistant (Vulule et al., 1999). Similarly, Brogdon et al. (1999a, b) have reported 
oxidase-based and esterase-based resistance mechanisms alone and in combination in 
permethrin-resistant An. albimanus from Guatemala.
2.7.1.3 Target-site mechanisms
Alterations of amino acids responsible for insecticide binding at its site o f  action cause the 
insecticide to be less effective or even ineffective. Non-silent point mutations within 
structural genes Eire the most common cause o f target-site resistance (Hemingway and 
Ranson, 2000). For selection o f the mutations to occur, the resultant amino acid change must 
reduce the binding o f  the insecticide without causing a loss o f primary function o f the target 
site. Therefore the number o f possible amino acid substitutions is very limited. Hence, 
identical resistance-associated mutations are commonly found across highly diverged taxa. 
The degree to which function is impaired by the resistance mutation is reflected in the fitness
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of resistant individuals in the absence of insecticide selection. This fitness cost has important 
implications for the persistence of resistance in the field.
2.7.1.3.1 Knock-down resistance
The target o f organochlorines especially DDT and synthetic pyrethroids are the sodium 
channels of the nerve sheath (Hemingway et al., 2004). In insects, an important mechanism 
of pyrethroid resistance is due to a modification of the voltage-gated sodium channel protein 
recently shown to be associated with mutations o f the para-type sodium channel gene, 
(Martinez-Torres et al., 1988). A reduction in the sensitivity o f the insect’s voltage-gated 
sodium channels to the binding of insecticides causes the resistance phenotype known as 
knockdown resistance (kdr) (Hemingway and Ranson, 2000). DDT-pyrethroid cross­
resistance may be produced by single amino acid changes in the axonal sodium channel 
insecticide-binding site.
Pyrethroid insecticides have a rapid ‘knock-down’ effect. However, the intensive use of DDT 
and pyrethroids has led to the development of kdr in many insect species including 
Anopheles mosquitoes (Hemingway et al., 2004). The use o f  both DDT and pyrethroids in 
the control o f rice and cotton pests is likely to have contributed significantly to the 
development o f resistance in An. gambiae from West Africa (Hemingway et al., 2004; 
Martinez-Torres et al., 1998). Already Pyrethroid resistance has been noted in An. 
albimanus, An. stephensi, and An. gambiae among the malaria vectors.
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The development o f pyrethroid resistance in An. gambiae is particularly important given the 
recent emphasis by the WHO and other organizations on the use of pyrethroid-impregnated 
bednets for malaria control. Two cases o f pyrethroid resistance in An. gambiae, from the 
Ivory Coast (Chandre et al., 1999) and Kenya (Vulule et al., 1999) are well documented. 
However, in Cameroun, Senegal and Botswana, An. gambiae populations have been reported 
to be fully susceptible to pyrethroids (Chandre et al., 1999a). The West African focus 
appears to be larger and has higher levels o f resistance than that in East Africa (Hemingway 
and Ranson, 2000). Resistance due to kdr can develop in a short period o f introduction of 
pyrethroids for mosquito control. Approximately one year after bed nets impregnated with 
permethrin were introduced as a malaria control measure in the northern part o f Thailand, 
evidence o f physiological resistance was reported (Chareonviriyaphap et al., 2002).
Several studies have reported that, the kdr mutation has been found widely distributed in the 
Savanna form o f An. gambiae s.s., but never in wild populations o f the Mopti form or An. 
arabiensis, even in areas where both occur in sympatry with resistant Savanna populations 
(Chandre et al., 1999; Awolola et al., 2003; Berzosa et al 2002). As a result it was suggested 
that, the absence o f  the kdr mutation in the M form involves an additional pyrethroid 
resistance mechanism in An. gambiae s.s. (Awolola et al., 2003). However, already low 
levels o f the kdr mutation has recently been detected in An. arabiensis during an extensive 
survey of pyrethroid resistance in An. gambiae s.l. in Burkina Faso (Diabate et al., 2004), 
Ethiopia (Balkew et al., 2003) and Western Kenya (Stump et al., 2004). Similarly, Weill et 
al., (2000) has reported kdr mutation in the Mopti form o f An.gambiae s.s. The detection o f 
this mutation in both An. arabiensis and the M form o f An. gambiae s.s., is important at both 
epidemiologic and fundamental levels. Most susceptibility/resistance studies in Ghana have
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reported insensitivity of the sodium channels in An. gambiae populations, as characterized by 
increased knockdown and high frequency o f the kdr mutation allele (Adasi et al., 2000: 
Adeniran, 2002: Otieno, 2004).
2.7.1.3.2 Insensitive acetylcholinesterase
The target o f organophosphate and carbamate insecticides is acetylcholinesterase (AcChE) in 
nerve synapses. AcChE is a key enzyme in the cholinergic synapses where it rapidly 
terminates nerve impulses by catalyzing the hydrolysis o f the neurotransmitter acetylcholine. 
Organophosphates and carbamates are substrates o f AcChE and their hydrolysis results in the 
phosphorylation or carbamylation of the active serine followed by dephosphorylation or 
decarbamylation. The deacetylation of the acetylated enzyme by its natural substrate 
acetylcholine is a rapid process, 1000 s '1 in insects. However, dephosphorylation or 
decarbamylation is very long and takes several days while synaptic transmission remains 
blocked, resulting to the death o f the insect, (Hemingway et al., 2004: Shi et al., 2004).
This resistance is frequently due to a loss o f sensitivity o f  the insect's acetylcholinesterase 
enzyme to organophosphates and carbamates. This insensitivity results from a single amino- 
acid substitution in the enzyme (Weill et al., 2003). At least five point mutations in the 
acetylcholinesterase insecticide-binding site have been identified that singly or in concert 
cause varying degrees o f reduced sensitivity to organophosphates and carbamate insecticides. 
Already resistance to carbosulfan, a carbamate insecticide, due to insensitive 
acetylcholinesterase has been detected in field populations o f Anopheles gambiae in Ivory 
Coast (N'Guessan et al., 2003; Corbel et al., 2003).
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2.7.1.3.3 GABA receptor changes
The Y-Aminobutyric acid (GABA) is the major inhibitory neurotransmitter in both insects 
and vertebrates (Ffrench-Constant et al., 2000). The GABA receptors belong to a 
superfamily of neurotransmitter receptors that also includes the nicotinic acetylcholine 
receptors (Hemingway and Ranson, 2000). These receptors are formed by the 
oligomerization o f five subunits around a central transmitter-gated ion channel. An alanine- 
to-serine substitution in the putative channel-lining domain o f the GABA receptor confers 
resistance to cyclodienes such as dieldrin (Hemingway and Ranson, 2000). Resistance to 
dieldrin was recorded in the 1950s, but the involvement of the GABA receptors in this 
resistance was not elucidated until the 1990s.
Although cyclodiene resistance is historically very widespread and in the past accounted for 
over 60% o f reported cases o f resistance, cyclodienes themselves have been largely with­
drawn from use, and therefore, in relative terms, the overall frequency o f  resistance cases is 
declining (Ffrench-Constant et al., 2000). However, resistance seems to be able to persist in 
the absence of extensive insecticide selection, representing a threat for novel insecticides 
interacting with the cyclodiene binding site, such as the fipronils. Further, cyclodiene type 
insecticides, such as endosulfan, are still used to control multiple resistant pests including the 
whitefly.
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2.8 M anagement of Insecticide Resistance
Insecticide resistance has been a problem in all insect groups that serve as vectors o f 
emerging diseases. Although mechanisms by which insecticides become less effective are 
similar across all vector taxa, each resistance problem is potentially unique and may involve 
a complex pattern of resistance foci. The main defense against resistance is close surveillance 
of the susceptibility of vector populations.
In the present situation of insecticide resistance status in malaria vectors, the future o f vector 
control mainly relies on the strategies for the management of insecticide resistance. So far the 
approach has been the replacement of insecticide by an effective and preferably by a new 
group of insecticides. Subsequent replacement of insecticides has led to the development o f 
multiple-resistant malaria vectors. It may be mentioned that subsequent change o f 
insecticides has burdened the programme with increased costs. Not many new insecticide 
molecules are available for vector control in the immediate future. What is needed for the 
present day vector control programme is an approach for the management o f  existing 
resistance in malaria vectors and to limit its further spread. The strategy for this approach is 
to use the available insecticides rationally.
The most important aspect o f the management o f  resistance is to either avoid or delay the 
onset o f resistance by effectively manipulating or influencing the factors responsible for the 
development o f resistance (Hemingway et al., 2004). Various methods emphasize on the 
strategic use o f  available insecticides to delay the onset o f  resistance. The methods include 
avoidance o f use o f insecticides that induce broad-spectrum resistance mechanisms and
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2.8 Management of Insecticide Resistance
Insecticide resistance has been a problem in all insect groups that serve as vectors o f 
emerging diseases. Although mechanisms by which insecticides become less effective are 
similar across all vector taxa, each resistance problem is potentially unique and may involve 
a complex pattern o f resistance foci. The main defense against resistance is close surveillance 
of the susceptibility of vector populations.
In the present situation of insecticide resistance status in malaria vectors, the future o f vector 
control mainly relies on the strategies for the management of insecticide resistance. So far the 
approach has been the replacement of insecticide by an effective and preferably by a new 
group of insecticides. Subsequent replacement o f insecticides has led to the development o f 
multiple-resistant malaria vectors. It may be mentioned that subsequent change o f 
insecticides has burdened the programme with increased costs. Not many new insecticide 
molecules are available for vector control in the immediate future. What is needed for the 
present day vector control programme is an approach for the management o f existing 
resistance in malaria vectors and to limit its further spread. The strategy for this approach is 
to use the available insecticides rationally.
The most important aspect o f the management o f  resistance is to either avoid or delay the 
onset o f resistance by effectively manipulating or influencing the factors responsible for the 
development o f resistance (Hemingway et al., 2004). Various methods emphasize on the 
strategic use of available insecticides to delay the onset o f resistance. The methods include 
avoidance of use of insecticides that induce broad-spectrum resistance mechanisms and
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confer cross-resistance to chemically related and unrelated insecticides. The sequential use of 
insecticides in rotation is preferred. The need of the hour is intensive research on 
management tactics and integration of such tested strategies in the ongoing vector control 
programmes. In malaria endemic areas, there is a need for comparative studies on susceptible 
and refractory populations for as many known vectors as possible (Chareonviriyaphap et al., 
2002).
Among the strategies proposed for resistance management is the use a pyrethroid and a non- 
pyrethroid insecticide in combination on the same mosquito net, either separately or as a 
mixture (Darriet et al., 2003; Corbel et al., 2003). Mixtures are particularly promising if  there 
is potentiation between the two insecticides as this would make it possible to lower the 
dosage o f each hence an advantage in terms o f  lower cost and toxicity (Hougard et al., 2003). 
The possible use o f non-pyrethroid insecticides, such as carbamates, on nets is a promising 
alternative solution because these insecticides are effective against susceptible and 
pyrethroid-resistant populations o f Anopheles and Culex mosquitoes (Corbel et al., 2003).
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confer cross-resistance to chemically related and unrelated insecticides. The sequential use o f 
insecticides in rotation is preferred. The need o f the hour is intensive research on 
management tactics and integration of such tested strategies in the ongoing vector control 
programmes. In malaria endemic areas, there is a need for comparative studies on susceptible 
and refractory populations for as many known vectors as possible (Chareonviriyaphap et al., 
2002).
Among the strategies proposed for resistance management is the use a pyrethroid and a non- 
pyrethroid insecticide in combination on the same mosquito net, either separately or as a 
mixture (Darriet et al., 2003; Corbel et al., 2003). Mixtures are particularly promising i f  there 
is potentiation between the two insecticides as this would make it possible to lower the 
dosage of each hence an advantage in terms o f  lower cost and toxicity (Hougard et al., 2003). 
The possible use o f non-pyrethroid insecticides, such as carbamates, on nets is a promising 
alternative solution because these insecticides are effective against susceptible and 
pyrethroid-resistant populations o f Anopheles and Culex mosquitoes (Corbel et al., 2003).
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CHAPTER THREE
M ATERIALS AND M ETHODS
3.0 Standard solutions
All solutions and reagents used were prepared according to standard procedures as shown in 
Appendix I.
3.1 Study area
The studies were carried out in three suburb villages o f Hohoe town; Adabraka (N 07° 
09.881’ E 000° 28.514’), Atabu Newtown (N 07° 08.153’ E 000° 28.815’) and Kledzo (N 07° 
07.353’ E 000° 27.523’) and Likpe (N 07° 09.799’ E 000° 35.313’), a rural village lying on 
the foot o f mountain ranges bordering Ghana and Togo.
Hohoe District is one of the twelve administrative districts o f the Volta-Region and has a 
land surface area o f 1,172km2. It is located within longitude 0°15E and 0° 45E and latitude 6° 
45’N and 7° 15’N and lies almost at the heart o f the Volta-Region. It shares borders with the 
Republic o f Togo on the east, on the southeast with Ho District, on the southwest with 
Kpando District and on north with Jasikan District. The District capital Hohoe is 220km 
from Accra, the national capital.
The District houses part o f the Akwapim-Togo ranges extending beyond the country’s 
eastern boundary all the way to Western Nigeria. Within these ranges is the Afadjato -  the 
highest elevation in Ghana (880.3m). Some of the low-lying areas have swamps which are
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used for rice cultivation. Annual rainfall is between 1016m m-121 Omm. There is 4-5 month 
dry season between November and April. Hohoe District falls within the forest-savanah 
transitioned ecological zone o f Ghana with the forest part at its southern and eastern sector 
and tapering into the middle o f the district. The eastern highlands are clothed with high 
forest. The soils generally tend to be sandy overlying iron pans. Bottom lands carry heavy 
silts and cracking clays and as a consequence, drainage is very poor, subjecting the area to 
extreme variations in soil moisture. Both savannah and forest crops do well in the district. 
Some o f these are cocoa, coffee, oil palm, bananas and plantains, rice, cassava, yams, maize, 
millet and groundnuts. The year 2000 population figure for Hohoe district (based on DSDA
II headcount) is 144,511 with the gender breakdown established as 70,754 males and 73,547 
females.
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3.2 Field collection of mosquitoes
Mosquitoes were collected as adults and larvae in the above mentioned sites between August 
2004 and April 2005.
3.2.1 Adult mosquito collection
In each village, houses (Figure 3.1 and 3.2) were randomly selected for adult mosquito 
collection in rooms where people sleep. The rooms of selected houses were thoroughly 
searched during the day for indoor resting Anopheles mosquitoes with the aid o f a torch light. 
Adult anophelines were identified by their characteristic resting position, with the body 
resting at an angle o f 45° to the surface. Only bloodfed and gravid adult females were 
collected using plastic aspirators and carefully transferred into paper cups by gently blowing 
them out of the aspirator. They were then transported to the laboratory where they were 
maintained until they were ready to lay eggs.
3.2.2 Larval mosquito collection
Where the number o f adults was very low, mosquitoes were collected as larvae from 
randomly selected larval sites around the sampled houses. The larval sites comprised mainly 
of abandoned fish ponds, tyre tracks, shallow water wells and an open concrete water tank 
(Figure 3.3 and 3.4). The Anopheles larvae were identified by their characteristic resting 
position, with the body lying parallel to the surface and just below the surface film. Larvae 
were collected using the dipping method with the aid o f copper ladles, transferred into plastic 
jars which had perforated covers for ventilation and transported to the laboratory for rearing.
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Figure 3.1: Typical adult mosquito collection house at Adabraka village.
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Figure 3.4: Typical larval mosquito collection site at Atabu Newtown village. A 
shallowly-dug water well.
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Figure 3.3: Typical larval mosquito collection site at Adabraka village. An abandoned 
fish pond filled with rain water.
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3.2.3 Laboratory rearing of mosquitoes
In the laboratory gravid females were carefully transferred into 1 foot square cages and 
provided with 10% sugar solution soaked in cotton wool. Petri dishes lined with moist filter 
paper were placed at the base of the cages for them to lay eggs on. The eggs when laid were 
then carefully transferred into plastic trays containing water for them to hatch. Larvae were 
later reared in plastic containers of 5cm x 27cm x 36cm that contained water to the depth of 
not more than 2cm (Figure 3.5 a). Field collected larvae were also transferred into similar 
trays and in cases where the water from their natural habitats was unsuitable, tap water was 
used. Larvae were fed on a mixture of two parts finely ground gold fish food (Nutrifin™, 
Warden Corporation U.S.A) and one part brewer’s yeast (Saf-instant , France) in water. 
Pupae were collected each day using Pasteur pipettes and transferred into small beakers and 
placed in labelled cages (Figure 3.5 b) for adult emergence. The emerging adults were 
similarly fed on 10% sugar solution soaked in cotton wool. Only 2-5 days old females were 
picked from the cages and used for bioassays.
The temperature and relative humidity during the rearing period was within the range 25- 
30°C and 55-78% respectively and a natural photoperiod was maintained. The cages were 
mounted on oil moats to prevent ants from entering the cages. Dead mosquitoes were also 
picked up daily to prevent attraction o f ants and mould formation. Other precautions such as 
overcrowding of larvae in the tray and not using much feed to avoid scum formation were 
taken during rearing.
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b)
Figure 3.5: Mosquito rearing system; a) Plastic pans for rearing larvae 
b) Wooden cages for rearing adults.
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3.2.4 Morphological identification of Anopheles gambiae s.l. mosquitoes
The Anopheles mosquitoes were identified using the keys by Giles and de Mellion (1968) 
and Hervy et al. (1998). The adults usually rest with the body at an angle to the surface and 
most of them have spotted wings. The number, length and arrangement o f the spots differ 
considerably in different species. The females have non-plumose antennae and palps as long 
as the proboscis and usually lie closely alongside (Service, 1980) while the males have 
plumose antennae. Apart from the morphological identification o f Anopheles from other 
mosquitoes, Morphological identifications were carried out to separate culex and Aedes 
species and to distinguish An. funestus from An. gambiae s.l. Anopheles gambiae complex 
species have 5 pale spots on the coastal margin o f  the wings, anal vein colouration with 3 
white spots and a dark apical fringe and white speckle (or spots in the median part) tibia 
ornamentation. In contrast, An.funestus and other Anopheles species have 4 pale spots on the 
costal margin on the wings, entirely dark anal vein colouration and entirely dark tibia 
ornamentation.
3.2.5 Preservation of mosquito samples
Randomly selected mosquitoes were placed in paper cups and killed by freeing at -20° C for 
at least 10 minutes. Specimens for identification of species and molecular forms and for Kdr 
analysis using molecular methods were preserved dry over silica gel in individual tubes. 
Specimens for biochemical analysis were quickly transferred to tubes and stored at -80° C 
until ready to use.
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3.3 Bioassay Experiments
The aim of the bioassay was to measure the time it took for a given insecticide to kill the 
adult mosquitoes. The susceptibility tests were carried out using the World Health 
Organization test kits for adult mosquitoes (WHO, 1998b). The kit is basically comprised of 
insecticide impregnated test papers and non-impregnated papers for control and plastic tubes 
that are marked red for exposure and marked green for holding (Figure 3.6). The papers were 
impregnated with the WHO-recommended discriminating dosages of 5% Malathion, 0.1% 
Propoxur, 0.05% Deltamethrin, 0.75% Permethrin and 4% DDT. For each test, batches o f  25 
female mosquitoes aged between 2-5 days old were aspirated from the cages and transferred 
into paper cups where they were held for 1 hour. They were then aspirated into exposure 
tubes lined with the insecticide impregnated papers for 1 hour; during which the number of 
mosquitoes knocked down was recorded after 10, 15, 20, 30, 40, 50 and 60 minutes. Where 
the number knocked down after 60 minutes was less than 80%, number knocked down was 
recorded for 20 more minutes (i.e after 80minutes) in the holding tube. A mosquito was 
considered knocked down if  it lay on its side on the floor o f  the exposure tube and was 
unable to fly (WHO, 1998b).
Mosquitoes were then transferred into holding tubes by gently blowing them through the 
open space between the exposure and the holding tubes. Cotton soaked in 10% sucrose was 
placed on top o f  the holding tube. This was to avoid death by starvation. The mortality was 
scored after 24 hours post-exposure and each test in each site was replicated four times. None 
of the impregnated papers was used for more than four times. The resistance or susceptibility 
status was evaluated based on the WHO criteria i.e. 98-100% mortality indicated
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susceptibility; 80-97% mortality required confirmation and less than 80% mortality indicated 
resistance (WHO, 1981; WHO, 1998b). When the control mortality was between 5% and 
20% the mean observed mortality was corrected using Abbott’s formula (Abbott, 1925). An 
experiment was repeated if control mortality was more than 20%. The susceptibility o f  the 
wild populations to the tested insecticides was compared with that o f the susceptible Kisumu 
strain.
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b)
Figure 3.6: Bioassay experiment set set-up; a) Exposure tubes with 
insecticide impregnated papers (b) Holding tubes.
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3.4 Molecular Studies
Molecular methods were used to identify species o f Anopheles gambiae complex, molecular 
forms of An. gambiae s.s and the Kdr alleles.
3.4.1 DNA Extraction
Genomic DNA extraction was carried out using two methods: the Bender buffer method (a 
modified protocol o f Collins et al., 1987) and homogenised legs/ parts o f mosquito. For the 
Bender buffer extraction, each mosquito was homogenised in 1.5ml Eppendorf tube 
containing 100/d of the buffer (preheated at 65°C) using a sterile plastic pestle followed by 
incubation at 65°C for 30 minutes. Then 125 fil o f  phenol was added to the homogenate, 
mixed well by vortexing and spun at 14,000 rpm for 10 minutes. The supernatant was 
transferred into a fresh tube and 250 /d o f chloroform added, vortexed briefly and spun as 
described. The supernatant was again transferred into a fresh tube and 250/nl o f  pre-chilled 
absolute ethanol and 10 /d o f  8M potassium acetate added followed by incubation at -40°C 
for 1 hour or -20°C overnight. The DNA was pelleted by centrifugation at 10000 rpm for 10 
min and the supernatant discarded. Two hundred micro litres o f 70% ethanol added to the 
pellet, the tube gently swirled and the DNA repelleted by centrifugation at 10000 rpm for 5 
minutes. The supernatant was discarded and the tube inverted over a tissue paper to dry. The 
DNA pellet left was re-dissolved in 25/d TE plus RNase (5|ig/ml). It was then stored at -20°C 
until ready for use. This method was used mainly for the kdr analysis because the PCR with 
the DNA extracted from mosquito legs proved unsuccessful.
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The ground leg extract involved grinding a single mosquito leg or wing with a plastic pestle 
in 50 /zl of sterile double distilled water (sdd H20 )  in 1,5ml Eppendoff tube. The homogenate 
was boiled for lOminutes, spun briefly and used directly as a DNA template for PCR.
3.4.2 PCR identification o f Anopheles gambiae complex
Anopheles gambiae sibling species identification was carried out according to the method o f 
Scott et al. (1993). Five sets o f primers abbreviated as UN, GA, ME, AR and QD designed 
from the DNA sequence o f the intergenic spacer region o f An. gambiae complex o f 
ribosomal DNA (rDNA) were used for species identification (Table 1). The sequence details 
of these primers, expected sizes o f the PCR products and their melting temperatures are also 
given in Table 2. The UN primer anneals to the same position on the rDNA sequences o f all 
five species, GA anneals specifically to An. gambiae s.s., ME anneals to both An. merus and 
An. melas, AR to An .arabiensis and QD to An. quadriannualatus.
The PCR reaction mixture o f  20 /*1 contained lx  reaction buffer (Buffer C), 200/iM each of 
the four oligonucleotide triphosphate (dNTPs), 0.25 /xM each o f oligonucleotide primers and 
0.5U of DNA Taq polymerase enzyme. Five microliters o f mosquito DNA (from single 
ground leg extraction method) template was used as the template for the amplification 
reaction. The reaction mixture was made up to 20 fi\ with sterile double distilled water.
The reaction mixture was spun down briefly and overlaid with mineral oil to avoid 
evaporation and refluxing during thermo cycling. The PCR thermal cycling was as follows; 
an initial step o f 3 min at 94°C, followed by 35 cycles with denaturation at 94°C for30s, 
annealing at 50°C for 30s and extension at 72 °C for 60s and ended with a final cycle at 94°C
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for 30s, annealing at 50 °C for 30s and extension at 72°C for lOmins using a PCR Express 
Thermal Cycler (Hybaid Ltd., UK).
The amplified products were analysed by agarose gel electrophoresis. Ten microliters o f each 
PCR product were added to 1/il o f lOx Orange G loading dye and electrophoresed in 2% 
agarose gel stained with 0.5/tg/ml o f ethidium bromide. The electrophoresis was run in IX  
Tris acetate-EDTA (TAE) buffer at 100V for one hour and were visualized and photographed 
over a UVP dual intensity transilluminator at short wavelength using a Palorid direct screen 
instant camera fitted with an orange filter, a hood and a Polaroid Type 667 film. The film 
was processed as recommended by the manufacturer (Polaroid Inc., USA). The amplified 
PCR product was identified to the sibling species on the basis o f the diagnostic band size 
determined by comparison with the mobility o f a standard lOObp DNA ladder (Sigma, USA).
3.4.4 Identification of the molecular forms of Anopheles gambiae s.s 
The identification o f An. gambiae s.s to the molecular forms was done using polymerase 
chain reaction-restriction fragment length polymorphism (PCR-RFLP). The method of 
Fanello et al. (2002) involving a combination o f the protocols established by Scott et al. 
(1993) and Favia et al. (1997) was used. This method allows for simultaneous identification 
of An. gambiae s.l. as well as the M and S forms within the An. gambiae s.s. It is based on 
the fact that GCG* C restriction site for Hhal enzyme (Favia et al., 1997) lies within the An. 
gambiae specific fragment (Scott et al.,1993) which makes it possible to digest this fragment 
directly in order to differentiate M and S molecular forms.
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The PCR reaction is the same as described in Section 5.4.2. After amplification, 1U o f Hhal 
enzyme (Promega, USA) in 10X enzyme buffer (Promega, USA) and nuclease free water 
were added to lO^il o f PCR product to make a 20|il o f the reaction mixture. The digestion 
was carried out at 37°C for 6 hours in a thermal cycler and the products electrophoresed 
through ethidium bromide-stained 2% agarose gel and visualised under UV light. Different 
band sizes are obtained between the PCR-amplified fragments and the fragments obtained 
after digestions are due to the presence o f a restriction site for Hhal (Fanello et al., 2002) at 
position 469 in all taxa except An. merus, and o f a second restriction site at position 475 in 
An. quadriannulatus, An. melas and An. merus. The An. gambiae S-form digestion is 
characterized by two fragments, 257 and 1 lObp long, resulting from the presence o f the Hhal 
restriction site. The An. gambiae M-form does not have this restriction site and thus is 
characterized by a single 367bp fragment.
3.4.5 PCR detection of the kdr alleles in Anopheles gambiae complex 
The PCR-based method o f Martinez-Torres et al. (1998) was used to detect kdr genes in the 
mosquitoes. DNA extraction was performed as described in Section 3.5.1. The primers used 
were Agdl and Agd2 (Oligos Etc. Inc., USA) and Agd3 and Agd4 (Oswel, UK) [Table 2], A 
total of 40 survivors from the bioassay plus 20 individuals o f the general population o f each 
of the sites were chosen at random for the kdr analysis. Kdr genotyping o f susceptible and 
resistant individuals was possible after amplifying the DNA template from mosquitoes 
following the PCR conditions o f  94°C for 3mins (initial denaturation), followed by 45cycles 
of 94°C for 30sec, 50°C for 30sec and 72°C for lmin. There was a final extension cycle of 
94°C for 30sec, 50°C for 30 sec and 72°C for lOmin followed by 4°C for cooling.
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The products were electrophoresed through ethidium bromide-stained 2% agarose gel and 
visualized under UV light. Kdr genotypes o f both the susceptible and resistant individuals 
were then recorded. Expected sizes for susceptible, resistant and control were 137bp, 195bp 
and 293bp respectively. The positive controls were laboratory susceptible strains o f An. 
gambiae from Kenya (Kisumu).
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Table 1: Oligonucleotides primer sequences, melting temperatures and the expected band 
sizes o f the PCR amplified DNA products for identification o f An. gambiae species complex 
(Scott etal., 1993).
Primer Sequence(5"-3") TM(”C) Band size (bp)
UN GTG TGC CCC TTC CTC GAT GT 56 468
GA CTC GTT TGG TCG GCA CGT TT 62 390
ME TGA CCA ACC CAC TCCCTT GA 90 464
AR AAG TGT CCT TCT CCATCCTA 78 315
QD CAG ACC AAG ATG GTTAGT AT 54 153
Table 2: Sequence details o f the kdr primers and their melting temperatures
Primer Sequence (5"-3") TM(°C)
A g d l ATA GAT TCC CCG ACC ATG 54
Agd 2 AGA CAA GGA TGA TGA ACC 64
Agd 3 AAT TTG CAT TAC TTA CGA CA 40
Agd 4 CTG TAG TGA TAG GAA ATT TA 52
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3.5 Biochemical assays
The purpose of conducting these assays was to determine activity o f  detoxification enzymes 
in the field populations as compared to that of a susceptible ‘Kisumu’ strain. The Centre for 
Disease Control (CDC, USA) microplates assays protocol with minor modifications was used 
(CDC, 2002). The enzymes tested were oxidases, glutathione S-transferases, 
acetylcholinesterase and non-specific esterases.
3.5.1 Sample preparation
Batches of 30 randomly selected frozen mosquitoes were placed individually in 1.5ml 
Eppendorf tubes using fine forceps and 100|il o f phosphate buffer (pH 7.2) added. 
Mosquitoes were then thoroughly homogenized using either Teflon or plastic pestles and the 
crude homogenate, diluted to a final volume of 1ml with additional 900|il of the buffer.
For each individual mosquito, lOOjxl each of the homogenate was loaded into flat-bottomed 
microplate wells in triplicate (Figure 3.7). For an enzyme assay, homogenates o f the first 
mosquito were loaded into the first three wells across i.e. A 1-3. The homogenate of next 
mosquito was loaded in the wells directly below the first i.e. B 1-3 and this continued down 
the plate until the end, then moving to the right and beginning at the top again. Wells A 4-6 
contained the ninth mosquito. The last 6 lower wells on the plate were loaded with the 
positive and negative controls. Fresh pipettor tip was used for each homogenized mosquito 
sample to avoid cross-contamination.
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1 2 3 4  5 6 7 8 9 10 11 12
a 0Q©000©000U© 
B @0OO©@©@©@©© 
C © © © © © © © © © © O O  
D ©OO©©©©©©©©©
E
F • • • • • • • • # § •
G o o o o o o o o o f • • • POS CTRL
H NEG CTRL
Figure 3.7: An illustration of a flat-bottomed microtiter plate used for biochemical assays.
Homogenates were loaded in the following order; Mosquito # 1 went into wells 
A l, A2, A3, mosquito # 2 went into wells B l, B2, B3, mosquito # 8 went into 
wells H I, H2, H3 and so on.
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3.5.2 Calibration curves
Calibration curves were obtained for protein, oxidases and the non-specific esterases.
3.5.2.1 Protein
Bovine serum albumin (BSA) was used to obtain a calibration curve for protein by dissolving 
0.02g o f BSA in 10ml 0.1M potassium phosphate buffer. Then serial dilutions o f the stock 
solution were made using the same buffer to obtain concentrations within the range 0.25-2.0 
mg/ml. For each dilution 5jj.1 were loaded into wells in triplicates and 250j_il o f Bradford 
reagent (protein dye concentrate) added. The plate was incubated and read after 5 minutes 
using an ELISA plate reader at 620nm. The buffer was used as the negative control. 
Concentrations were plotted against absorbance to obtain a calibration curve which was then 
used to determine the corresponding protein content in mosquito samples.
3.5.2.2 Oxidase
Cytochrome-C was used to obtain a calibration curve for oxidase. 80mM solution of 
cytochrome -C  was prepared (O.Olg in 100ml 0.25M sodium acetate buffer pH 5) and serial 
dilutions were made using the same buffer to obtain concentrations within the range 7.467­
0.933mM. Hundred microlitres o f each dilution were loaded into each well in triplicates 
followed by the addition o f 200|il o f 16mM 3, 3’, 5, 5’-Tetramethyl-Benzidine 
Dihydrochloride (TMBZ) solution. Twenty-five microlitres o f  3% hydrogen peroxide were 
then added to each well and the plate incubated for five minutes. Potassium phosphate buffer 
was used as the negative control. The plates were read as described at 620nm. Concentrations 
were plotted against absorbance to obtain a calibration curve.
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3.5.2.3 Non- specific esterases
Alpha and beta-napthol were used to obtain calibration curves for non-specific esterases. 
Thirty millimolar (30mM) solutions of a- and j8-napthol were prepared (0.0433g in 10ml 
acetone). Serial dilutions of the two stock solutions were made using 0.1M Potassium 
phosphate buffer to obtain concentrations within the range 7.5-0.17mM. Five microlitres of 
each dilution was loaded into wells in triplicates and 100|il o f  Fast Blue B salt solution 
added. The same buffer was used as the negative control. The plates were incubated at room 
temperature and read after 2 minutes as described at 540nm. Concentrations were plotted 
against absorbance to obtain a calibration curve.
3.5.3 Enzyme activity assays
For each test for activity o f  oxidases, Acetylcholine esterase and glutathione S-transferase 30 
mosquitoes were used, while for the non-specific esterases 20  mosquitoes were used.
3.5.3.1 Protein assay
Total protein content was determined in 2 batches of mosquito samples. Batch one comprised 
of samples from 4 field populations and the susceptible Kisumu strain which were used for 
oxidase, glutathione S-transferase and acetylcholinesterase assays. Batch two comprised o f 
samples from 3 field populations and the susceptible Kisumu strain which were used for the 
elevated non-specific esterase assays. For each mosquito homogenate 20|il was loaded into 
appropriate wells in the microplate, followed by 80pl o f 0 .1M potassium phosphate buffer 
Two hundred microlitres protein dye (Bradford reagent) were added and the plate read 
immediately (To) using an ELISA plate reader at 620 nm. Potassium phosphate buffer was
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used as the negative control. The quantity of protein (mg) per mosquito was estimated from 
the BSA calibration curve.
3.S.3.2 Oxidase assay
This assay measures the heme peroxidase levels in the test population. To conduct the assay, 
100|il o f mosquito homogenate were transferred into appropriate wells followed by lOOpl 
Tetramethyl-Benzidine Dihydrochloride (TMBZ) solution. Then 25 j l l 1 of 3% hydrogen 
peroxide was added into each well. Cytochrome-C and potassium phosphate buffer were 
used as positive and negative controls respectively. The plates were read after 5 minutes 
incubation as described at 620nm. The concentration o f  oxidase was calculated from a 
calibration curve obtained for cytochrome-C and the specific activity o f  oxidase expressed as 
mmole of product/min/mg protein per mosquito.
3.S.3.3 Acetylcholinesterase assay
This assay measures the amount o f acetylcholine esterase (AcChE) present. To conduct the 
assay, lOOpl o f  mosquito homogenate was transferred into in appropriate wells followed by 
the addition o f 100 pi o f 26mM Acetylthiocholine iodide (ATCH) solution into each well. 
0.1M potassium phosphate buffer was used as the negative control. Then lOOpl o f 3.3mM 
dithio-bis-2-nitrobenzoic acid (DTNB) solution was added into each well and the plate read 
immediately (To) with ELISA microplate reader using 414 nm filter. The plate was incubated 
and read again after ten minutes (Tio) and the difference between the two readings used for 
statistical analysis. The concentration o f AcChE produced was calculated using Beer’s law
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used as the negative control. The quantity of protein (mg) per mosquito was estimated from 
the BSA calibration curve.
3.5.3.2 Oxidase assay
This assay measures the heme peroxidase levels in the test population. To conduct the assay, 
100|al o f mosquito homogenate were transferred into appropriate wells followed by 1 OOp.1 
Tetramethyl-Benzidine Dihydrochloride (TMBZ) solution. Then 25(il o f 3% hydrogen 
peroxide was added into each well. Cytochrome-C and potassium phosphate buffer were 
used as positive and negative controls respectively. The plates were read after 5 minutes 
incubation as described at 620nm. The concentration o f oxidase was calculated from a 
calibration curve obtained for cytochrome-C and the specific activity o f  oxidase expressed as 
mmole of product/min/mg protein per mosquito.
3.5.3.3 Acetylcholinesterase assay
This assay measures the amount of acetylcholine esterase (AcChE) present. To conduct the 
assay, lOOul of mosquito homogenate was transferred into in appropriate wells followed by 
the addition o f 100 |il o f 26mM Acetylthiocholine iodide (ATCH) solution into each well. 
0.1M potassium phosphate buffer was used as the negative control. Then 100|al o f 3.3mM 
dithio-bis-2-nitrobenzoic acid (DTNB) solution was added into each well and the plate read 
immediately (To) with ELISA microplate reader using 414 nm filter. The plate was incubated 
and read again after ten minutes (Tio) and the difference between the two readings used for 
statistical analysis. The concentration o f AcChE produced was calculated using Beer’s law
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used as the negative control. The quantity o f  protein (mg) per mosquito was estimated from 
the BSA calibration curve.
3.5.3.2 Oxidase assay
This assay measures the heme peroxidase levels in the test population. To conduct the assay, 
100|il of mosquito homogenate were transferred into appropriate wells followed by 100 |il 
Tetramethyl-Benzidine Dihydrochloride (TMBZ) solution. Then 25\i\ o f 3% hydrogen 
peroxide was added into each well. Cytochrome-C and potassium phosphate buffer were 
used as positive and negative controls respectively. The plates were read after 5 minutes 
incubation as described at 620nm. The concentration o f oxidase was calculated from a 
calibration curve obtained for cytochrome-C and the specific activity o f oxidase expressed as 
mmole o f product/min/mg protein per mosquito.
3.5.3.3 Acetylcholinesterase assay
This assay measures the amount of acetylcholine esterase (AcChE) present. To conduct the 
assay, lOOfxl of mosquito homogenate was transferred into in appropriate wells followed by 
the addition of 100 jj,1 o f 26mM Acetylthiocholine iodide (ATCH) solution into each well. 
0.1M potassium phosphate buffer was used as the negative control. Then 100|il o f 3.3mM 
dithio-bis-2-nitrobenzoic acid (DTNB) solution was added into each well and the plate read 
immediately (To) with ELISA microplate reader using 414 nm filter. The plate was incubated 
and read again after ten minutes (Tio) and the difference between the two readings used for 
statistical analysis. The concentration of AcChE produced was calculated using Beer’s law
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(Hemingway, 1998). The specific activity was then expressed as mmole o f product/min/mg 
protein per mosquito.
3.5.3.4 Glutathione S-transferases
This assay measures the level o f Glutathione S-Transferase (GSTs) present in each mosquito. 
To conduct the assay, 1 OOjj.1 mosquito homogenate was transferred into each well in 
triplicates. Then 100^1 a 0.02mM reduced glutathione solution was added into each well 
followed by the addition o f 100 p.1 o f ImM l-chloro-2, 4 ’-dinitrobenzene (cDNB) solution 
and the plate read immediately (To) at 340nm filter. The plate was then incubated and read 
again after five minutes (T5) and the difference between the two readings (T5-T0) used for 
statistical analysis. The concentration o f GSTs produced was calculated by following the 
method of Hemingway (1998) using Beer’s law. The specific activity was then expressed as 
mmole of product/min/mg protein per mosquito.
3.5.3.5 Non-specific esterases
This assay measures levels o f non-specific B-esterases present. To conduct the assay, 100 jj.1 
mosquito homogenate was pippeted into each well and lOOpl o f 30mM a  and /? naphthyl 
acetate solutions added, and incubated at room temperature for ten minutes. One hundred 
microlitres of Fast Blue B salt solution was then added to each well and further incubated for 
two minutes. Potassium phosphate buffer and a//3-naphthol were used as the negative and 
positive controls respectively. The plates were read as described at 540 nm. The 
concentration of naphthol produced from the esterase reactions was calculated from standard
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curves obtained for a  and /3 naphthol. Results were expressed as mmole o f  product/min/mg 
protein per mosquito triturate.
3.6 Data Analysis
Abbott’s formula was used to correct the observed mortality in adult susceptibility tests 
(Abbott, 1925). The KT50 and KT95 values were estimated from the time mortality 
regression curves using probit analysis (Finney, 1971). Observed differences in resistance 
between susceptible and wild populations were analyzed by Student’s t-test. A one-way 
analysis of variance (ANOVA) was used to compare the protein content and enzyme 
expression levels between susceptible and wild populations. Chi-square test (X2) was used to 
test for relationships between site, molecular forms and kdr. All levels o f  statistical 
significance were determined at P<0.05.
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CHAPTER FOUR
RESULTS 
4.1 Bioassays
A total of 3,125 adult female An. gambiae s.l. were studied. For each site and for each test, 
100 test and 25 control mosquitoes were used. For permethrin, deltamethrin, malathion and 
propoxur 100 test and 25 control susceptible Kisumu strain mosquitoes were used while for 
DDT 80 test and 20 control were used.
4.1.1 Permethrin (0.75%)
Mortality in the field populations ranged between 24 and 48% across the 4 replicates with a 
mean of 32% (±10.83) for Adabraka, 60 to 100% with a mean o f 78% (±16.49) for Atabu 
Newtown. For Kledzo mortalities ranged from 72 to 100% with 82% (±13.27) mean 
mortality while in Likpe, mortality was between 44 and 64% with mean o f 57% (2.31). No 
mortality was scored o f the wild mosquito populations used for control tests. There was 
however 100% mortality in the susceptible Kisumu strain (Figure 4.1).
The median knockdown time (KT5o) obtained ranged between 39.1 - 65 minutes for the wild 
populations. In the Kisumu susceptible strain KDT5o was 10 minutes thus indicating a 3.9 to
6.5 fold increase in the wild populations. The KDT95 in wild populations ranged from 113.4 
to 147.1 minutes while in the Kisumu susceptible strain it was 25 minutes. The details are 
given in Table 3.
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4.1.2 Deltamethrin (0.05%)
Mortalities in the field populations ranged from 96-100% across the 4 replicates with a mean 
of 97% (±2.00) for Adabraka and Atabu Newtown while Kledzo had 97% (±3.83), thus 
indicating susceptibility in these populations. However for Likpe mortality ranged between 
84-100% with a mean o f 91% (±6.83), thus indicating reduced susceptibility (Figure 4.1). No 
mortality was scored o f the wild mosquito control populations but 100% mortality was 
recorded in the susceptible Kisumu strain.
The median knockdown time in wild populations was in the range o f 24.5 to 33.5 minutes 
while in the susceptible Kisumu strain KDT50 was 17 minutes thus indicating a 1.4 to 1.9 fold 
increase in wild population. The KDT95 ranged from 42.8 to 55 minutes in the wild 
populations while in the Kisumu susceptible strain it was 19 minutes (Table 3).
4.1.3 DDT (4%)
The mortalities observed for Adabraka mosquito population ranged from 8-24% with mean 
of 16% (±6.53), while in Atabu Newtown mortalities ranged from 40-64% with a mean of 
51% (±10.00). The least mortality rate was observed in the Kledzo population which ranged 
from 0-20% with a mean of 6% (±9.52). For the Likpe population, mortality ranged from 36­
56% with a mean o f 46% (±8.33). Results from these tests thus indicate very high levels o f 
resistance to 4% DDT in all the field populations while there was full susceptiblity in the 
susceptible Kisumu strain (Figure 4.1). No mortality was recorded among the control 
mosquito populations from the villages.
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The median knockdown time (KDT50) was recorded in only Atabu Newtown village and the 
susceptible strain populations. This is because less than 50% had been knocked down at the 
end of the exposure period for the other three villages. Thus the KDT50 was 77.5 and 23.4 
minutes for Atabu Newtown and the Kisumu susceptible strain respectively, thus indicating a
3.3 fold increase in the wild population as compared with the susceptible strain. KDT95 was
138.3 and 43.1 minutes for the two populations respectively (Table 4).
4.1.4 Malathion (5%)
Mortality in wild populations was 100% (±0.00) in all the 4 replicates tested across the four 
wild populations as well as with the Kisumu susceptible strain (Figure 4.2). Thus there was 
full susceptibility in all the test populations. Control mortality was 0% for both field 
populations and the susceptible strain.
4.1.5 Propoxur (0.1%)
In the wild populations, the mortality rates recorded were between 92-96% with a mean of 
95% (±2.00) for Adabraka and Kledzo and 94% (±2.31) for Atabu Newtown and Likpe 
populations respectively (Figure 4.2). There was 100% mortality for the Kisumu susceptible 
strain while with the control populations no mortality was recorded.
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4.1.6 Resistance ratios
Resistance ratios were determined for the pyrethroids. For permethrin the RR50 ranged from 
3.9 to 6.5 with the lowest and highest RR50 occuring in Kledzo and Adabraka populations 
respectively (Table 3). Similarly the highest mortality with permethrin was recorded in the 
Kledzo while the lowest was in Adabraka and therefore there was a similar trend in 
mortalities and RR50 in these populations. However this was not the case for Atabu Newtown 
in which despite having relatively high RR50 like in Adabraka, mortality was instead higher. 
With deltamethrin RR50 was very low when compared with permethrin and was in the range 
of 1.4 and 1.9 and unlike permethrin, the lowest RR50 occurred in Adabraka while the highest 
was in Kledzo.
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Table 3: Knockdown times (in minutes) for Permethrin, Deltamethrin and DDT in the four wild populations and the susceptible 
Kisumu strain and Resistance ratios.
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4.2 Molecular studies
A total o f200 specimens (50 per site) were processed by polymerase chain reaction (PCR) 
for species identification, molecular forms o f An. gambiae s.s. and kdr alleles.
4.2.1 Species and molecular forms identification
PCR amplification was successful in 174 (87%) specimens which were all positively 
identified as An. gambiae s.s. In Adabraka, of the 42 identified An. gambiae s.s, 40 were 
successfully identified to molecular forms, with 26 being ‘S’ and 14 being ‘M ’. Out o f the 46 
identified An. gambiae s.s in Atabu Newtown, 36 were successfully identified to molecular 
forms, and they comprised of 7 ‘S’ and 26 ‘M ’ forms. In Kledzo 33 An. gambiae s.s were 
successfully identified to molecular forms; 27 were ‘S’ and 6 were ‘M ’ forms. In Likpe, 38 
An. gambiae s.s were successfully identified to molecular forms; 35 were ‘S’ and 3 were ‘M ’ 
forms (Table 4).
4.2.2 Distribution of the kdr allele in An. gambiae s.s. population
In the four field populations, 63% o f the samples tested were kdr positive while 37% were 
kdr-negative. In Adabraka 24 specimens were kdr positive while 14 were kdr negative. In 
Atabu Newtown, 15 were kdr positive and 18 were negative. In Kledzo, o f  the 32 specimens 
23 possessed the kdr allele and 9 were kdr negative and in Likpe 32 o f the 45 studied were 
kdr positive. Over 70% o f the kdr mutation was associated with the ‘S’ form in the wild An. 
gambiae s.s populations.
Chi-square tests revealed no significant relationship between molecular forms and kdr 
(Pearson Chi-square value = 1.019, p = 0.313).
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Table 4: Molecular identification of Anopheles gambiae s.s samples into ‘M ’ and ‘S ’ forms and distribution of kdr alleles
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Table 5: Distribution of kdr alleles into the ‘M ’ and ‘S ’ forms of Anopheles gambiae s.s in the four field populations.
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8 M
— 600bp
_ 400bp
— 300bp
Figure 4.3: An example o f  electrophoresed ethidium bromide-stained 2.0% agarose gel o f 
PCR products for the detection of species o f An. gambiae complex. Lanes 1-6 = An. 
gambiae s.s. ; lane 7 = negative control; lane 8 = positive control; lane M = lOObp DNA size 
marker.
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M
400 bp 
300 bp
200 bp 
100 bp
Figure 4.4: An example o f electrophoresed ethidium bromide-stained 2.0% agarose gel o f 
Hhal restriction enzyme analysis o f An. gambiae s.s. PCR products for the detection o f M 
and S molecular forms. Lane M =100bp DNA size marker; lanes 1-3 = M form; lane 4 = S 
form; lanes 5-6 = M form; lane 7 = S form; lane 8 = undigested An. gambiae s.s. PCR 
product.
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400 bp —►
300bp —►
200bp —>
100 bp —►
Figure 4.5: An example o f electrophoresed ethidium bromide-stained 2.0% agarose gel of 
An. gambiae s.s. PCR products for the detection o f kdr alleles. Lane M =1 OObp DNA size 
marker; lanes 1-2 = kdr susceptible; lane 3 = kdr resistant; lanes 4 = kdr susceptible; lane 5-6 
= kdr resistant; lane 7 = negative control.
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4.3 Biochemical assays
A total of 190 wild mosquitoes and 80 from the susceptible Kisumu strain were assayed for 
the activities of oxidase, acetylcholine, glutathione S-transferase and esterase enzymes. 
Detailed information on absorbances obtained and enzyme activities o f individual 
mosquitoes are shown in appendix HI.
4.3.1 Calibration curves
The results obtained are illustrated in Figure 4.6. All values were in the range o f 92 -  99% 
and therefore all the curves exhibited good linearity hence indicating the reliability o f  the 
methods used.
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2.5
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4.3.2 Total protein content
In batch one mean total protein content was in the range o f 0.2 to 0.52 mg/ml in the wild 
populations while the susceptible Kisumu strain had a mean o f 0.53 mg/ml protein content 
(Table 6). For batch two, mean total protein content ranged from 0.22 to 0.35 mg/ml in the 
wild populations (Table 7). The susceptible Kisumu strain had a mean o f 0.47 mg/ml protein 
content.
4.3.3 Oxidase
Distribution pattern of oxidase activity is shown in Figure 4.7. Mean enzyme activity was in 
the range of 0.11 to 0.53 mmole product/min/mg protein in the wild populations. The 
susceptible Kisumu strain had mean activity o f 0.10 mmole product/min/mg protein (Table
6).
There was a significant increase in enzyme activity in Likpe population when compared to 
the susceptible Kisumu strain (P = 0.005). Similarly oxidase activity within the field 
populations was significantly higher in Likpe as compared to Adabraka (P = 0.005), Atabu 
Newtown (P = 0.005) and Kledzo (P = 0.005) populations.
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o or- o
d o O
H 4? -h -Hoo rr*- coco v00n co •VrOj
U o o d
o
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w 0  oA CN d 0 0 ft,
uo A 1  VO 1  1  (N co CN
< o d d cr~>
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CL ■s 3 1 
T3 cu
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Table 6  : Mean activity of Oxidase, Acetylcholinesterase (AcChE) and Glutathione-S-transferase (GST) in the four wild populations 
and the susceptible Kisumu strain.
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Figure 4.7: Distribution patterns of Oxidase activity in the four wild populations and the susceptible Kisumu strain.
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4.3.4 Acetylcholinesterase
Distribution pattern of AcChE activity is shown in Figure 4.8. Mean enzyme activity in wild 
populations ranged from 0.22 xlO '6 to 1.12 xlO '6 mmole product/min/mg protein. In the 
susceptible Kisumu strain mean activity o f  was 0.28 xlO"6 mmole product/min/mg protein 
(Table 6).
Enzyme activity was significantly increased in the Adabraka population as compared to the 
susceptible Kisumu strain (P = 0.009). Among the field populations, enzyme activity was 
also significantly higher in Adabraka than Atabu Newtown (P = 0.009), Kledzo (P = 0.009) 
and Likpe (P = 0.009) populations respectively.
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Adabraka mean=l 1.2 Atabu mean-2.14
Figure 4.8: Distribution patterns of AcChE activity in the four wild populations and the susceptible Kisumu strain.
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4.3.5 Glutathione S-Transferase
Distribution pattern o f GST activity is shown in Figure 4.9. In the wild populations mean 
enzyme activity was in the range of 0.31 xlO '3 to 0.77 xlO '3 mmole product/min/mg protein 
(Table 6). The susceptible Kisumu strain had a mean enxyme activity o f 0.56 xlO '3 mmole 
product/min/mg protein. Adabraka and Kledzo populations showed an increase in enzyme 
activity as compared to the susceptible Kisumu strain although it was not significant (P = 
0.056), while Atabu Newtown and Likpe populations had a much lower enzyme activity than 
the susceptible strain.
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( ^ 0 | . X )  Aj i a i j o v  ( , . 0 1 . * )  ^ 4 ! A ! J ° V
Activity = mmole product/min/mg protein
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4.3.6 Non-specific esterases
Distribution pattern o f  a-esterase activity is shown in Figure 4.10. Mean enzyme activity was 
in the range o f 0.23 xlO ' 1 to 0.94 xlO ' 1 mmole product/min/mg protein in the wild 
populations and 0.19 xlO ' 1 in the susceptible Kisumu strain (Table 7). Enzyme activity in 
the field populations o f Adabraka and Likpe was significantly higher when compared to the 
susceptible Kisumu strain (P = 0.000). However enzyme activity in Kledzo population was 
not significantly increased (P = 0.000)
Distribution pattern of ^-esterase activity is shown in Figure 4.11. Mean enzyme activity 
ranged from 0.90 xlO"1 to 2.55 xlO ' 1 mmole product/min/mg protein in wild populations 
while the susceptible Kisumu strain had a mean activity o f 0.72 xlO"1 mmole product/min/mg 
protein (Table 7). Likewise, enzyme activity in the field populations o f Adabraka and Likpe 
was significantly elevated when compared to the susceptible Kisumu strain (P = 0.000).
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Table 7: Mean activity of Non-specific esterases in the four wild populations and the susceptible Kisumu strain.
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c
□
(C0TO O
3'
(A
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II
n ^ « o o o
(j.Olx) A}iA!pv
(, 01.x) A4!auov
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University of Ghana http://ugspace.ug.edu.gh
4.3.7 Level of increase in enzyme activity
Increase in activity of oxidase was in the range o f 1.1 to 5.4 fold, with only the highest level 
of increase being significant and occurring in Likpe population. For AcCHE the level o f 
increase ranged from 0.8 to 4 fold and as with oxidase the highest level was significant but 
instead was in Adabraka population. Activity o f GSTs increased by between 0.6 to 1.4 fold, 
but there was no significant increase among the field populations. The level o f increase in 
activity had a similar pattern for both a- and /3-esterases (Table 8).
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<cDx
9
rG<D
JO
<D
>CO
c3oxcx
2o OS
;a£
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;3
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Table 8  : Level of increase in enzyme activity in field populations as compared with the 
susceptible Kisumu strain.
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CHAPTER FIVE
DISCUSSION
Insecticide resistance has been a problem in all insect groups that are vectors of diseases. 
Although mechanisms by which insecticides become less effective are similar across all 
vector taxa, each resistance problem is potentially unique and may involve a complex pattern 
of resistance foci. Regular testing is therefore vital for mosquito control operations because 
resistant populations o f mosquitoes reduce the effectiveness o f  control procedures. The main 
defense against resistance is close surveillance of the susceptibility o f  vector populations 
(Brogdon and McAllister, 1998). The initial step in identifying a potential problem is to 
detect changes in the susceptibility o f a population o f vectors through bioassays. Biochemical 
and molecular methods can detect resistance mechanisms in individual insects and therefore 
resistance can be confirmed with the use of only a small number of insects.
The actual molecular mechanisms that regulate insecticide resistance are poorly understood. 
Anopheles gambiae has multiple resistance mechanisms that have been field-selected in both 
East and West Africa through exposure to DDT and pyrethroids (Hemingway et al., 2002). 
Identification of resistance mechanisms helps determine the cross-resistance spectrum, 
facilitates the choice o f alternative insecticides and allows detailed mapping o f areas with 
resistant populations. This study therefore set out to determine the susceptibility status of An. 
gambiae complex towards various commonly used insecticides and to determine the 
underlying mechanisms of resistance by using molecular and biochemical assays.
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In this study, susceptibility tests results generally showed high levels of resistance to 
permethrin and DDT in filed populations of An. gambiae as shown by the very low 
mortalities. This is strongly supported by the greatly increased median knockdown times in 
the field populations. Higher KT5o values in field populations have been suggested to give an 
early indication of the involvement of kdr mechanism of resistance (Chandre et al., 1999a,
b). Deltamethrin and propoxur resulted in slightly reduced susceptibility levels while all 
tested samples were fully susceptible to Malathion. Deltamethrin, an a-cyano pyrethroid and 
more toxic than permethrin, despite giving very high mortality, still had a resistance ratio 
ranging from 1.4 to 2, thus implying low levels of resistance. These results are therefore 
consistent with those of other studies conducted in Accra, where An. gambiae s.l. has been 
reported to be resistant to permethrin and DDT (Adasi et al., 2000: Adeniran, 2002: 
Achonduh, 2005). However the results obtained are in contrast with those of Kristan et al.
(2003) who reported full susceptibility of An. gambiae s.l. field populations to permethrin 
and DDT in the Western Region of Ghana.
Morphological identification revealed An. gambiae s.s. as the dominant Anopheles mosquito 
species at all the study sites. These results are consistent with that of other studies carried out 
in the country (Adasi et al., 2000; Adeniran, 2002; Midega, 2002; Kristan et al., 2003 and 
Otieno, 2004). This is explained by the fact that the study area is comprised of a high altitude 
humid forest zone with higher rainfall making it is more favourable to An. gambiae s.s while 
An. arabiensis prefers more of dry savanna areas (Colluzi et al., 2000). In Ghana An. 
arabiensis has been found to predominate in the arid savanna of the northern region (Appawu 
etal., 1994).
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Both the ‘M’ and ‘S’ molecular forms of An. gambiae s.s were present and occurred in 
sympatry at all the study sites. Generally the ‘S’ form occurred at higher frequencies than the 
‘M’ form in the study area with an exception of one site, Atabu Newtown where the ‘M’ 
form occurred at a very high frequency than the ‘S’ form. This is a low lying swampy area 
that is normally flooded during the rainy season. The ‘M’ form has generally been associated 
with urban environments and flooded/irrigated areas while the ‘S’ form has been associated 
with rural areas. The fact that the ‘S’ form occurred at a very high frequency has serious 
implications in malaria control efforts since it has been strongly associated with the kdr 
mechanism of resistance (Chandre et al., 1999a,b). In this study the resistant phenotype of 
kdr was 70% in the ‘S’ form. Nevertheless it is very remarkable that 30% of resistant 
phenotype occurred in the ‘M’ form.
PCR identification of the kdr allele revealed both the resistant and susceptible phenotypes 
across all the collection sites. The resistance phenotype generally occurred at a higher 
frequency and was mainly associated with the ‘S’ form of An. gambiae s.s. Atabu Newtown 
however had more of the susceptible phenotype and this is mainly due the fact that the ‘M’ 
form which is less associated with the kdr occurred at a very high frequency. The observed 
high frequency of the resistant phenotype of kdr is supported by the high increase in 
knockdown times in field populations as compared with the susceptible Kisumu strain. This 
high kdr frequency by no means accounts for the very low mortalities observed in field 
populations with permethrin and DDT.
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Biochemical assays showed elevated levels of activity of detoxification enzymes in some 
field populations. Mean enzyme activity of oxidase was significantly higher in Likpe 
population as compared with the susceptible Kisumu strain. Likewise this population 
exhibited very high levels of permethrin and DDT resistance and thus strongly suggesting the 
implication of oxidases in addition to kdr in the observed resistance in this population. Mixed 
junction oxidases may be involved in the detoxification of almost all insecticides but are 
most notably associated with pyrethroid resistance (Hemingway & Ranson, 2000; Brooke et 
al., 2001; Chareonviriyaphap et al., 2003). In An. gambiae, metabolic resistance to 
pyrethroids has been shown to involve increased levels of cytochrome P450 (Ranson et al., 
2000). Work by Vulule et al. (1999) demonstrated the involvement of P450 enzymes in 
pyrethroid resistance in a Kenyan population of An. gambiae, where treatment with P450 
inhibitor piperonyl butoxide (PBO) partially reversed permethrin resistance in this strain.
Activity of both a. and /3 esterase was significantly increased in Adabraka and Likpe 
populations. Bioassays show that the Adabraka population had the highest levels of 
resistance to both permethrin and DDT. This strongly suggests the involvement of esterases 
as well as kdr and oxidases in the observed high resistance to permethrin and DDT in this 
population. Pyrethroids are insecticidal esters derived from primary alcohols and are thus 
susceptible to hydrolysis by esterases. Elevated esterase activity has been linked to 
pyrethroid resistance patterns in a variety of insects. Anopheles albimanus resistant to 
deltamethrin demonstrated elevated esterase levels in Guatemala (Brogdon et al., 1998). 
Chareonviriyaphap et al. (2000) found extremely high elevated esterase levels in the
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pyrethroid resistant An. albimanus colony in Guatemala, and they concluded pyrethroid use 
against them in Central America may be limited.
GST enzyme activity in field populations was not significantly elevated when compared with 
the susceptible Kisumu strain. Several studies have reported similar findings. 
Chareonviriyaphap et al. (2003) did not find any association between insecticide resistance in 
An. minimus with activity of GSTs. The mean level of GST activity across test populations of 
An. funestus in South Africa was not significantly higher than that of the corresponding 
susceptible reference strain (Brooke et al., 2001). There are currently no published records 
implicating GSTs in pyrethroid resistance in mosquitoes and the mixed function oxidase 
synergist PBO has not been implicated in GSTs activity inhibition (Brooke et al. 2001).
However GSTs have been implicated in resistance to DDT (Penilla et al., 1998; Ranson et 
al., 2000; Prapanthadara et al., 2002). GST activity was implicated in the observed high 
resistance to DDT and permethrin field populations of An. gambaiae in the Greater Accra 
area (Achonduh, 2005). The high level of DDT resistance detected in field populations of An. 
albimanus in southern Mexico was attributed to elevated levels of GST activity leading to 
increased rates of metabolism of DDT to DDE (Penilla et al., 1998). The GSTs of Anopheles 
gambiae have been studied extensively because of their involvement in DDT resistance. 
GSTs from a DDT-resistant strain of An. gambiae had an altered GST profile compared with 
susceptible insects (Prapanthadara et al., 1993). Ortelli et al. (2003) have indicated that DDT 
resistance in An. gambiae is associated with both qualitative and quantitative changes in 
multiple GST enzymes. Vontas et al. (2001) have implicated elevated GSTs with a
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predominant peroxidase activity to pyrethroid resistance in a laboratory selected colony of 
the brown planthopper Nilaparvata lugens.
There was a significantly elevated level of non-specific acetylcholinestrease in the Adabraka 
population as compared to the susceptible Kisumu strain. This does not seem to be consistent 
with the bioassay results which showed that Anopheles gambiae was fully susceptible to 
Malathion, an organophosphate while susceptibility to propoxur, which is a carbamate, was 
only slightly reduced in this population. However this slightly reduced susceptibility to 
propoxur could still be attributed to the activity of this enzyme. Studies by Penilla et al. 
(1998) attributed the carbamate resistance in An. albimanus population in Mexico to an 
altered acetylcholinesterase based resistance mechanism. Carbamate resistance as a result of 
insensitive acetylcholinesterase has recently been detected in Anopheles gambiae s.s. 
populations from C6te d'Ivoire (Corbel et al., 2003). In Accra Anopheles gambiae was 
reported to be fully susceptible to propoxur (Adeniran, 2002).
Elevated esterase occurs in a number of mosquitoes that are resistant to organophosphate and 
carbamate insecticides (Hemingway et al., 1998). Beach et al. (1989) implicated elevated 
esterases for organophosphate and pyrethroid resistance in An. albimanus from the coastal 
areas of Guatemala where organophosphate and carbamate insecticides have been used for 
agriculture. Malathion-specific carboxylesterase mechanisms were found in A. culicifacies 
and A. subpictus, both giving high rates of insecticide metabolism in Sri Lanka (Karunaratne 
and Hemingway, 2001). Thus the slight reductions in susceptibility to propoxur could be due 
to the activity of both esterases and acetylcholinesterase.
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Resistance is not evolving through unique new mechanisms but instead existing mechanisms 
are being enhanced, and cross-resistance is occurring. Multi-resistance (two or more 
resistance mechanisms in the same insect) is becoming widespread as control programs make 
sequential use of one chemical class after another (Brogdon et al., 1998; Brogdon et al., 
1999b). Vulule et al. (1999) suggested that use of impregnated nets selects for higher oxidase 
and esterase levels in An. gambiae to metabolize permethrin acquired from the nets. It has 
been suggested that deltamethrin selection appears to select initially a monooxygenase-based 
mechanism and when this mechanism is blocked by treatment with PBO, selection of a Mr- 
type mechanism is accelerated, as is evident from increased cross-resistance to DDT in the 
adults selected with deltamethrin-PBO (Kumar et al., 2004).
Similarly, P450-dependent oxidases were implicated in permethrin resistance in Culex 
quinquefasciatus Say from, C6te d'Ivoire and Burkina Faso in which kdr associated with 
DDT cross resistance and a dramatic loss of permethrin knockdown effect on adults was also 
involved (Chandre et al., 1998). Studies by Brogdon et al. (1999a) identified an oxidase 
resistance mechanism producing DDT-pyrethroid cross-resistance in Guatemalan An. 
albimanus Wiedemann. Interestingly, only adult female mosquitoes express the oxidase 
mechanism which coexists with an elevated esterase mechanism. Brooke et al. (2001) found 
the evidence for mixed function oxidase based cross-resistance between pyrethroids and the 
carbamate, propoxur in An. funestus rather unusual.
The findings of the current study are consistent with most of the other studies elsewhere as 
there is evidence of cross and multi-resistance. Results of bioassays, molecular and 
biochemical assays show an interesting trend. For example the occurrence of high levels of
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permethrin and DDT resistance is associated with the occurrence of significant levels of 
esterases, oxidases and a high frequency of the resistant kdr phenotype. In Adabraka and 
Likpe there was a significant increase in esterase activity as well as a high frequency of the 
resistant kdr phenotype. Furthermore activity of oxidases was significantly higher in the 
Likpe population. Thus there is a strong indication that both oxidase and esterase enzyme 
systems and kdr mechanisms are working in concert to confer the observed high levels of 
permethrin and DDT resistance in these An. gambiae populations.
The lack of a significant increase in enzyme activity in the other two field populations of 
Atabu Newtown and Kledzo is rather surprising as they both had high resistance to DDT and 
significantly reduced susceptibility to permethrin. Although esterase assay was not conducted 
in Atabu Newtown population none of the enzymes tested was significantly high either. 
However it is worth noting that occurrence of kdr was equally high in both populations and 
that it is probably the only mechanism conferring resistance. Another point worth 
considering is the fact that since it is the means which were used to compare the enzyme 
activity, there are higher chances that some individual mosquitoes in these populations might 
have enzyme activity high enough to metabolize some of these insecticides as is shown by 
the distribution patterns of enzyme activity.
Currently, the most advocated worldwide malaria vector control strategy appears to be the 
use of pyrethroid-impregnated bed nets. This is because they are less expensive than spraying 
walls with residual insecticides, are effective in reducing child deaths and can be better 
administered through a horizontal community-based program. The evolution of the kdr and 
biochemical mechanisms of resistance by no means poses a threat to this strategy. Initially
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there was an early optimism that because of their rapid toxicological action pyrethroids 
would not produce resistance.
In this study, resistance was generally high against permethrin, which is commonly used for 
impregnating bednets and curtains as well as indoor spraying against disease causing and 
biting nuisance mosquitoes. This could have serious implications in malaria transmission as 
well as compliance to ITN programmes. It has already been pointed out that any failure in 
nuisance control due to resistance is likely to demotivate people in using impregnated 
materials (Chandre et al., 1998). The high mortalities with deltamethrin gives hope that other 
pyrethroids are still effective in controlling An. gambiae populations. However the slight 
reduction in susceptibility as indicated by the slight increase in KT50 should be closely 
monitored. The same case applies to propoxur and malathion, since already the carbamates 
and organophosphates have been suggested as alternatives, should the pyrethroids fail to be 
effective in ITN programmes due to high levels of resistance (Darriet et al., 2003; Corbel et 
al., 2003; Hougard et al., 2003).
To compromise vector control with insecticides, the level of resistance must be high enough 
to adversely affect disease transmission and in many cases, vector control may not be 
affected by the observed levels of resistance in the vectors (Brogdon and McAllister, 1998). 
Even in areas of high pyrethroid resistance and high M r gene frequency in An. gambiae, 
deltamethrin impregnated bednets remain effective in vector control and should still be 
considered as a practical means of personal protection against malaria (Darriet et al., 2000). 
If for example, the level of resistance is lower than 10%, resistance will not affect disease 
control efforts (Brogdon and McAllister, 1998). Thus in this situation, increasing surveillance
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and monitoring level and frequency of resistance would be sufficient without the need for 
change in control methods.
Insecticide susceptibility has been described as a resource and resistance surveillance as an 
essential step in resistance management (Brogdon and McAllister, 1998).. Resistance 
surveillance has three objectives; first to provide baseline data for program planning and 
pesticide selection before the start of control operations. Secondly, to detect resistance at an 
early stage so that timely management can be implemented. Even detection of resistance at a 
late stage can be important in elucidating the causes of failure of disease control. However, in 
such cases, any management other than replacement of the pesticide may not be possible. 
Thirdly, resistance surveillance is essential to continuously monitor the effect of control 
strategies on resistance.
In conclusion, the current study forms a baseline survey in the study area and since no earlier 
study has been conducted, resistance has been inferred by comparison with the susceptible 
Kisumu strain. Since it is the first study in Hohoe and although resistance levels in An. 
gambaie populations in agree with those of other areas, there is need for regular surveillance 
to find out if resistance is rapidly evolving or it has stabilized. For example, in western 
Kenya, pyrethroid resistance appeared soon after bed nets were introduced. After 2 years, the 
resistance level had not changed significantly, possibly because of the continual massive 
introduction of susceptible genes (Vulule et al., 1996). This study has demonstrated the 
existence of several resistance mechanisms operating together in An. gambaie populations. It 
is therefore recommended that investigations on multi-resistance mechanisms in An. gambaie
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populations be conducted in different ecological zones of the country as most previous 
studies have been focusing only on the kdr mechanism.
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APPENDIX I
Preparation of standard solutions
The following standard solutions were prepared using sterile double distilled water 
(sddw).Where appropriate, the solutions were autoclaved at 1211 b/sq in. for 15minutes in an 
Eyela Autoclave (Rikikkaki, Tokyo)
1. Solutions for DNA extraction
a) Bender buffer
0.1M NaCl, 0.2 M sucrose, 0.1M Tris-HCl pH 7.5, 0.05M EDTA pH 9.1, 0.5% SDS. 
Stored at 4°C.
b) 0.5M EDTA (pH 8.0)
186.1 g/1 in water, pH adjusted with NaOH pellets and stored room temperature.
c) KAc(8M)
60ml of 5M KAc and 11.5ml glacial acetic acid in 28.5ml distilled water.
d) RNase
lOmg/ml. Sterilised by filtration and stored at-20°C
e) TE (pH 8.0)
lOmM Tris-HCl (pH 8.0), ImM EDTA (pH8.0). Stored at room temperature.
f) TE + RNnase (5 jig/ml)
5p.l of RNase (lOmg/ml) solution, 995 (il of TE (pH8.0). Stored at -20°C.
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2. Solutions for Electrophoresis
i) Agarose gels
10X TAE buffer
242g Tris base, 57.1ml glacial acetic acid, 100ml of 0.5 m EDTA, pH 
adjusted to 7.7 (with glacial acetic acid) and the volume made to 1000ml 
with ddw.
EtBr (lOmg/ml)
lg  of EtBr was completely dissolved in 100ml ddw and stored in the dark at 
room temperature.
ii) Gel loading buffer
Orange G
20% (w/v) Ficoll, 25 mM EDTA, 2.5% (w/v) orange G. Stored at 4°C.
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3. Solutions for Biochemical Assays
a) Buffers
i. 0.1M potassium phosphate buffer(pH7.2)
1.7g monobasic potassium phosphate in 454.4ml distilled water plus 6.6g 
dibasic potassium phosphate in 545.5ml distilled water. Adjust pH to 7.2 
Stored at room temperature
ii. 0.25M Sodium acetate (NaAc) buffer (pH 5.0).
Mix 83ml of 3M sodium acetate solution with 900ml of distilled water. 
Adjusted to pH5 using glacial acetic acid and the final volume made to 1 litre 
with distilled water. Stored at room temperature
b) Solutions for enzyme determination assays
a) j3-naphthyl acetate: 56 mg /3-naphthyl acetate was dissolved in 20 ml acetone
followed by addition of 80 ml of 0.1M K3PO4 buffer and stored at 
4°C.
b) a-naphthyl acetate: 56 mg a  -naphthyl acetate was dissolved in 20 ml acetone followed
by addition of 80 ml of 0.1M KjPCUbuffer and stored at 4°C.
c) Fast Blue solution: 150mg fast blue B salt was dissolved 15ml distilled water followed by
the addition of 35ml of 5% sodium lauryl sulfate. This was stored at in 
the dark and used within 2 hours after preparation
125
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d) TMBZ: 50mg 3,3',5,5'-Tetramethyl-Benzidine Dihydrochloride (TMBZ) was
dissolved in 25ml methanol and 75ml 0.25M NaAc buffer added. This
was at stored for a few days at 4°C.
e)cytochrome -C: O.Olg cytochrome-C was dissolved in 100ml 0.25M NaAc buffer and
used immmediately
f) Bradford reagent (protein dye concentrate): 20 ml Protein dye concentrate was mixed with
80 ml distilled H20  and stored indefinitely at 4°C in a light proof bottle
g) Reduced glutathione: 61 mg reduced glutathione was dissolved in 100 ml K3PO4 buffer
and stored for 3-4 days at 4°C.
h) cDNB: 20 mg l-chloro-2,4'-dinitrobenzene(cDNB) was dissolved in 10 ml
acetone followed by the addition of 90 ml KPO4 buffer . This was
stored for 3-4 days at 4°C.
i) ATCH: 75 mg acetylthiocholine iodide (ATCH) was dissolved inlO ml acetone
and made up to 1ml with 0.1M K3PO4 buffer. This was stored for 3-4
days at 4°C
j) DTNB: 13 mg dithio-bis-2-nitrobenzoic acid (DTNB) was dissolved in 100ml
K3PO4 buffer and stored for 3-4 days at 4°C.
126
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APPENDIX II 
W.H.O Susceptibility Tests Results
i) 0.75% Permethrin
a) Adabraka village
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 1 2 0 3
15 3 2 2 4
20 5 2 4 6
30 6 7 9 8
40 9 12 11 12
50 10 12 11 12
60 10 12 11 13
80 10 12 13 16
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 6 7 7 12
Observed mortality (%) 24 28 28 48
Control mortality (%) 0 0 0 0
Corrected mortality (%) 24 28 28 48
127
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b) Atabu Newtown
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 1 0 1 0
15 3 2 3 3
20 5 6 5 6
30 10 10 6 7
40 13 14 7 9
50 13 15 7 9
60 13 17 8 12
80 13 18 9 12
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 15 19 19 25
Observed mortality (%) 60 76 76 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 60 76 76 100
128
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c) Kledzo village
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 1 1 1 1
15 3 2 7 4
20 6 3 11 15
30 11 11 16 19
40 15 14 16 20
50 15 14 16 20
60 17 16 18 21
80 19 18 19 21
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 21 18 18 25
Observed mortality (%) 84 72 72 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 84 72 72 100
129
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d) Likpe village
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 0 0 1
15 4 4 2 5
20 7 5 4 8
30 11 12 9 13
40 13 14 10 13
50 13 15 12 13
60 14 15 15 14
80 16 20 19 15
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 16 11 15 15
Observed mortality (%) 64 44 60 60
Control mortality (%) 0 0 0 0
Corrected mortality (%) 64 44 60 60
130
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e) Susceptible Kisumu strain
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 7 14 15 16
15 14 17 18 21
20 20 20 22 23
30 23 22 24 25
40 25 25 25 25
50 25 25 25 25
60 25 25 25 25
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 24 24 23 24
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 4 0 4
Corrected mortality (%) 100 100 100 100
131
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ii) 0.05% Deltamethrin
a) Adabraka
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 5 4 0 6
15 6 7 3 7
20 11 13 11 9
30 15 21 20 16
40 20 23 22 19
50 22 22 24 23
60 25 25 25 23
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 24 24 25 24
Observed mortality (%) 96 96 100 96
Control mortality (%) 0 0 0 0
Corrected mortality (%) 96 96 100 96
132
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b) Atabu Newtown
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 1 0 1 1
15 4 3 2 3
20 11 7 5 10
30 15 23 18 17
40 24 24 24 23
50 24 25 25 24
60 24 25 25 24
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 24 24 25 24
Observed mortality (%) 96 96 100 96
Control mortality (%) 0 0 0 0
Corrected mortality (%) 96 96 100 96
133
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c) Kledzo
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 0 1 0
15 0 0 2 2
20 6 3 5 7
30 11 8 8 14
40 17 21 13 22
50 20 23 22 25
60 21 25 24 25
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 23 24 25 25
Observed mortality (%) 92 96 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 92 96 100 100
134
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d) Likpe
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 3 0 4
15 4 7 3 8
20 8 13 6 12
30 15 20 22 20
40 22 20 22 24
50 24 21 22 24
60 24 23 24 24
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 25 21 23 22
Observed mortality (%) 100 84 92 88
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 84 92 88
135
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e) Susceptible Kisumu strain
No. knocked down after exposure Replicates 
1 2 3 4
for minutes
10 6 5 6 8
15 13 7 10 14
20 20 14 15 16
30 24 24 20 25
40 25 25 23 25
50 25 25 25 25
60 25 25 25 25
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 4 4 0 0
Corrected mortality (%) 100 100 100 100
136
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iii) 4% DDT
a) Adabraka
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 0 0 0
15 0 0 0 0
20 0 0 0 0
30 0 0 0 0
40 0 0 0 0
50 2 0 1 0
60 2 0 1 0
80 2 0 1 0
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality at the end of holding period 6 4 2 4
Observed mortality (%) 24 16 8 16
Control mortality (%) 0 0 0 0
Corrected mortality (%) 24 16 8 16
137
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b) Atabu Newtown
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 0 0 0
15 0 1 0 0
20 1 1 0 0
30 2 5 2 2
40 3 7 10 7
50 5 8 11 8
60 6 9 11 10
80 7 10 11 11
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality at the end of holding period 13 16 10 12
Observed mortality (%) 52 64 40 48
Control mortality (%) 0 0 0 0
Corrected mortality (%) 52 64 40 48
138
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c) Kledzo
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 0 0 0
15 0 0 0 0
20 0 0 0 0
30 0 0 0 0
40 0 0 0 0
50 2 0 0 0
60 0 0 0 0
80 2 0 0 0
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality at the end of holding period 5 0 0 1
Observed mortality (%) 20 0 0 4
Control mortality (%) 0 0 0 0
Corrected mortality (%) 20 0 0 4
139
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d) Likpe
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 0 0 0
15 0 0 0 0
20 0 0 0 1
30 0 0 0 3
40 1 0 0 5
50 2 2 1 6
60 2 2 1 6
80 4 5 2 9
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality at the end of holding period 11 9 12 14
Observed mortality (%) 44 36 48 56
Control mortality (%) 0 0 0 0
Corrected mortality (%) 44 36 48 56
140
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e) Susceptible Kisumu strain
No. knocked down after exposure Replicates
1 2 3 4
for minutes
10 0 2 4 2
15 1 2 7 4
20 15 13 19 19
30 16 16 20 20
40 20 18 20 20
50 20 18 20 20
60 20 18 20 20
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality at the end of holding period 20 20 20 20
Observed mortality (%) 100 100 100 100
Control mortality (%) 4 0 0 4
Corrected mortality (%) 100 100 100 100
141
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iv) 5% Malathion
a) Adabraka
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 25 25 23 25
Mortality a the end of holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 100 100 100
b) Atabu Newtown
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 25 25 24 25
Mortality a the end of holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 100 100 100
142
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c) Kledzo
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 25 23 25 23
Mortality a the end of holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 100 100 100
d) Likpe
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 24 24 25 24
Mortality a the end of holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 4 0 0 0
Corrected mortality (%) 100 100 100 100
143
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e) Susceptible Kisumu strain
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 25 25 25 25
Mortality a the end of holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 100 100 100
144
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v) 0.1% Propoxur
a) Adabraka
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 25 25 25 25
Mortality a the end of holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 100 100 100
b) Atabu Newtown
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 23 21 24 23
Mortality a the end of holding period 24 23 23 24
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 4
Corrected mortality (%) 96 92 92 96
145
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c) Kledzo
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end of exposure period 21 21 18 25
Mortality a the end of holding period 24 23 24 24
Observed mortality (%) 96 92 96 96
Control mortality (%) 0 0 0 0
Corrected mortality (%) 96 92 96 96
d) Likpe
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end o f  exposure period 25 25 25 25
Mortality a the end o f  holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 100 100 100
146
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e) Susceptible Kisumu strain
Replicates
1 2 3 4
Total exposed 25 25 25 25
Total control 25 25 25 25
Mortality a the end o f  exposure period 25 25 25 25
Mortality a the end o f  holding period 25 25 25 25
Observed mortality (%) 100 100 100 100
Control mortality (%) 0 0 0 0
Corrected mortality (%) 100 100 100 100
147
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0.990667  I 0.239933  | 1.1273 0.4686 | 0 .935 0.146 0.7008
0.974667 10 .213087  | 1.019 | 0 .286913  | 1.1617 0.5263 1.113 0 .4446 1.1797 0.5565
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1 1.021667  | 0 .290268  | 1.275333 | 0 .716443  | 1.1373 0 99 0.2383 1.205 0.599
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