ASSESSMENT OF DIFFERENT STAINS AND STAINING PROCEDURES
FOR MICROSCOPIC DETECTION OF MALARIA PARASITES
By:
Awad Alla Hamza Osman Kashif
BSc of Medical Laboratory Sciences
(1995)
A thesis submitted for fulfillment requirement of master degree
in Medical Laboratory Sciences (Medical Parasitology)
At the
Department of Parasitology & Medical Entomology
Faculty of Medical Laboratory Sciences
University of Khartoum
Supervisor: Dr. Eldirdieri Salim Ahmed College of Medicine –University of Juba
September 2004
LIST OF CONTENTS
DEDICATION………………………………………………………………….. I ACKNOWLEDGEMENT.…………………………………………………….. II LIST OF TABLES……………………………………………………………... III LIST OF FIGURES……………………………………………………………. IV LIST OF APPENDICES……………………………………………………….. V ABSTRACT…………………………………………………………………….. VI ABSTRACT (ARABIC)………………………………………………………... VII
1- INTRODUCTION & LITREATURE REVIEW 1
1.1 General introduction……………………………………………………….…... 1 1.2 Historical background…………………………………………………………. 1 1.3 Malaria parasites………………………………………………………………. 3 1.4 Transmission of malaria……………………………………………………….. 4 1.5 The life cycle of malaria……………………………………………………….. 8 1.5.1 Pre-erythrocytic phase……………………………………………………….. 8 1.5.2 Erythrocytic phase…………………………………………………………… 9 1.5.3 Vector phase (Sporogony)……………………………………………….…... 9 1.6 Pathogenesis of malaria………………………………………………………... 12 1.7 Immunity to malaria…………………………………………………………… 14 1.8 Diagnosis of malaria……………………………………………………….…... 15 1.8.1 Clinical diagnosis……………………………………………………………. 15 1.8.2 Microscopic diagnosis……………………………………………………….. 16 1.8.2.1 Conventional thick and thin blood smears………………………………… 18 1.8.2.1.1 Blood drawing technique…………………………………………….….. 19 1.8.2.1.2 Preparation of the smears…………………………………………….….. 19
1.8.2.1.3 Staining procedures…………………………………………………. 21 1.8.2.1.4 Microscopic examination. ………………………………………….. 23 1.8.2.2 Quantitative Buffy Coat (QBC) Test…………………………………. 24 1.8.2. 3 Saponin lyzing technique…………………………………………….. 24 1.8.3 Rapid Diagnostic Tests (RDTs)………………………………………… 25 1.8.4 Deoxyribonucleic acid (DNA) probes and Polymerase Chain Reaction 26 (PCR)...... 1.8.5 Detection of Plasmodia specific antibodies……………………………. 26 1.9 Chemotherapy of malaria…………………………………………………. 27 1.10 Malaria in the Sudan…………………………………………………….. 29 1.10.1 Transmission …………………………………………………………... 29 1.10.2 Morbidity and mortality ………………………………………………. 30 1.10.3 Diagnosis of malaria...... 30 1.10.4 Chemotherapy of malaria...... 31 1.10.5 Control………………………………………………………………… 32 1.11 Justification……………………………………………………………… 34 1.12 Objectives………………………………………………………………... 34 1.12.1 General objective……………………………………………………… 34 1.12.2 Specific objectives…………………………………………………….. 34 2. MATERIALS & METHODS 35 2.1 Study population………………………………………………………….. 35 2.2 Study design ……………………………………………………….……… 35 2.3 Samples collection ………………………………………………………... 35 2.4 Techniques………………………………………………………………... 35 2.4.1 Giemsa stained smears………………………………………………….. 35 2.4.2 Saponin lyzed venous blood technique …………………………………. 36 2.4.3 Field’s stain smears ……………………………………………………... 37 2.4.4 Preparation of different Giemsa stains stock solution…………………... 37 2.4.5 Concentration of Giemsa working solution…...………………………… 38 2.4.6 Dilution of stock Giemsa stain with distilled or tap water……………… 39 2.4.7 Stability of Giemsa stain working solution……………………………... 40 2.5 Ethical consideration……………………………………………………… 41 2.6 Data analysis……………………………………………………………… 41
3. RESULTS 42 3.1 Giemsa stained blood smears…………………………………………….. 42
3.2 Saponin lyzed venous blood technique …………………………………... 46
3.3 Comparison between Giemsa and Field’s stain in detecting malaria 48 parasites......
3.4 Preparation of different Giemsa stains stock solutions…………………… 50
3.5 The use of different concentrations of Giemsa stain working solutions. ... 52
3.6 Preparation of Giemsa stain working solution using different diluents.... . 54
3.7 Stability of Giemsa stain working solution……………………………….. 56
4. DISCUSSION 58
CONCLUSION & RECOMMENDATIONS……………………………… 63
REFERENCES………………………………………………………………. 64
APPENDICES……………………………………………………………….. 83
DEDICATION
To my mother, soul of my father & my brothers
To all my teachers from primary school to the university
I ACKNOWLEDGMENT
I am greatly indebted to my supervisor Dr. Eldirdieri Salim Ahmed, Department of Microbiology & Parasitology, College of Medicine, University of Juba, for his continuous encouragement and valuable advices during the whole period of the
s t u d y .
My thanks are due to all those who helped me, Mr. Omer Mohamed Ali, Mr.
Yousif Adam Omer, and my great thanks also to Mrs. Sara Mohamed Al Aalim,
Mr. Anawar Ahmed, Mr. Al sadig Al bakheet & Mr. Yousif Manoly.
My thanks to my colleagues, Mr. Said Ali Mustafa & Mr. Luai Osman
Ibrahim, for their help in examination of the blood smears and my thanks also to
Mrs. Enaam Husein, Mr. Mamoun Magzoub, Mr. Mohamed Abdel hadi, Mr
Khaliefa Al tayeb & Mr Ahmed Baher f o r t h e i r g r e a t s h e l p s .
Special thanks should go to Mr. Awad Ahmed Nasr the Head Department of
Parasitology & Medical Entomology, Faculty of Medical Laboratory Sciences,
University of Khartoum for his encouragement and great help to conduct this study.
II LIST OF TABLES
Table 1. Comparison between Giemsa , Saponin lyzed and Field’s stained thick 44 blood smear in detecting malaria parasite and in determining the parasite count/µl blood ......
Table 2. Comparison between Giemsa stained-thick blood smears and Giemsa 47 stained-Saponin lyzed venous blood smears for detecting malaria parasites examined by three technicians ......
Table 3. Comparison between Giemsa and Field stained thick blood smears 49 examined by three technicians in the detection of malaria parasites......
Table 4. Comparison of the stain quality of thick & thin blood smears using 51 different Giemsa stain stock solutions ......
Table 5. Comparison between different concentrations (10%, 15% and 20%) of 53 Giemsa stain working solution in staining thick & thin blood smears ......
Table 6. Comparison of Giemsa stain working solution for staining blood 55 smears using different diluents ......
III LIST OF FIGURES
Figure1. Distribution of Malaria parasites over the world ...... 7
Figure 2. Life Cycle of Malaria parasites ...... 11
Figure 3. Transmission of Malaria in the Sudan ...... 33
Figure 4. Age group distribution of the study population ...... 43
Figure 5. Relationship between detection of malaria parasites according to level 45
of parasiteamia in blood smears stained with different stain ......
Figure 6. The percentage of distinguishable component of blood smears (stain 57 quality) following different intervals of storage………………………………….
LIST OF APPENDICES
IV Appendix 1. Relationship between detection of malaria parasites according to 83 level of parasiteamia in blood smears stained with different stains......
Appendix 2. Comparison between the quality of Giemsa stain working 84 solution and time after preparation of the working solution ......
Appendix 3. Preparation of stock Giemsa stain (Stock I )...... 85
Appendix 3a. Preparation of stock Giemsa stain (Stock I I)...... 86
Appendix 3b. Preparation of stock Giemsa stain (Stock III) ...... 87
Appendix 4. Preparation of Field’s stain solution A ...... 88
Appendix 4a. Preparation of Field’s stain solution B ...... 89
Appendix 5. Preparation of Saponin lyzing solution...... 90
Appendix 6. Preparation of phosphate buffered solution pH 7.2 ...... 91
Appendix 7. Questionnaire ...... 92
Appendix 8. Form used to assess the blood smears of study techniques ...... 93
Abstract
V This study was carried out during the period from July 2002 to September 2003 in order to assess the use of different stains and staining procedures in microscopic diagnosis of malaria.
A total of 203 individuals were included in the present study, Field-stained thick blood smear, Giemsa-stained thick and thin blood smears and Saponin lysed Giemsa- stained smears were prepared from each individual. Three different technicians examined stained blood smears independently.
The result showed superiority of Giemsa-stained thick blood smears in detecting malaria parasites over others stained smears. Using Giemsa-stained thick blood smears malaria parasites were detected in 31, 32.5, 33 % whereas it was detected in
28.6, 29.5,30%, by Saponin lyzed venous blood smears and only in 26.1, 27.1%,
27.1% of Field stained smears as examined by three technicians.
Further, three stocks solutions of Giemsa stain prepared by different methods were evaluated in staining blood smears, the finding showed a comparable result between the three Giemsa stain stock solutions.
Unsatisfactory results were obtained when higher concentrations of Giemsa working solutions (15% & 20%) were used for shorter staining period versus convential concentration (10%).
Adequate results were obtained by working solutions of Giemsa stain prepared using distilled water instead of phosphate buffered solutions, whereas poor results were obtained when a tap water was used.
The study findings showed that it is possible to obtain Giemsa working solution with satisfactory staining properties, which did not decrease during the first 3 hours of preparation.
اﻟﺨـــﻼﺻــﺔ
VI أﺟﺮﻳﺖ هﺬﻩ اﻟﺪراﺳﺔ ﻓﻲ اﻟﻔﺘﺮة ﻣﻦ ﻳﻮﻟﻴﻮ 2002 إﻟﻰ ﺳﺒﺘﻤﺒﺮ 2003 ﻟﺘﻘﻴﻴﻢ اﺳﺘﺨﺪام أﺻ ﺒﺎغ و ﻃ ﺮق
ﺻﺒﻎ ﻣﺨﺘﻠﻔﺔ ﻓﻲ ﻣﺠﺎل اﻟﻔﺤﺺ اﻟﻤﺠﻬﺮي ﻟﻄﻔﻴﻠﻴﺎت اﻟﻤﻼرﻳﺎ .
أﺧ ﺬت ﻋﻴﻨ ﺎت دم ﻣ ﻦ 203 ﺷ ﺨﺺ و اﻟ ﺬﻳﻦ ﻳﻤﺜﻠ ﻮن ﺷ ﺮﻳﺤﺔ اﻟﺪراﺳ ﺔ اﻟﺤﺎﻟﻴ ﺔ ﺛ ﻢ ﺑﻌ ﺪ ذﻟ ﻚ ﺗ ﻢ ﺗﺤ ﻀﻴﺮ
ﻣﺴﺤﺎت دم ﺳﻤﻴﻜﺔ ﺻﺒﻐﺖ ﺑﺼﺒﻐﺔ اﻟﻔﻴﻠﺪ و ﻣ ﺴﺤﺎت دم ﺳ ﻤﻴﻜﺔ و رﻗﻴﻘ ﺔ ﺻ ﺒﻐﺖ ﺑ ﺼﺒﻐﺔ اﻟﺠﻴﻤ ﺴﺎ ﺑﺎﻹﺿ ﺎﻓﺔ إﻟ ﻲ
ﻣ ﺴﺤﺎت ﻣ ﻦ اﻟ ﺪم اﻟﻮرﻳ ﺪي اﻟﻤﻌ ﺎﻟﺞ ﺑﻮاﺳ ﻄﺔ ﻣ ﺎدة اﻟ ﺴﺎﺑﻮ ﻧﻴﻦ ﺻ ﺒﻐﺖ أﻳ ﻀﺎ ﺑ ﺼﺒﻐﺔ اﻟﺠﻴﻤ ﺴﺎ . ﻋﻨ ﺪ ﻓﺤ ﺺ آ ﻞ
ﻣﺴﺤﺎت اﻟﺪم اﻟﻤﺤ ﻀﺮة ﺑﻮاﺳ ﻄﺔ ﺛﻼﺛ ﺔ ﺗﻘﻨﻴ ﻴﻦ ﺑ ﺼﻮرة ﻣ ﺴﺘﻘﻠﺔ آ ﺎن هﻨﺎﻟ ﻚ ﺗﻤﻴ ﺰ واﺿ ﺢ ﻟﻠﻤ ﺴﺤﺎت اﻟﻤ ﺼﺒﻮﻏﺔ
ﺑﺼﺒﻐﺔ اﻟﺠﻴﻤﺴﺎ ﻓﻰ ﺗﺤﺪﻳﺪ وﺟ ﻮد ﻃﻔﻴﻠﻴ ﺎت اﻟﻤﻼرﻳ ﺎ، ﺣﻴ ﺚ آﺎﻧ ﺖ ﻧ ﺴﺒﺔ اﻟﻤ ﺴﺤﺎت اﻟﻤﻮﺟﺒ ﺔ ﻟﻠﺜﻼﺛ ﺔ ﺗﻘﻨﻴ ﻴﻦ ﻋﻠ ﻰ
اﻟﺘﻮاﻟﻰ هﻰ 31 ، 32.5 و 33% ﺑﻴﻨﻤ ﺎ آ ﺎﻧ ﺖ اﻟﻨ ﺴﺒﺔ ﻟﻤ ﺴﺤﺎت اﻟ ﺴﺎﺑﻮﻧﻴﻦ ه ﻲ 28.6،29.5 و 30% و ﺑﺎﻟﻨ ﺴﺒﺔ
ﻟﻠﻤﺴﺤﺎت اﻟﻤﺼﺒﻮﻏﺔ ﺑﺼﺒﻐﺔ اﻟﻔﻴﻠﺪ آﺎﻧﺖ اﻟﻨﺴﺒﺔ ﻓﻘﻂ هﻲ 26.1،27.1 و %27.1.
ﻓﻴﻤﺎ ﻳﺨﺘﺺ ﺑﺼﺒﻐﺔ ﺟﻴﻤﺴﺎ ﺗﻢ اﻟﺤﺼﻮل ﻋﻠﻰ ﻧﺘﺎﺋﺞ ﻣﺘﺸﺎﺑﻬﺔ ﻋﻨﺪ اﺳﺘﺨﺪام اﻻﺻﺒﺎغ اﻟﻤﺨﺰﻧﺔ اﻟﻤﺤ ﻀﺮة
ﺑﻨﺴﺒﺔ آﺤﻮل اآﺒﺮ ﻣﻦ اﻟﺠﻠﻴﺴﺮﻳ ﻦ وﺑ ﺎﻟﻨ ﺴﺒﺔ ﻟﻠﻤﺤﺎﻟﻴ ﻞ اﻟﻌﻤﻠﻴ ﺔ ﻳﻤﻜ ﻦ اﻟﺤ ﺼﻮل ﻋﻠ ﻰ ﻧﺘ ﺎﺋﺞ ﻣﺮﺿ ﻴﺔ ﺑﺘﺨﻔﻴ ﻒ
اﻟﻤﺤﻠﻮل اﻟﻤﺨﺰن ﺑﻮاﺳﻄﺔ ﻣﺤﻠﻮل اﻟﻔﻮﺳﻔﻴﺖ اﻟﺪارئ ب .ﻩ 7.2 أو اﻟﻤﺎء اﻟﻤﻘﻄﺮ و ﻟﻜﻦ ﻟﻴﺲ ﻣﺎء اﻟﺼﻨﺒﻮر و
ﻟﻢ ﻳﺘﻢ اﻟﺤﺼﻮل ﻋﻠﻰ ﻧﺘﺎﺋﺞ ﻣﺮﺿﻴﺔ ﻋﻨﺪ زﻳﺎدة ﺗﺮآﻴ ﺰ اﻟﻤﺤﻠ ﻮل اﻟﻌﻤﻠ ﻲ ﻣ ﻊ ﺗﻘﻠﻴ ﻞ زﻣ ﻦ اﻟ ﺼﺒﻎ و أﺧ ﻴ ﺮا
وﺟﺪ أن اﻟﻤﺤﺎﻟﻴﻞ اﻟﻌﻤﻠﻴﺔ ﻟﻠﺼﺒﻐﺔ ﺗﻔﻘﺪ ﺧﺼﺎﺋﺼﻬﺎ ﺑﻌﺪ ﻣﺮور 3 ﺳﺎﻋﺎت ﻣﻦ زﻣﻦ اﻟﺘﺤﻀﻴﺮ.
VII CHAPTER ONE
INTRODUCTION & LITERATURE REVIEW
1.1 General introduction:
Malaria is the highest single cause of morbidity and mortality throughout the tropic and subtropics. (Greenwood & Mutabingwa, 2002). Despite of the effort directed towards the control of malaria over the world for many years, nevertheless, malaria remains one of the most prevalent vectors – borne diseases in terms of life lost and economic burden. According to the World Health Organization (WHO), approximately
300 million to 500 million clinical cases of malaria are reported worldwide yearly; they live unprotected from malaria due to lack of the necessary physical and human resources. (Sacks & Malaney, 2002). Malaria is endemic in over 90 countries lying between latitude 60 N° and 40 S° about 40 % of the world population is at risk and 90% of them are in Africa (south of the Sahara) (Guyatt & Snow, 2000). Worldwide, malaria is responsible for about 2 million cases of deaths annually, particularly, among children below 5 years of age. (Jennifer, 2003).
1.2 Historical background:
Malaria has been known since antiquity, with recognizable description of the disease recorded in various Egyptian papyri. The Ebers papyrus (1550 BC) mention fever, splenomegaly and use oil of Balanites tree as a mosquito repellent. Hieroglyphs on the wall of the ancient temples of Denderah in Egypt describe an intermittent fever following the flooding of the Nile (Nunn & Tapp. 2000).
1 The clinical manifestations of seasonal and intermittent fever have been recorded in ancient Asian Chinese and Indian medical texts and classical malaria or Roman fever was common in the vicinity of Rome and cyclical epidemics of malaria continued in many parts of Europe and other continents throughout many centuries. The early opinion was that the disease was spread by miasma, mist, arising in marshes so the name malaria is driven from Italian mal aria, meaning “ bad air”. (Cox, 2002).
In the fifth century Bc, in Greece, Hippocrates was the earliest physician who made logical observations of the relationship between the appearance of the disease and the season of the year or the places where his patients lived. (Bruce-Chwatt, 1988). The most important events in the history of malaria took place towards the end of the nineteenth century, when the sciences of bacteriology and pathology were discovering the causes of infectious diseases, observing the morbid changes in the organs and tissues and also perceived the role of the insect in transmission of some diseases, but no progress was made in etiology of malaria until 1847,when Meckel observed black pigment granules in the blood of a patient who died of the disease and in 1879 Afanasiev suggested that granules caused the disease. In 1880 the French army surgeon Laveran in Algeria, was the first scientist to see and described malaria parasites in the red blood cells of man. Laveran accurately described the male and female gametes, the trophozoite, and the schizont in unstained preparations, by 1890 several scientists in different parts of the world verified Laveran’s finding. In 1891,Romanowsky, in Russia, developed a new method of staining blood smears based on methylene blue and eosin to demonstrate malaria parasites as stained objects, and modifications of this stain remain in wide use. (Bruce-Chwatt, 1985).
2 However, the way by which the disease was transmitted from man to man was still a mystery although a few early and inspired guesses pointed the possible association between swamp, mosquitoes, and fevers Sir Patrick Manson, in 1894 put forward the theory that mosquitoes transmit malaria from man to man, he was conditioned by the proof of mosquitoes as vectors of filariasis. In 1897, Ross in India discovered pigmented cysts (oocysts) on the stomach of an Anopheles mosquito that had previously fed on a patient with plasmodium in his blood (Rajakumar & Weisse, 1999). The role of female
Anopheles mosquito in the transmission of malaria has been conformed by the combined field experiments carried out by Patrick Manson and his colleagues near Rome and
London in 1900. This was proved by showing that protection from the bites of
Anopheles mosquito prevent the occurrence of the infection (Schmidt & Robert, 1989).
The whole complex picture of the life cycle of development malaria parasites in man and in Anopheles mosquito became clear as a result of further studies by the Italians:
Amico Bignami, Giuseppe Bastianelli and Battista Grassi in 1898-1899 (Ascenzi, 1999).
1.3 Malaria parasites:
Malaria parasites of man are species of the genus Plasmodia, family Plasmodiidae, suborder Haemosporina, order Eucoccidiida, subclass Coccidia, and the class Sporozoa
(Beaver et al., 1983). There are four species (falciparum, vivax, ovale and malariae) of the genus plasmodium known to infect man and able to cause significant disease
(Snounou et al., 1993). P. falciparum causes malignant tertian malaria; it is the most dangerous among the above- mentioned species and is responsible for the majority of malaria related deaths (WHO, 1996). In falciparum malaria fever occurs every 48 hours, however, the periodicity often masked because the stages are not always synchronous.
3 The term tertian was applied for such periodicity because fever occurs every third day.
The development of P. falciparum needs an average ambient temperature of at least
20°C that is found mainly in warmer parts of the world. P. vivax, causes benign tertian malaria, which rarely have a fatal outcome. It can exist in places with an average summer temperature of only 16°C, together with P. ovale is considered relapsing malaria, so named because it can remain in liver as a dormant hypnozoite stage for very long periods (years) in the liver. The adaptive value of this ability is that the parasite can persist in areas that experience long winters with no opportunities for transmission.
Plasmodium ovale causes rare tertian malaria with long incubation period and relapse at three-month intervals. It is found mainly in tropical Africa but with sporadic reports from elsewhere. Plasmodium malariae causes quartan malaria with fever returning every
72 hours. It is remarkable in that it can persist in the blood of a host for decades at very low densities, but it does not have a dormant stage in liver. Recrudescence was reported after half a century following the infection (Knight, 19985).
1.4 Transmission of malaria:
Malaria occurs in most tropical regions of the world with P. falciparum predominating in Africa, New Guinea and Haiti. P.vivax is more common in the Indian sub-continent and Central America with the prevalence of these two infections roughly equal in Asia, Oceania and South America. Whereas, P. malariae is found in most endemic areas, especially sub-Saharan Africa, but less frequently. P. ovale is relatively unusual outside Africa although some cases are now being identified in others regions (e.g. Southern States of India) (WHO, 1989). Additionally " imported”
4 cases of malaria may present in any country, so called "airport malaria” has now been identified in a number of countries over the world. (Freedman, 1992)
In nature malaria is transmitted from person to person through the bite of infected female Anopheles mosquitoes of which there are hundreds of species. (Ponnuduari et al., 1988). The Anopheles belongs to the order of Diptera; sub-order Nematocera, family
Culicidae, Sub-family Culicinae and tribe Anophelini and the genus Anopheles. There are about 400 species of Anopheles mosquitoes throughout the world, but only some 60 species are vectors of malaria under natural conditions; of these some 30 species are of major importance (Gordon & lavoipierre, 1969).
Malaria can be transmitted by other relatively infrequent ways for example by inoculation of blood from an infected person to a healthy person (blood transfusions or using contaminated needles and syringes). By this way the asexual blood stages continue to develop in their own periodicities in the peripheral blood, producing attacks of fever in the recipient, but pre-erythrocytic and exo-erythrocytic schizonts are not formed in liver, as these forms originate from sporozoites inoculated by mosquito. Malaria transmitted by inoculation of blood is relatively easy to cure and relapses do not occur.
P. falciparum infection transmitted by this way can be fatal (Hang et al., 1995).
Furthermore, malaria can be transmitted congenitally, this can occurs, especially, when a mother is non-immune (Boukari et al., 1991).
. Anopheles mosquitoes are essential for development, multiplication and spread of plasmodia. Therefore, any area harboring Anopheles mosquitoes may be at risk for malaria transmission Specific environmental conditions optimal for anopheline mosquito vector and parasite development include temperature between 20° and 30°C and a mean relative humidity of 60%. The sporogony phase requires temperatures between 16° and
5 33°C. High relative humidity increases mosquito life span, thereby increasing the probability of mosquitoes becoming infective. Areas with high rainfall have increased malaria incidence because of an increase in breeding sites. The accompanying high humidity increases survival rates of female anopheline mosquitoes. Elevation, along with cooler temperatures and lower humidity, is also a factor as transmission rarely occurs above 2000-2500 meters. (Charlwood & Billingsley, 2000).
Malaria in a community could either be stable or unstable, and is classified as stable when there are high repeated infections in the presence of a constant transmission and the population acquires high degree of partial immunity with no epidemics occurring. On the other hand it is classified as unstable in areas where there are marked changes in transmission from one season to anther and from one year to the next. (WHO,
1996).
There are terms used to define the increasing levels of prevalence of malaria as estimated by surveys of spleen rates in particular age group. According to endemicity malaria is grouped into hypoendemic, mesoendemic, hyperendemic and holoendemic.
The lowest level is described hypoendemicity, and highest level is described as mesoendemicity, hyperendemicity, and holoendemicity. Holoendemic and hyperendemic malaria are found in areas of stable transmission whereas mesoendemicity and hypoendemicity are found in areas of unstable transmission. (Smith et al., 1993).
6 Figure1. Distribution of Malaria parasites over the world:
7 1.5 The life cycle of malaria:
The life cycle of malaria is complex with developmental stages and corresponding symptoms differing according to the Plasmodium species involved.
Sporozoites, the infective stage of plasmodia, are injected from the salivary glands of infected mosquitoes (definitive host) during feeding. Following inoculation, the sporozoites disappear from the blood within 30 minutes. Many are destroyed by white blood cells, but some of them reaches different organs and tissue but they succed to develop further only those, which enter liver cells where tissue (pre-erythrocytic) phase started. (Joklik et al., 1980).
1.5.1 Pre-erythrocytic phase:
Sporozoites that enter liver cells multiply asexually in a process called tissue schizogony. Thousands of uninucleate merozoites form, displacing the nucleus of the liver cell, but causing no inflammatory reaction in the liver. Eventually, invaded liver cells rupture, releasing thousands of merozoites into the bloodstream. This occurs 6 - 16 days after initial infection depending on the infecting Plasmodium species.(Lennette et al.,1985). All infections due to P. falciparum and P. malariae have a single exoerythrocytic form. All infected liver cells parasitized with P. falciparum and P. malariae rupture and release merozoites at about the same time. In contrast, P. vivax and
P. ovale have pre-erythrocytic and exo-erythrocytic forms. The primary type develops, causes liver cell rupture, and releases merozoites just as described for P. falciparum and
P. malariae. The other form, which develops concurrently, is known as the hypnozoite.
Sporozoites that enter liver cells differentiate into hypnozoites that remain dormant for weeks, months, or years. At some future time, the hypnozoites activate and undergo
8 exoerythrocytic schizogony, forming a wave of merozoites that invade the blood and cause a delayed case or a clinical relapse (Oh et al., 2001).
1.5.2 Erythrocytic phase:
Released merozoites invade red blood cells, where they develop into trophozoites. After a period of growth, the trophozoite divide and develop eventually forming a number of merozoites in each red blood cell depends on the species of plasmodia. When this process is completed, the host red blood cells rupture, releasing mature merozoites. The symptoms associated with malaria occur at this point. The merozoites then invade fresh erythrocytes and another generation of parasites develops in the same manner. This process occurs repeatedly during the course of infection and is called erythrocytic schizogony. The length of this development cycle differs according to the species of parasite, varying from 48 hours in vivax, ovale and falciparum malaria, to 72 hours in P. malariae infection. In the early stages of infection there is no characteristic periodicity as groups of parasites develop at different times. The febrile episodes caused are inconsistent. Later, the erythrocytic schizogony development cycle becomes synchronized, and the febrile paroxysms become more consistent. Some merozoites differentiate into sexual stages (macrogametocyte and microgametocyte) and develop in invaded red blood cells (Manson-Bahr & Apted, 1985).
1.5.3 Vector phase (Sporogony):
Anopheles mosquitoes feeding on infected hosts ingest sexual forms developing in red blood cells. The macrogametocytes and microgametocytes mature in the mosquito’s stomach and combine forming a zygote that undergoes mitosis. The products of mitosis
9 are ookinetes, which force themselves between the epithelial cells to the outer surface of the stomach, and form into small spheres called oocysts. The oocysts enlarge as the nucleus divides, eventually rupturing and releasing thousands of motile sporozoites into the body cavity. The sporozoites migrate to the salivary glands, making the female mosquito infective. The vector phase (Sporogony) is complete in 8- 35 days depending on species and environmental conditions (Brooks et al., 1998).
10 Figure 2. Life Cycle of Malaria parasites:
11 1.6 Pathogenesis of malaria:
The most serious form of malaria disease is caused by P. falciparum, mainly due to the erythrocytic schizogony-taking place in deep capillaries of internal organs such as brain, heart, spleen, intestine, bone marrow, lungs and placenta. And also due to the changes on the surface of the parasitized red blood cells especially that containing mature forms of the parasites in effect making them "sticky", which cause the cells adhere one another and to the cells that lining the walls of capillaries. This leads to the sequestration of infected red blood cells in the capillaries of internal organs, with subsequent congestion, hypoxia, and blockage and sometimes ruptures of small vessels
(Miller, 2002).
Clinically falciparum malaria is divided into uncomplicated, severe and complicated.
Severe and complicated falciparum malaria are associated with serious conditions, which may lead to death, particularly in non or semi immune patients such as pregnant women and children below 5 years of age. One of the most common complications is anaemia, which can be severe and occurs rapidly, particularly, in young children.
Anaemia in malaria is mainly due to mechanical distraction of parasitized red blood cells, parasitized red cells also lose their deformability and are rapidly phagocytosed and destroyed in spleen (Weatherall & Abdalla, 1982). Cerebral malaria, it is the commonest cause of death in falciparum malaria. Manifestations of cerebral malaria are caused by microvascular obstructions that prevent the exchange of glucose and oxygen at the capillary level, hypoglycemia, lactic acidosis and high-grade fever. These effects impair brain function, yet cause little tissues damage in most cases, as rapid and full
cerebral recovery follows prompt treatment. Ten to twelve percent of patients surviving
12 malaria has persistent neurologic abnormalities upon hospital discharge (Beare et al.,
2003). Renal failure due to acute tubular necrosis is a common complication of severe falciparum malaria in non-immune person. Acute tubular necrosis in severe falciparum infections is caused by two mechanism: renal tubules become clogged with haemoglobin and malaria pigment released during massive haemolysis, and microvascular obstruction causes anoxia and glucose deprivation at the renal capillary or tissue level (Mehta et al.,
2001). Pulmonary edema, this complication is rare but often fatal, acute pulmonary edema can develop rapidly and is associated with excessive intravenous fluid therapy.
Fast labored respiration, with shortness of breath, a non-productive cough and physical findings of moist rales and rhonchi present (Boulos et al., 1993). Hypoglycemia, commonly seen in falciparum malaria, is due to increase glucose consumption by parasitized red blood cells. In addition, quinidine or quinine may stimulate insulin secretion, causing clinically significant hypoglycemia (Shalev et al., 1992). Black water fever, this is rare but acute condition, in which there is a rapid and massive intravascular haemolysis of both parasitized and non-parasitized red cells, resulting in haemoglobaemia, haemoglobinuria, and fall in haemoglobin. It is accompanied with high fever, vomiting, jaundice, and the urine appears dark red to brown-black due to the presence of free haemoglobin. This condition is often fatal due to renal failure
(Delacollette et al., 1995). Algid malaria, it is falciparum malaria attacks characterized by rapid development of hypo tension and impairment of vascular perfusion, the temperature falls rapidly and the patient may become delirious. Symptoms of generalized vascular collapse and shock develop quickly (Bygbjerg & Lanng, 1982).
Dysenteric malaria, is an uncommon but extremely serious complication of falciparum malaria characterized by abdominal pain, nausea, vomiting and upper gastrointestinal
13 bleeding, which may be related to focal ischemic changes in the intestinal wall capillaries bed (Kochhar et al., 1990).
1.7 Immunity to malaria:
After repeated infections, people who live in regions where malaria is prevalent develop a limited immunity to the malaria. This partial protection does not prevent people from developing malaria again, but does protect them against the most serious effects of the infection. So, they develop a mild form of the disease that does not last very long and is unlikely to be fatal (Smith et al., 2002).
Most of the deaths and severe illnesses caused by malaria occur in children below
5 years of age and pregnant women. Children below 5 years are vulnerable because they have had fewer infections and have not yet built up immunity against the parasite.
Pregnant women are more susceptible to malaria because the immune system is somewhat suppressed during pregnancy. Moreover, malaria parasite uses a specific molecule to attach to the tiny blood vessels of the placenta. After exposure to this molecule during her first pregnancy, woman’s immune system learns to recognize and fight against the molecule. This phenomenon makes a woman, particularly, vulnerable to malaria during her first pregnancy, and somewhat less susceptible during subsequent pregnancies (Baird, 1995).
Some people have genetic traits that help them resist malaria by preventing the parasites from growing and developing normally, even in people who are infected with malaria for the first time, sickle cell anemia and thalassemia are two inherited blood diseases linked to malaria resistance. Various sickle cell or thalassemia genes are
14 widespread among people in Africa, Mediterranean region, Middle East, India, and
Southeast Asia (Astolfi et al., 1999; Bayoumi, 1987).
Another genetic condition that results in an increased resistance to malaria is ovalocytosis. In ovalocytosis, a protein found in the membrane of red blood cells is abnormal, causing these cells to have an oval shape. This trait, which is common in
Southeast Asia and the Pacific Islands, causes chronic anemia, but protects people from developing cerebral malaria. Finally, P. vivax cannot infect people whose red cells lack the Duffy antigen, a protein that is usually found on the surface of red blood cells. This trait, known as Duffy negativity, is common in people of African ancestry and causes no apparent health problems (Foo et al., 1992; Miller et al., 1976).
1.8 Diagnosis of malaria:
In malaria patients, early and accurate diagnosis is the keys to effective disease management and addressing morbidity and mortality due to malaria. (Dorsey et al.,
2000 ; WHO, 1993). However, malaria is commonly diagnosed clinically, nevertheless, laboratory confirmation should always take place whenever there is a possibility for that.
Clinical diagnosis, the most widely used approach, is unreliable because symptoms of malaria are very non- specific. (Van der Hoek et al., 1998). Rapid diagnostic tests
(RDTs) are another recent diagnostic approach introduced for malaria diagnosis. (Mason et al., 2002).
1.8.1 Clinical diagnosis:
Clinical diagnosis is the most widely used approach. It has been the only feasible one in many situations, particularly in rural areas and at the periphery of the health care
15 system where laboratory support to clinical diagnosis does not exist (Bassett et al.,
1991). Symptoms commonly associated with malaria include fever, mild to moderate malaise, fatigue, muscle aches, backache, headache, dizziness, loss of appetite, nausea, vomiting, abdominal pain and diarrhea. (Genton et al., 1994; Gilles, 1984). These various signs and syndromes caused by malaria are resulting from the pathological and physiological involvement of certain organs including brain, liver and kidney. (Clark &
Schofield, 2000). However, the symptoms of malaria are very non-specific and overlap with those of other febrile illnesses such as viral infections, hepatitis, bacterial infections
(typhoid fever, brucellosis, tuberculosis, meningitis, tonsillitis, wound infection, septicaemia, respiratory tract and urinary tract infections), and also protozoan infections such as visceral leishmaniasis and trypansomiasis. Therefore, diagnosis of malaria based on clinical grounds alone is unreliable, and when possible should be confirmed by laboratory tests. (Hanscheid, 2003).
1.8.2 Microscopic diagnosis:
Light microscopy is the conventional method for confirmation of malaria diagnosis (Chiodini & Moody, 1989). Careful examination of a well prepared and well stained blood smear remains currently the “gold standard” for detecting and identifying malaria parasites. In most settings, the procedure consists of: collection of a finger-prick blood sample; preparation of thick and thin bood smears and staining of the smear with
Giemsa stain and examining the smear under light microscope using the 100X oil immersion objective for the detection of malaria parasites (Thakor, 2000). Microscopy offers many advantages; it is sensitive when performed by skilled and careful technicians, it can detect the parasite at density level as low as 5– 10 parasites per µl of
16 blood. Under general field conditions, however, the detection capabilities of a typical microscopist might be more realistically placed at 100 parasites per µl of blood (Thellier et al., 2002). It is informative, when parasites are found, they can be characterized in terms of their species (falciparum, vivax, ovale, or malariae) and of the circulating stages (trophozoite, schizont, gametocytes). In addition, the parasite densities can be quantified (from ratio of parasites per number of leukocytes or erythrocytes). Such quantifications are needed to demonstrate hyperparasitaemia (which may be associated with severe malaria) or to assess parasitological response to chemotherapy (Van den
Ende et al., 1998). Microscopic diagnosis is relatively inexpensive. Cost estimates for endemic countries ranges from 0.12 US $ to 0.4 US $ per slide examined. (Bualombai et al., 2003). It is a general diagnostic technique that can be shared with other disease control programmes, such as those against leishmaniasis, filariasis, schistosomiasis or tuberculosis and finally it can provide a permanent record (the smears) of the diagnostic findings and be subject to quality control (Collier & Longmore, 1983).
However microscopy suffers from three main disadvantages. It is labour-intensive and time-consuming, normally requiring at least 30 minutes from specimen collection to result. It is exacting and depends absolutely on good techniques, reagents, and microscopes and, most importantly, well trained and well supervised microscopists.
Unfortunately these conditions are often not met, particularly at the more peripheral levels of the health care system. In these circumstances, microscopic diagnosis risks becoming an unreliable tool that uses up scarce resources for doubtful results and there are often long delays in providing the microscopy results to the clinician, so that decisions on treatment are often taken without the benefit of the results (Igbal et al.,
2002).
17 The Microscopic demonstration of malaria parasite may be accomplished by several methods, from the old and simple (but still golden standard) direct microscopic observation of stained blood specimens to the more recent and sophisticated concentration and staining techniques (Quantitative Buffy Coat, acridine orange method)
(Kawamoto, 1991).
1.8.2.1 Conventional thick and thin blood smears:
Microscopic examination of blood smears is still the cornerstone of malaria diagnosis since 1880, when the French Army surgeon Alphonse Laveran first observed and described malaria parasites in human blood in Algeria (Congpuong et al., 2001). At the end of the 19th century, it was recommended that the direct microscopic observation of malaria parasites be done in fresh unstained blood specimens flattened between a slide and a cover slip to appreciate the mobility of the parasite. It was then soon recognized by Romanowsky (1891) that the fixation and staining of blood specimens permitted a more accurate definition of the different stages and species, even though mobility was lost. The current standard methodology of Giemsa stained blood thick smears for the diagnosis of malaria dates back to 1903. (Adam & Maegriath, 1966).
Though simple in principle, the microscopic observation of stained blood specimens requires specific instrumentation and reagents (microscope, microscopic slides, pricking needle, staining reagents, water, electric or solar light, etc) and a trained professional to perform a correct steps of diagnosis. (Lema et al., 1999).
18 1.8.2.1.1 Blood drawing technique:
Since parasite concentration is fairly constant in internal and peripheral blood, it is routine to draw the blood by pricking a finger (or the heel in young infants) with a sharp sterile needle. The best site to prick is the lateral side of the third phalanx of the second or third finger of the left hand (unless the patient is left handed). After accurate cleaning with spirit- moisted cotton, of the finger, it will be pricked and the first drop of blood must be removed with cotton. The following drops of blood will be obtained by gentle squeezing of the finger. Then the blood will be collected on a glass slide that is lowered to touch the top of the drop. (Joss, 1999). It is to be stressed that the glass slides are to be carefully cleaned before use with clean water and detergent followed by 95% ethyl alcohol polishing to every trace of grease that may spoil the results. Slides, particularly in warm tropical climates, should be stored packaged in a dry atmosphere to prevent contamination with dust and mould. Blood obtained by syringe for routine examination purposes may also be used to prepare specimens for malaria diagnosis, preferable before anticoagulant is used. (Cheesbrough, 1999).
1.8.2.1.2 Preparation of the smears:
The microscopical observation of malaria parasites is optimal when parasites are fixed and observed in their natural location within red blood cells after appropriate staining. This is best accomplished with the thin smear preparation technique. (Day,
2003). Unfortunately, thin smear has a low sensitivity (100- 200 parasites/m l of blood) and is thus inadequate for low parasiteamia infection. An adequate parasite concentration method is obtained by osmotic lysis of the red blood cells releasing the parasites, as is the case with the thick smear preparation technique, the sensitivity of
19 which is then increased (about 10 parasites/m l of blood). Experienced technicians may also prepare thick and thin smears on the same slide, even though particular attention is to be paid in this case to avoid fixation of the thick smear. (Houwen, 2002). The methodology of the thin smear is as follow; one end of the slide is allowed to touch the top of the blood drop on the patient’s finger. Only the top of the drop should come into contact with the slide. The quantity of blood to be transferred to the slide should not exceed 1.5ml, usually corresponding to a diameter of 3- 4 mm. The edge of a second slide (or a cover slip) is then laid on the drop of blood that will spread on the entire line of contact between the two slides. The second slide, steadily held by the technician to form a 45 ° angle with the original slide, is then moved to the opposite end of t he slide to which the drop was originally located. In the well-prepared thin smear, the blood smear should end with multiple tails not touching the edges of the slide. Red blood cells should be visible one by one without overlapping. Abnormally thick slide may be the result of an exaggerated volume of blood or of an angle larger than 45 °. (Song et al.,
2000). On the other hand on the thick smear, the volume of blood twice to thrice the one used for a good thin smear is needed for the preparation of a thick smear (3.0 to 4.0m l).
This is usually accomplished by touching three times the top of the blood drop at the top of the finger to obtain a triangle on the slide. The blood is then gently mixed for 20- 30” using the corner of a second slide to defibrinate the blood and to obtain a round smear of about, 1cm in diameter. The thickness of the obtained smear should allow the reading of the printed text of a usual newspaper through it (Cheesbrough, 1992). Thin and thick smears must be allowed to dry in the air protected from dust, or actively dried by waving it. It is important to avoid abrupt exposure to heath (fire, sun light) that may lead to fixation and fissure the preparations. Thin smears must both processes of fixation and
20 staining, while thick smears must not be fixed to allow haemolysis of red blood cells and consequent dehemoglobinization. It is to be noted that fixation may also occur spontaneously (auto fixation) within time (7 to 15 days, varying with humidity and temperature of the atmosphere). It is then important to process thick smear as soon as possible to permit complete dehemoglobinization. (Coombs & Sandosham, 1975).
Fixation may be achieved by heat and alcoholic solutions. Methanol (methyl alcohol) is the most widely used fixative for malaria thin smears. Contact with methanol should be maintained for10- 20 seconds. If both thin and thick smears are on the same slide, it is mandatory to avoid the contact of methanol (or even of its fumes) with the thick smear to avoid fixation. (WHO, 1980).
1.8.2.1.3 Staining procedures:
The discovery by Laveran in 1880 that the disease of malaria was caused by parasites which developed in erythrocytes was made by examination of fresh unstained smears, therefore Laveran and others scientist at that time, only saw the vague outline of the living organism (Shute & Margon, 1966). The first attempts to colour blood smears was made in 1879 by Ehrlich when he described the use of 'neutral' dyes (mixtures of acidic and basic dyes) for the differentiation of cells in peripheral blood smears. In 1891
Romanowsky and Malakowsky independently developed a method, which used mixtures of Eosin Y and ‘ripened’ Methylene Blue that not only differentiated blood cells, but also demonstrated the nuclei of malarial parasites. A number of 'ripening' (oxidation or polychroming) techniques were investigated by different groups (Unna 1891, Nocht
1898) but the aqueous dye solutions produced were unstable and precipitated rapidly.
Subsequently, methanol was introduced as a solvent for the dye precipitate (Jenner 1899,
21 England; May and Grünwald 1902, Germany) and techniques were developed that utilised the fixative properties of the methanolic solution, prior to aqueous dilution for staining (Leishman 1901, England; Wright 1902, Germany). Giemsa (1902) further improved these techniques by using more controlled methods of oxidation with measured amounts of known dyes, and adding glycerol to the methanol solvent to increase the solubility and stability of the dyes. (Power, 1982).
Traditional Romanowsky-type dyes are produced by oxidation (polychroming) of Methylene Blue in aqueous solution, using heat and alkali. The resulting solution contains a mixture of Azure A, Azure B, Methylene Violet and Methylene Blue
(aqueous Romanowsky stain). A measured quantity of Eosin Y is then added to produce a 'neutral' dye. The precipitate formed is dissolved in methanol, or a mixture of equal volumes of methanol and glycerol, to produce a stable stock solution (alcoholic
Romanowsky stain). Working solutions are prepared by diluting the stock solution with an aqueous buffer, to allow ionisation of the dyes. (Field & Sandosham, 1964).
The aqueous type of Romanowsky stains that are in use up to date are Field’s stain and J.S.B (Jaswart, Singh, & Bhattacharji) stain, Field's stains is only suitable for thick blood smear while J.S.B.may be used for thick and thin blood smears. Brilliant results can be obtained with the aqueous stain but their great advantage is in the rapidity with which the blood smears may be stained, staining with Field is completed in a few seconds and J.S.B. in little more than that also the aqueous stains are cheap to produce.
But the main disadvantages of the aqueous stains are that they are not adapted to mass staining techniques, each blood smear requiring individual staining and the all-aqueous staining solutions are susceptible to the growth of mold, particularly in tropical countries. The moulds become rapidly prolific and are deposited over the blood smears
22 stained in solution, ruining the smear from the point of view of malaria parasite. To prevent this the stain should be filtered or better still boiled and filtered. (Makler et al.,
1998). The alcoholic types of Romanowsky stain are those in which the azures have been dissolved in methyl alcohol, sometimes with addition of glycerol. The azures do not dissolve readily in methyl alcohol and are usually ground into fine powder to aid solution. The addition of glycerol to the solution aids its keeping properties as Giemsa stain, which is always made up with a proportion of glycerol, will be keep indefinitely.
Leishman and Wright stains are usually prepared from methyl alcohol only and in warm climate rarely keeps for more than a few months, sometimes for only a few weeks. It may be that moisture plays an important part in this and that if moisture from atmosphere is prevented from reaching the stock solution of the stain, it might keep for a considerably longer time. In practice, particularly in warm, humid climate, this is difficult to achieve. The addition of glycerol to the solution reduces the affinity of methyl alcohol and this may be why glycerol inated staining solutions have such good keeping properties. (Pandey et al., 1995).
From all types of Romanowsky stains, Giemsa stain still is the 'gold standard ‘ stain for staining blood smears to demonstrate the malaria parasites. It can be used with thick blood smears as well as the thin blood smears, also it can be used to stain mass of slides at the same time, whereas in the all other types of Romanowsky stains each slide needs treated individually. (Igbal et al., 2003)
1.8.2.1.4 Microscopic examination:
When the smears completely dry, a drop of immersion oil is applied to the blood smear and they are examined by using the objective lens. ×100. The thick blood smear is
23 examined firstly to detect the presence of malaria parasite and then the thin blood smear examined is to identify the species of plasmodia. (Ohrt et al., 2002).
1.8.2.2 Quantitative buffy coat (QBC) test:
It is a new method for identifying the malarial parasite in the peripheral blood. It involves staining of the centrifuged and compressed red cell layer with acridine orange and examination it under UV light source. (Lowe et al., 1996).
The QBC tube is a high-precision glass hematocrit tube, pre-coated internally with acridine orange stain and potassium oxalate. It is filled with 55-65 microliters of blood from a finger, ear or heel puncture. The tube is centrifuged at 12,000 rpm for 5 minutes.
The components of the buffy coat separate according to their densities, forming discrete bands. The leukocyte and the thrombocyte cell bandwidths and the top-most area of red cells are enlarged to10 times normal. The QBC tube is placed on the tube holder and examined using a standard white light microscope equipped with the UV microscope adapter, an epi - illuminated microscope objective. Fluorescing parasites are then observed at the red blood cell / white blood cell interface. (Htut et al., 2002; Keiser et al., 2002).
1.8.2. 3 Saponin lyzing technique:
Detection of parasitized erythrocytes in the blood is a requirement in the diagnosis of malaria by the standard blood-smear technique, but parasiteamia in patients is often scanty and sometimes difficult to detect by this technique. The poor sensitivity of this method can be attributed, at least in part, to the small quantity of blood examined. Thick smear examination for malaria is often hampered by the presence of cellular debris
24 obscuring parasites. This is a major problem for diagnostic laboratories that do not have a high exposure to material identification. Smears that are relatively free of cellular debris, allowing easier identification of parasites are an obvious advantage. Researchers to liberate malarial parasites for harvesting from infected erythrocytes have used saponin. It has also been used for thick smear preparations for diagnosis, but has not gained widespread acceptance, possibly due to the persistence of cellular debris inherent in the technique. (Gleeson, 1997). The method for thick smear examination has been modified by the inclusion of a centrifugation step to remove cellular debris. The blood is suspended in a 0.015 % saponin solution and incubated for 30 min to give time for red blood cells to lyses. Then the suspension is centrifuged in 2000 rpm for 5 min. The supernatants discarded, thin smears made from pellets and examined under microscope.
(Orjih, 1994).
1.8.3 Rapid diagnostic tests (RDTs):
These tests are based on the detection of antigens derived from malaria parasites in lyzed blood, using immunochromatographic methods. (Moody, 2002; Playford &
Walker, 2002). Current interest in rapid diagnosis is focused primarily on detection of histidine- rich protein II from P. falciparum and parasite specific lactate dehydrogenase from all species of malaria. Current tests use immunocapture techniques to detect malaria antigens. (Piper et al., 1999; Beadle et al., 1994). However RDTs are very easy to perform and the result obtained within 15 minutes but compared to microscopy, the main disadvantages of currently available RDTs are: lack of sensitivity at low levels of parasitaemia; inability to quantify parasite density; inability to differentiate between P. vivax, P. ovale and P. malariae, as well as between the sexual and asexual stages of the
25 parasite; persistently positive tests (for some antigens) in spite of parasite clearance following chemotherapy; and relatively high cost per test. (Wongsrichanalai et al., 2003;
Dietze et al., 1995).
1.8.4 Deoxyribonucleic acid (DNA) probes and Polymerase Chain Reaction (PCR):
Conventional microscopy is usually satisfactory for the large majority of clinical situations, but may be inadequate for very low parasiteamia; the possibility that modern molecular biologic techniques overcome these drawbacks has thus been explored
(Morassin et al., 2002; Srinivasan et al., 2000). PCR assays to detect Plasmodium DNA in human blood by using different approaches to increase sensitivity and specificity have been described. (Patsoula et al., 2003; Haiana et al., 2001) Nevertheless, a number of drawbacks hamper the use of the PCR technique for the diagnosis of malaria infection not only in the field but also in well equipped laboratories in industrialized countries, the most important of which being contamination, yielding false positive results, and high costs. of contamination of PCR with previously amplified products.(Chaorattnakawee et al., 2003; Rubio et al., 2001).
1.8.5 Detection of Plasmodia specific antibodies:
The presence of the malaria parasite in the patient’s organism elicits the production of a wide range of antibodies, both specific against Plasmodia antigens and non-specific against leukocytes, red blood cell, rheumatoid factor, etc. (Dion, 1995; Ray,
1984). The first serological test to be used for malaria antibodies was immunofluorescence (IFAT), which may give quantitative results for both G and M specific immunoglobulins. Its specificity and sensitivity largely rely on the laboratory
26 technician’s expertise .Results higher that 1:20 are considered to be positive and high titers (>1: 200) probably reflect recent infection. The indirect haemoagglutination test
(IHA) is simple and suitable for field studies, but its sensitivity and specificity are poor.
The enzyme- linked immunosorbent assay (ELISA) has similar sensitivity and specificity characteristics than the IFA test, but the interpretation of the results may be better standardized. For research purposes, radio immunoassay (RIA) is sometimes used but needs well-equipped research laboratories and personnel. (Schapira et al., 1994;
Ferreira, 1990).
1.9 Chemotherapy of malaria:
Proper use of antimalarial drugs is based on the knowledge of their effects on the parasite at various stages of the life cycle. (WHO, 1973). Suppressive or protective therapy (chemo prophylaxis), this implies that the drugs are used before infection occurs or before it become evident, with aim of preventing either the occurrence of the infection or any of it symptoms. Curative (therapeutic), therapeutic use of the drugs refers to the action on the established infection and comprises the treatment of acute attacks, and radical treatment. (Christopher, 2003). There are several chemical groups of antimalarial compound in general use, for practical purposes, the drugs belonging to the most nine important groups are given here under their international nonproprietary names; quinine, mepacrine, chloroquine, amodiaquine, proguanil and chlorproguanil, pyrimethamine, sulfones and sulfonamides, Quinolinemethanols, tetracycline and clindamycin. (Philip,
2003). Resistance to some of these drugs was noted early but was of little clinical significance until chloroquine resistance appeared in strains of P.falciparum from South
America and Southeast Asia in the 1950s, subsequently spreading to the most areas of
27 intense malaria transmission. (Sidhu et al., 2002). The most important reasons for chloroquine resistance malaria is incorrect use of the drugs particularly when taken by non or semi immune person, when chloroquine is taken to relive the symptoms of malaria attack but not enough to kill all parasites, the stronger parasites survive and multiply. It also though that selective pressure favoring naturally occurring resistance mutation increase in areas of intense malaria transmission. Resistant strains may also developed in area where a small drug doses are used in prophylaxis. (Foote & Cowman,
1994). P. falciparum strains that are resistant to chloroquine are often resistant to others synthetic antimalarial as well , and quinine has once more remerged as mainstay of treatment in such cases . (Reed et al., 2000). Chloroquine resistant has also made its appearance in P.vivax in Oceania but as yet in limited clinical significance.
(Murphy, 1993). There are three grades of resistance are recognize, ranging from RI, in which administration of the drug achieves apparent clinical cure only to be followed by recrudescence, to RIII, in which therapy seems completely ineffective (Markell et al.,
1999).
Many older drugs are still effective in some areas of the world. Of these, chloroquine is the most desirable because of its low cost, high effectiveness, and few side effects. But
P. falciparum has developed resistance to the drug, making it ineffective for use in
Africa and other areas. Resistance exists also in other drugs include atebrin,
doxycycline, mefloquine, and quinine. (Jennifer, 2003).
Some alternatives to current malaria drugs are now in the works. These include a new formula of the new, long-acting prophylactic drug tafenoquine that is suitable for children, artemisinin derivatives (products derived from a Chinese herbal remedy).
Although artemisinins reportedly have a high failure rate when used alone, they may be
28 effective when combined with another drug such as mefloquine. Malaria research is moving away from single-drug treatment, and treatment recommendations now emphasize combination therapies. The drawback to these combination drugs is that their cost may be up to 10 times that of current antimalarial agents. (Jennifer, 2003)
1.10 Malaria in the Sudan:
1.10.1 Transmission:
Endemicity of malaria ranges from hypo- endemic in northern Sudan, meso- endemic in the center to hyper or holo-endemic in central and southern parts of Sudan.
This considerable variation in transmission is due to that, in northern states the breeding of the vector limited to the river bed in the remaining pools created by the receding flood. In the central states (irrigated areas), transmission season usually in the rain season (June to October), and this followed by another transmission season during irrigation of winter crops and vegetables, which extended from November to March. In southern Sudan states, malaria transmission is stable throughout the year, due to the availability of breeding places to vector all the year. Generally the malaria transmission reaches its maximum peak in the heavy rainy season with mild fluctuation; therefore malaria could be classified into unstable and high incidences of malaria and intensive transmission correlated with the heavy rain and Nile flood. Easy movement of population, displaced population during the natural disaster and war and the refugees from neighboring countries particularly in the East, could help in the spreading of the disease and new strains might imported, so strains resistant to drug was reported in recent years.(Hamad et al., 2002).P. falciparum is responsible for more than 90% of malaria cases. However, there is an increase in malaria cases caused by P.vivax outside
29 its classical zone, Anopheles' arabiensis is the principle vector of the malaria in the
Sudan. (El sayed et al., 2000).
1.10.2 Morbidity and mortality:
Sudan is the largest country in Africa, comprising more than 8% of the entire continent. The total population is estimated to be 35 million inhabitants, of whom 75% live in rural areas. In the Sudan, Malaria is a major health problem causing 7 - 8 million episodes annually, and 35,000 deaths every year due to malaria, it accounts for about
21% of all diseases seen at outpatient department in health facilities in the country, ranging from 20-40% and 30-40% of admissions. In terms of mortality, malaria accounts for 20% of all hospital deaths in the country. Whilst the whole population is at risk from the disease, there is higher incidence among pregnant women and children under five years of age. This results in complicated pregnancies, low birth weight and infant mortality. (UN, 2003).
1.10.3 Diagnosis of malaria:
Diagnosis of malaria in the Sudan based mainly on examination of stained blood smears under microscope and recently immunological detection (ICT) and the PCR are used. In a comparison between microscopic diagnosis and rapid tests, both are accurate, but only microscope can count the parasite number. In the Sudan, the performances of the laboratories are generally poor, including malaria microscopy. When’re- examining blood films, there is a high proportion of false positives. A survey conducted July2002-
Sept 2003 included 288 labs: about half of the microscopes are in bad condition with
30 20% needing replacement. In general, less than 50% of all the equipment are of acceptable quality. (Sudan Federal Ministry of Health, 2003)
1.10.4 Chemotherapy of malaria:
Chloroquine has been the antimalarial drug of choice for the past 50 years Malaria treatment is today in a critical stage of transition. Chloroquine resistance has reached unacceptably high levels in most malaria endemic countries, and this has been realised also in Sudan through 30 studies of chloroquine resistance completed over the last 5 years in various parts of the country. In Sudan, resistance was reported in Sinnar in
1983, Khartoum in 1983, and Gedaref in 1985. Therefore National Protocol for Malaria treatment options in 1998 for simple malaria: first line: chloroquine second line: SP
(Fansidar ) third line: quinine, mefloquine and halofan . For severe malaria quinine / artemether. For prevention: chloroquine, mefloquine or proguanil. A high prevalence rate of drug resistance is noticed by health professionals and the public, some areas in the Sudan have a low resistance rate against chloroquine and SP as monotherapies.
According to the recommendation of options against malaria in Sudan conference that held in Khartoum in October 2003, Sudan Federal Ministry of Health choice combination therapy as National Protocol for Malaria, first line: Artesunate + SP
(Fansidar) second line: Coartem for uncomplicated malaria, quinine / artemether for pregnant women and quinine for severe complicated malaria . (Sudan Federal Ministry of Health, 2003).
31 1.10. 5Control:
Sudan has a long history of malaria control activities dating back to pre-colonial times. The attempt at malaria eradication started 1954-64 with the introduction of DDT and it focused on vector control, residual spraying, environmental management and public education. The program had very limited success due to managerial, technical and financial constrains. In 1956, Malaria Eradication Center was founded in Sennar with assistance of WHO, the center was started as a malaria control project and extended for eradication programme and training courses for different health workers. The establishment of the Blue Nile Health Project (BNHP) in 1978, with support from WHO,
World Bank, Kuwait, Japan and USA, led to successful control of malaria for 10 years with the prevalence of the disease coming down to <1%. Unfortunately, external funds stopped, leading to discontinuation of control operations in 1989. In 1995, the Blue Nile
Research and Training Institute (BNRTI), started instead of BNHP with a new approach, the BNRTI founded in joined between University of Gezira, the WHO and the Federal
Ministry of Health. The BNRTI formulated a curriculum and programmes for research and training for awarding higher diploma in malariology for medical and health officers and established reference laboratory for malaria research in Wad Medani
Teaching Hospital. Now Malaria National Administration – Federal Ministry of health acts as a co-ordination authority on international health works concerning malaria, also one of its functions is to formulate a national strategy to control the disease and to exercise leadership in providing support for the implementation of the national policy.
(Mohamed-Nour, 1998).
32 Figure 3. Transmission of Malaria in the Sudan:
33 1.11 Justification:
The accurate diagnosis of malaria is the base of effective management, although, many new techniques have been developed for the laboratory diagnosis of malaria, nevertheless, in the Sudan microscopic detection of malaria parasites in stained blood smears is still commonly used method.
Microscopic diagnosis of malaria face some difficulties and is very frequently not accurate due to the general condition of the microscope, the quality of stain and other consumables, the techniques used for preparation and staining of blood smears and wrong practices of laboratory personnel (mainly in staining of blood smears). Fortunelty, all of the above-mentioned differences could be correlated by both qualification and training of laboratory workers.
1.12 Objectives:
1.12.1 General objective:
To assess different stains and staining procedures for microscopic detection of malaria parasites.
1.12.2 Specific objectives:
1- To assess the use of saponin lyzed venous blood for microscopoic detection of
malaria parasites.
2- To compare between the use of Giemsa and Fields stains for the diagnosis of
malaria.
3- To assess quality of three stocks of Giemsa stain prepared by different methods,
used different concentration for Giemsa working solutions, used distilled water
or tap water to dilute the stock stain and check the stability of working solution.
34 CHATER TWO
MATERIALS & METHODS
2.1 Study population:
A total of 203 patients referred by clinicians to the laboratory for microscopic confirmation of malaria were included in this study after obtaining their verbal consent.
Selected patients comprise both sexes and aged between 10-60 years.
2.2Study design:
This is a comparative study consisted of six parts and the protocol of each part were carried out separately.
2.3 Samples collection:
Five ml of venous blood were collected from each individuals into labeled containers containing sequestrene anticoagulant (EDTA). Immediately after collection, from each one of study population the following blood smears were prepared, 15 thin and thick blood smears on same slide, a thick smear and smear from saponin haemolyzed venous blood.
2.4 Techniques:
2.4.1 Giemsa stained smears:
Thick and thin blood smears were prepared for each individual. Thin blood smears were fixed by absolute methyl alcohol before staining. Then they were stained with 10% Giemsa stain (Nen Tech.Ltd-UK) (pH 7.2) for 10 min and left to dry.
35 Microscopic examination of stained blood smears was carried out using x100 immersion oil lens by three technicians with a same qualifications independently. According to
WHO at least 100 thick fields were examined before the blood smear was determined as negative. In positive thick blood smears density of infection (parasite count per µl of blood) was calculated. Thus, 200 white blood cells (WBC) were counted if less than 10 asexual stages were detected then counting continue tell reaching 500 WBC, counting at the same time the number of parasites (asexual stages) in each field covered, the numbers of parasites in 1 µl of the blood was then calculated using the following formula:
Parasites counted against 200 WBC × TWBC count
200
According to WHO the total white blood cell was considered as 8000/µl for each individual (WHO, 2000).
2.4.2 Saponin lyzed venous blood technique:
According to Orjih (1994), 1 ml of blood from each individual was dispensed in a plastic test tube containing 7 ml of 0.015% saponin lyzing solution(Seelze Hanover-
Germany), then blood was mixed gently in the lyzing solution. The suspensions were left for 30 min in order to give time for all red blood cells to be lyzed. Then the, haemolysate was centrifuged at 2000 rpm for 5 min. The supernatant was then discarded, and the pellets were washed once with 7 ml of lyzing solution and centrifuged at 2000 rpm for 5 min. Thin smears were prepared from the pellet and stained with 10%
Giemsa stain for 10 min. Smears were examined microscopically by the three
36 technicians using oil immersion lens. The results were recorded and compared to the results obtained by Giemsa smear
2.4.3 Field’s stain smears:
A thick blood smear was made from each individual of the study, stained with
Field’s stain (BDH Chemical Ltd-UK). The blood smear was dipped in solution (A) for
5 seconds, washed by dipping in clean water to remove excess of solution (A), then dipped into solution (B) for 2 seconds and then washed in clean water to remove excess of stain. The three-technician using oil immersion lens examined all smears microscopically and results were recorded and compare to the results obtained by
Giemsa smears.
2.4.4 Preparation of different Giemsa stains stock solution:
Three different batches of Giemsa stock stain were prepared, Stock I was prepared using equal amounts of glycerol and methyl alcohol, Stock II was prepared using 2 parts of absolute methyl alcohol to 1 part of glycerol in dissolving the compound of the stain, and Stock III was prepared by using 3 parts of methyl alcohol to 1 part of pure glycerol.
These stock stains were used to stain 120 thick and thin blood prepared in the same slide, of these 60 smears were positive for malaria parasites and the other 60 were negative blood smears. Then 10% stain working solution (pH 7.2) was prepared from each stock stain and blood smears were stained for 10 min using different batches of stain for the same blood. Then blood smears were examined under the microscope using oil immersion lens and the remarks about the quality of the stain in staining parasites and blood cells were reported. The examined blood smears were assessed and evaluated
37 by rule of thumb, according to quality of staining different components of blood smear, each blood smear was given a score depending on the staining of parasites (nucleus & cytoplasm), blood cells and background of thick blood smear. One mark was given for each parameter examined (parasite nucleus, parasite cytoplasm, nuclei of white blood cells in thick smear, white blood cells & red blood cells in thin smears and the background of the thick blood smear) that showing good stain quality and 2 marks wee given for that one showing bad stain. The positive blood smear was regarded as distinguishable (good quality of stain) when it scored 7 or less and undistinguishable blood smear scored more than 7 marks. For negative blood smears the distinguishable had 5 marks or less while undistinguishable blood smears scored more than 5.
2.4.5 Concentration of Giemsa working solution:
Three different concentrations of Giemsa working solution were prepared (from stock stain containing equal part of methanol and glycerol). The first concentration was
10% of working solution for staining for 10 min, the second was 15 % of working solution for staining for 5 min, and the last one was 20 % of working staining for 2 min.
The working solutions were used to stain 120 thick and thin blood smears prepared from the same blood. Of these 60 blood smears were known positives and the others 60 one were known negatives. The stock stains diluted to working solution by buffered solution pH 7.2 and used to stain the smears for 10 min. The smears were examined under the microscope using oil immersion lens and the remarks about the properties of the stain in staining parasites and blood cells were reported. The examined blood smears were assessed and evaluated by rule of thumb, according to quality of stain, each blood smear was given a score depending on the staining of parasites (nucleus & cytoplasm), blood
38 cells and background of the thick blood smear. One mark was given for each parameter examined (parasite nucleus, parasite cytoplasm, nuclei of white blood cells in thick smear, white blood cells & red blood cells in thin smear and the background of the thick blood smear) that showing good stain quality and 2 marks were given for that one showing bad stain. The positive blood smear was regarded as distinguishable (good quality of stain) when it scored 7 or less and undistinguishable blood smear scored more than 7 marks. For negative blood smears the distinguishable had 5 marks or less while
undistinguishable blood smears scored more than 5.
2.4.6 Dilution of stock Giemsa stain with distilled or tap water:
Two Giemsa working solutions were prepared (from stock stain containing equal part of methanol and glycerol) one by using distilled water and the other by tap water, respectively instead of buffered solution pH 7.2. These working solutions were used to stain 120 thick and thin blood (the smear were made in one slide), 60 of these smears were known positives and the others 60 one were known negatives. The smears were examined under the microscope using oil immersion lens and the remarks about the properties of the stain for staining parasites and blood cells were reported and compared with smears from the same blood stained with working solution diluted by buffered solution pH 7.2. The examined blood smears were assessed and evaluated by rule of thumb, according to quality of stain, each blood smear was given a score depending on the staining of parasites (nucleus & cytoplasm), blood cells and background of the thick blood smear. One mark was given for each parameter examined (parasite nucleus, parasite cytoplasm, nuclei of white blood cells in thick smear, white blood cells & red blood cells in thin smears and the background of thick blood smear) that showing good
39 stain quality and 2 marks gave for that one showing bad stain. The positive blood smear was regarded as distinguishable (good quality of stain) when it scored 7 or less and undistinguishable blood smear scored more than 7 marks. For negative blood smear the distinguishable had 5 marks or less while undistinguishable blood smears scored more than 5.
2.4.7 Stability of Giemsa stain working solution:
A working solution of Giemsa stain (10 %) was prepared (from stock stain containing equal part of methanol and glycerol) by using buffered solution pH 7.2 and used immediately to stain120 thick and thin blood (the smear were made on the same slide), 60 of these smears were known positives and the others 60 one were known negatives. The same working solution was used at intervals of, 1 hour, 2 hours, 3 hours,
4 hours, 6 hours, 8 hours, 10 hours and 12 hours to stain 120 smears (60 known positive
& 60 known negative at each time. The all smears were examined under microscope using oil immersion lens and the remarks about the properties of the stain for staining parasites and blood cells were reported. The examined blood smears assessed and evaluated by rule of thumb, according to quality of stain, each blood smear was given a score depending on the staining of parasites (nucleus & cytoplasm), blood cells and background of the thick blood smear. One mark was given for each parameter examined
(parasite nucleus, parasite cytoplasm, nuclei of white blood cells in thick smear, white blood cells & red blood cells in thin smear and the background of the thick blood smear) that showing good stain quality and 2 marks were given for that one showing bad stain.
The positive blood smear was regarded as distinguishable (good quality of stain) when it scored 7 or less and undistinguishable blood smear scored more than 7 marks. For
40 negative blood smears the distinguishable had 5 marks or less while undistinguishable blood smears scored more than 5.
2.5 Ethical consideration:
All participants of the study were told about the aims of the study and they were willingly to participate in present study. Very young and very old individuals were excluded from the study
2.6 Data analysis:
Data were analyzed using the Statistical package for Social Sciences SPSS
(SPSS/PC + version 10.05). The descriptive statistics were carried out using frequency.
Proportion was compared using McNemer and Kendall tests. T test was used for comparison of means (mean parasite count). Significance was determined at the 0.05% probability level in all analysis.
41 CHAPTER THREE
RESULTS
3.1 Giemsa stained blood smears:
A total of 203 patients referred by clinicians to the laboratory for microscopic examination of blood for malaria were included in the study. 127 (62.6 %) were males and 76 (37.4%) were females, their age ranged between 15 and 60 years and the majority of them (48.3%0) were between 30-39 years (Figure 4).
Microscopic examination of Giemsa stained blood smears (gold standard) revealed the presence of malaria parasites in 67 (33%) of them. P. falciparum was the only species identified in all examined blood smears. It was detected in 52 (40.9%) of males and in 14 (18.4%) of females.
As presented in table 1, out of 203 examined blood samples, malaria parasites
(Plasmodium falciparum) were identified in 67 (33%) of Giemsa stained blood smears, in 61 (30%) of Saponin lyzed blood smears and in 55 (27.1%) of Field's stained blood smears. The result showed that Giemsa stained blood smears determined the highest mean of parasite count 5964/µl, followed by Saponin lyzed blood smears (4660/µl) whereas using Field's stains smears a mean of 4459 parasite per µl of blood was determined.
Moreover, the result showed that the density of infection ranged between 120 and
46400 parasites per microlitre of blood and the smears stained with Giemsa stain showed marked capability over all techniques of the study to detect malaria parasites in low densities, 37.4% of positive smears with level of parasiteamia < 1000 parasites/µL of blood were detected by Giemsa stain whereas 34.3% detected by Saponin and only
28.3% of them detected by Field stain. (Figure 5)(Appendix1).
42 Figure 4. Age group distribution of the study population:
70
60
50
40
Number 30
20
10
0 10 19 20 29 30 39 40 49 50 60 Age group
Male Female
43 Table 1. Comparison between Giemsa, Saponin lyzed and Field’s stained thick blood smear in detecting malaria parasite and in determining the parasite count/µl blood:
Staining technique P-
Thick blood smear value Giemsa Saponin Field’s
Total number examined 203 203 203
Number positive 67 (33%) 61 (30%) 55 (27%) .002*
Mean Parasites count/µl of blood 5964 4660 4459 .000**
* Kendall’s test ** T. test.
44
Figure 5. Relationship between detection of malaria parasites according to level of parasiteamia in blood smears stained with different stains.
30 27 24 21 18 15
Number 12 9 6 3 0
00 00 70 <1000 130 -19000 01- >22000 01 1000-400040 7001=1000010001- 13001-16000160 19001-22000 Range of parasiteamia / microlitre of blood
Giemsa Saponin Field
45 3.2 Saponin lyzed venous blood technique:
Two smears were prepared from each of the 203 blood samples, a thick blood smear and Saponin lyzed venous blood smear, further smears were stained with 10%
Giemsa for 10 minutes and examined by the three technicians.
Study findings showed that of 203 samples examined, technician A detected malaria parasite in 66 (32.5%) of thick blood smears and in 60 (29.5%) of Saponin lyzed venous blood smears. Technician B detected malaria parasites in 63 (31%) and 58 (28.6%) of thick blood smears and Saponin lyzed venous blood smears, respectively. Of 203 blood samples examined by Technician C malaria parasites were found in 67(33%) of thick blood smear and in 61 (30%) of Saponin lyzed venous blood smear. Statistical analysis showed similar result between the two techniques for the three technicians (Table 2).
46 Table 2. Comparison between Giemsa stained-thick blood smears and Giemsa stained-Saponin lyzed venous blood smears for detecting malaria parasites examined by three technicians:
Positive Technicians No. Examined P- Thick blood smears Saponin lyzed value* blood smear
A 203 66 (32.5%) 60 (29.5%) .070
B 203 63 (31%) 58 (28.6%) .063
C 203 67 (33%) 61 (30%) .070
* McNemar test Exact Sig. (2 sided).
47 3.3 Comparison between Giemsa and Field’s stain in detecting malaria parasites:
As presented in table 3, out of 203 Giemsa stained thick blood smears examined independently by each of the three technicians, malaria parasites were detected in 66
(32.5%) of examined blood smears by Technician A, in 63 (31%) by Technician B and in 67 (33%) by Technician C. Considering the use of Field's stain the study findings revealed that of 203 Field’s stained thick blood smears examined by each of the three technicians, malaria parasites were detected in 55 (27.1%) of examined blood smears by
Technician A and Technician B, whereas technician C determined the presence of malaria parasites in 53 (26.1%) of them. Statistical analysis showed that malaria parasites were detected by the three technicians in significantly greater proportion when
Giemsa stained thick blood smears compared to those stained with Field's stain.
48 Table 3. Comparison between Giemsa and Field stained thick blood smears examined by three technicians in the detection of malaria parasites:
Positive Technicians No. Examined P. value* Giemsa Field
A 203 66 (32.5%) 55 (27.1%) .001
B 203 63 (31%) 55 (27.1%) .039
C 203 67 (33%) 53 (26.1%) .013
* McNemar test Exact Sig. (2 sided).
49 3.4 Preparation of different Giemsa stains stock solutions:
The quality of differently prepared Giemsa stock solutions was assessed in the present study. Thus, three differently prepared Giemsa stain stock solutions were used for staining 120 (60 known positive & 60 known negative) thick and thin blood smears were prepared on the same slide. According to the criteria mentioned in chapter two components of blood smears were classified as distinguishable or indistinguishable based on the quality of staining blood cells and the parasites. Microscopic examination of 120 thick blood smears stained with Giemsa stock solution 1 (standard stain) showed that 93.3% of the components of the blood smears were easily identifiable during microscopic examination. This is comparable with 92.5% and 90.8% identifiable components of blood smears stained with stock solution II and stock solution III, respectively. (Table 4). Further statistical analysis revealed no significant differences between the three stock solutions of Giemsa stain in staining blood smears.
50 Table 4. Comparison of the stain quality of thick & thin blood smears using different Giemsa stain stock solutions:
Giemsa stain stock solution
Thick blood smear Stock 1 Stock II Stock III
Number examined 120 120 120
Distinguishable components of smear 93.3% 92.5% 90.8%
P. value* (Compare to Stock I) 1.000 .581
* McNemar test Exact Sig. (2 sided)
51 3.5 The use of different concentrations of Giemsa stain working solutions:
Three different dilutions of Giemsa stain of 10%, 15% and 20% were used to stain
120 (60 known positive & 60 known negative) thick and thin blood smears on the same slide. As shown in table 5, according to the appearance of different components of blood smear after staining the quality of stain was evaluated. Thus, the study findings showed that after staining with 10% Giemsa stain 95% of the blood smear components were distinguishable, when 15% of Giemsa stain was used 76.7% of the smear components were distinguishable however when Giemsa stain concentration was increased to 20% only 9.2% of blood smear components were distinguishable.
Statistical analysis revealed significant superiority of using low concentration of
Giemsa stain in staining blood smears.
52 Table 5. Comparison between different concentrations (10%, 15% and 20%) of
Giemsa stain working solution in staining thick & thin blood smears:
Concentration of Giemsa stain working solution
Blood smear 10% 15% 20%
Number examined 120 120 120
Staining of smears (Distinguishable components) (%) 95% 76.7% 9.2%
P- value* (Compare to concentration 10%) .000 .000
* McNemar test Exact Sig. (2 sided).
53 3.6 Preparation of Giemsa stain working solution using different diluents:
Giemsa stain stock solution was diluted with phosphate buffered solution pH 7.2
(gold standard), distilled water and tap water. Each of the prepared working solutions was used to stain 120 (60 known positive & 60 known negative) thick and thin blood smears.
Table 6, clearly demonstrated that 93.3% of blood smear components were instantly recognizable in blood smears when buffered solution was used to dilute Giemsa stain stock solution. Comparable finding was obtained when distilled water was used as diluents where 88.3% of blood smears components were found to be distinguishable (P- value = 0.18). However when Giemsa stain working solution was prepared using a tap water only 11.7% of blood smears components were distinguishable that is significantly lower than 93.3% when phosphate buffered solution (pH 7.2) for staining blood smears was used (P-value = 0.00).
54 Table 6 Comparison of Giemsa stain working solution for staining blood smears using different diluents:
Diluents
Blood smears Buffered solution Distilled water Tap water
Number examined 120 120 120
Staining of smears 93.3% 88.3% 11.7% (Distinguishable components) (%)
P- value * (Compare to buffered .180 .000 solution)
* McNemar test Exact Sig. (2 sided).
55 3.7 Stability of Giemsa stain working solution:
An attempt to find out a cutoff point for the duration of use after preparation of
Giemsa stain working solution was carried out in the present study. After preparation of
10% Giemsa stain working solution it was used to stain batches of 120 (60 known positive & 60 known negative) blood smears at different intervals after preparation.
Thus, Giemsa satin working solution (10%) was used to stain blood smears immediately, after one hour, 2 hours, 3 hours, 4 hours, 6 hour, 8 hours, 10 hours, and after 12 hours.
As illustrated in figure 6 it is evident that components of blood smear were clearer during the fist three hours after preparation where more than 89% of the components were clear then it started to decline significantly after the third hour of preparation reaching 52.5% after 4 hours and then the quality of the stain deteriorates to 39.2% after
6 hours, 20% after 8 hours, 10% after 10 hours and after 12 hours only 2.5% of the components of blood smears were distinguishable (Appendix 2).
56 Figure 6.The percentage of distinguishable component of blood smears (stain quality) following different intervals of storage:
100
90
80
70
60
50
40
30
20
10 Percentage of distinguishable component blood smears
0
rs rs rs s rs 0 h u u u u o o o o hours h h 1 hour 2 3 h 4 6 hours 8 h 10 hour 12 Time of staining after preparation
57 Chapter Four
DISCUSSION
Although, many new techniques have been developed for the laboratory diagnosis of malaria, nevertheless, the microscopic examination of stained blood smears still remains the recommended method and current gold standard method in use for routine laboratory diagnosis of malaria. In the most capable hands, microscopy can be expected to detect malaria parasites when the density of infection is 5 parasites/µl, moreover microscopy can identify species to the level of 98% of all parasites seen
(Moody, 2002).
Microscopic diagnosis of malaria requires a well functioning microscope, a good quality stain, immersion oil and other consumables. Moreover it needs qualified and well-trained personnel in addition to a quality assurance system (Milne et al., 1994).
This study was carried out in order to clarify some of the queries, which have been raised repeatedly by laboratory staff as regards the techniques used during microscopic diagnosis of malaria.
The superiority of Giemsa stained blood smear in detecting malaria parasites as compared to Saponin lyzed venous blood smear was evident in the present study. Thus, smears prepared after Saponin lysis of venous blood had some limitations due to, firstly incomplete lysis of red blood cells secondly smear occupies a relatively large area of slide, consequently it requires prolonged time for microscopic examination of the blood smear, in order to obtain an accurate result. A Giemsa stained blood smear required about 10 min for microscopic examination while 20 min were required for Saponin lyzed venous blood smear. This is in discrepancy with the finding of Orjih, (1994) who
58 noted that the parasiteamia in Giemsa stained smear preparations treated with Saponin consistently increased detection of malaria parasites.
In the present study the Giemsa-stained thick blood smear was found to be more sensitive in detecting malaria parasites in blood with low parasitaemia. Thus, examination of Giemsa-stained thick blood smears detected malaria parasites when the parasite count was 120 parasite/µL of blood, at this level of parasiteamia when Saponin lyzed venous blood smear was used malaria parasites were not detected. This is likely to be due to incomplete lysis of red blood cells; subsequently it is more difficult to detect malaria parasites especially in low parasitaemia. This is in contrast to the finding of
Orjih, (1994).
In the present study microscopic examination of blood smears stained with Giemsa stain and Fields' stain that were performed blindly by three different technicians independently, revealed that detection of malaria parasites in Giemsa-stained blood smears of 31%, 32.5% and 33% were significantly higher than 26.1, 27.1% and 27.1% the corresponding figures of Fields'-stained blood smears. Therefore, this finding clearly demonstrated that Giemsa stain is the most appropriate stain for staining blood smears in microscopic diagnosis of malaria. Similar finding had been reported in the Sudan by
Ibrahim, (1995) and also reported by World Health Organization, (2004).
The impact of stain in determining the density of infection was evaluated in the present study. Thus, as illustrated in figure 5, in Giemsa-stained blood smears the highest parasite counts were determined. Furthermore, Giemsa stain was found to be much better in detecting malaria parasites in blood with low levels of parasitaemia. The failure to detect malaria parasites in blood smears with low parasitaemia after staining
59 with field's stain is likely to be due to wash out of blood in greater amount compared to
Giemsa stain. A rather lower parasite counts determined after staining with Field’s stain is likely due to partial wash out of the parasite during staining.
In this study three different stocks of Giemsa stain were prepared in order to minimize the use of glycerol thus reducing the cost of stain and having a better stain with minimum of deposits.
Although Eisa, in 1985 noted that it is preferable to use stock stain with a large amount of methanol rather than equal to glycerol, nevertheless, the comparison of using the three batches of Giemsa stock stains in the present study showed similar finding in staining blood smears for malaria parasites detection. So, the three Giemsa stocks stain applied in present study had the same aptitude in staining of blood smears for detection of malaria parasites, therefore it is justifiable to increase the amount of methanol in preparing Giemsa stock stain in order to reduce the cost.
Unsatisfactory results obtained when higher concentration (15% & 20%) of
Giemsa working solutions were used versus conventional concentration (10%) in order to reduce the staining period, is likely to be due to incomplete lysis of red blood cells in thick blood smears which leads to overlapping of these cells. It assumes a dark colour with the results that thick blood smears lost their diagnostic value. In thin blood smears over staining of blood cells was observed and it appeared darker and detection of malaria parasites within the red blood cells was more difficult. The majority of thick blood smears showed a rather satisfactory staining when the concentration of working solution was (10%) and used for staining blood smears for 10 minutes.
60 Thus higher concentrations of working Giemsa stain resulted in over-staining of the blood cells with loss of a distinct parasite texture. Besides such factors as stain concentration and pH of the staining solution, standardization of staining time may be considered necessary for the reproducibility of the Romanowsky Giemsa staining pattern. This is in agreement with Cheesbrough, (1992) and Schulte, (1987).
To find out alternative diluents for stock Giemsa stain, distilled water and tap water were used instead of phosphate buffered solution (pH 7.2). Distilled water showed good quality and there were no significant differences in staining properties when it was used to dilute the stock Giemsa stain. This is in accordance with Shute, (1966), Manson-Bahr and Wilcocks, (1978) and Beaver et al., (1983).
The staining properties were significantly different when the blood smears were stained with working Giemsa stain prepared with tap water (unstable pH) instead of phosphate buffered solution (pH 7.2) the differences observed with this batch, in staining of parasites and red cells being particularly variable. They seem red in colour, lightly stained and there were considerable stain deposit in most smears. This is likely to be due to the pH and turbidity of water. This finding is in accordance with Cheesbrough,
(1992).
To determine the cutoff point of stability of Giemsa stain working solution, Giemsa stain working solution was prepared and used to stain blood smears at different hourly intervals. The working solution showed marked decline of staining ability with time after preparation. The findings showed that Giemsa stain working solutions remained stable in staining blood smears during the first 3 hours after preparation then the stain
61 markedly lost it properties with time. However Bins and others in a study carried out in
1985, concluded that the vigor of the staining solution decrease immediately after preparation, but it keep its properties during 8 hours after preparation, this because of change of its chemicals compounds and oxygenation.
There were observable variations in properties of the staining solutions when it was used after three hours of preparation, its loss its properties, the blood smears were palely stained and the parasites were missed. In additional there was greater formation of stain deposit when the working solution was left to stand for a long time. Thus the Giemsa stain working solutions was good enough to use during the first three hours of its preparation and then should be discarded. Similar findings were reported earlier by
Marshall and others, (1978).
62 CONCLUSION & RECOMMENDATIONS
Conclusion:
The findings of the present study showed that the performance of the standard
Giemsa staining method was more reliable in detecting malarial parasites as compared to
Saponin lyzed venous blood and Field stained smears. Moreover, the Giemsa staining method was sensitive and consistent.
The study findings concerning the Giemsa stain showed that comparable results obtained when the stock stain was made up of a large proportion of absolute methanol instead of equal volume of glycerol, the Giemsa working solutions made by diluting the stock stain with phosphate buffered solutions or distilled water but not tap water gave adequate results, poor results were obtained when higher concentrations of working solutions were used in staining blood smears for a short time. Finally the working solutions of Giemsa were stable and valid for use during 3 hours after preparation.
Recommendations:
1- For microscopy detection of malaria parasites, it is better to use standard Giemsa staining method.
2- Saponin lyzed venous blood technique needs more assessment in order to ovoid the difficulties encountered in this study.
2- It is recommended to prepare the working solutions of Giemsa stain by using phosphate buffered solutions.
3 - Although the working solution of Giemsa stain gives adequate result up to three hours after preparation, but it recommended to use it immediately after preparation.
4 - It recommended to use low concentration of Giemsa working solution.
63 REFERENCES
Adam, A. R. and Maegriath, B. G. (1966). Clinical Tropical Disease, 4th edition, P;
211-50, J & A Churchill Ltd-London.
Ascenzi, A. (1999 ). Malaria diseases and parasites. Parassitologia, 41(1-3):33-8.
Astolfi, P., Lisa, A., Degioanni, A., Tagarelli, A. and Zei, G . (1999). Past malaria, thalassemia and woman fertility in southern Italy. Annals of Human Biology, 26(2):
163-73.
Baird, J. K. (1995). Host age as a determinant of naturally acquired immunity to
Plasmodium. falciparum. Parasitology Today, 11:105–111.
Bassett, M.T., Taylor, P., Bvirakare, J., Chiteka, F. and Govere, E. (1991). Clinical diagnosis of malaria: can we improve? . Journal of Tropical Medicine and Hygiene,
94(1): 65-9.
Bayoumi, R. A. (1987). The sickle-cell trait modifies the intensity and specificity of the immune response against P. falciparum malaria and leads to acquired protective immunity. Medical Hypotheses, 22(3): 287-98.
64 Beadle, C., Long, W. R., Weiss, P. D., Mc Elroy, S. M., Maret, A. J., Oloo, S. and
Hoffman, L. (1994). Diagnosis of malaria by detection of P. falciparum HRP-2 antigen with a rapid dipstick antigen-capture assay. Lancet, 343:564-568.
Beare, N. A., Lewis, D. K., Kublin, J. G., Harding, S. P., Zijlstra, E. E. and
Molyneux, M. E. (2003). Retinal changes in adults with cerebral malaria. Annals of
Tropical Medicine and Parasitology, 97(3): 313-5.
Beaver, P. C., Tung, R. C. and Cupp, E. W. (1983). Clinical Parasitology, 9th ed.,
P; 35-40. Lea & Febiger- USA.
Bins, M., Huiges, W., Halie, MR. (1985). Stability of azure B-eosin Y staining solutions. British Journal of Haematology, 59(1): 73-8.
Brooks, G. F., Butel, J. S. and Mores, S. A. (1998). Jawetz Melnick & Adelberg's
Medical Microbiology, 20th ed., P; 632-634, Appleton & Lange - USA.
Bruce – Chwatt, E. J. (1985). Essential Malariology, 2end ed., p 1-12. ELBS William
Heinemann Medical Books, London- United Kingdom
Bruce-Chwatt, E. J. (1988). History of malaria from prehistory to eradication, p. 1-59.
In W. H. Wernsdorfer and Mc Grego , Malaria: principles and practice of malariology vol. 1. Churchill Livingstone, Edinburgh, United Kingdom.
65
Boukari, B. S., Napo-Koura, G., Kampatibe, N., Kpodzro, K., Rabineau, D. and
Vovor, M. (1991). Congenital malaria: clinical, parasitological and histological considerations. Apropos of 200 observations collected at the Lome University Teaching
Hospital and Kpalime Hospital. Bull Soc Pathol Exot., 84:448-57.
Boulos, M., Costa, J. M. and Tosta, C. E. (1993). Pulmonary involvement in malaria.
Rev Inst Med Trop Sao Paulo. 35(1): 93-102.
Bualombai, P., Prajakwong, S., Aussawatheerakul, N., Congpoung, K., Sudathip,
S., Thimasarn, K., Sirichaisinthop, J. and Indaratna, K. (2003) Determining cost- effectiveness and cost component of three malaria diagnostic models being used in remote non-microscope areas. Southeast Asian Journal of Tropical Medicine and Public
Health. 34(2): 322-33.
Bygbjerg, I. C. and Lanng, C. (1982). Septicaemia as a complication of falciparum malaria. Transactions of the Royal Society of Tropical Medicine and Hygiene, 76(5):
705.
Chaorattanakawee, S., Natalang, O., Hananantachai, H., Nacher, M., Brockman,
A., Krudsood, S., Looareesuwan, S. and Patarapotikul, J. (2003). Storage duration and polymerase chain reaction detection of Plasmodium falciparum from blood spots on filter paper. American Journal of Tropical Medicine and Hygiene, 69(1): 42-4.
66 Charlwood, J. D., Vij, R. and Billingsley, P. F. (2000). Dry season refugia of malaria- transmitting mosquitoes in a dry savannah zone of east Africa. American Journal of
Tropical Medicine and Hygiene, 62(6): 726-32.
Cheesbrough, M. (1992). Medical Laboratory Manual for Tropical Countries.Volume1,
Second ed., p; 221-249. Butter Worth -Heinemann Ltd Oxford – UK.
Cheesbrough, M. (1999). District Laboratory Practice Tropical Countries. Part I, first edition, p; 239 - 241. Cambridge University Press-UK.
Chiodini, P. L. and Moody, A. H. (1989). Techniques for the detection of malaria parasites. Royal Society of Medicine, 82(17): 41-3.
Clark, I. A. and Schofield, L. (2000). Pathogenesis of malaria. Parasitology Today,
16(10): 451-4.
Christopher, V. P. (2003). Monitoring antimalarial drug resistance: making the most of the tools at hand. Journal of Experiment Biology, 206: 3745-3752.
Collier, L.A. and Longmore, J.M. (1983). The reliability of the microscopic diagnosis of malaria in the field and in the laboratory. Annals Tropical Medicine and Parasitology,
77(2): 113-7.
67
Congpuong, K., Bualombai, P., Jitchamroen, S. and Konchom, S. (2001).
Comparison of the Optimal rapid test with routine microscopic examination of Giemsa-
Stained Thick Blood Smear for diagnosis of malaria. Thailand Medical Association
Journal, 84(3): 357-63.
Coombs, G. L. and Sandosham, A. A. (1975). The staining of combined thick and thin blood smears on the same slide with modified Field’s rapid stain Medical Journal
Malay, 20(1): 41- 53.
Cox, F.E.G. (2002). History of Human Parasitology. Clinical Microbiology Reviews,
15(4): 595-612.
Day, N. (2003). Malaria and the blood smear. Tropical Doctor.33 (2): 65.
Delacollette, C., Taelman, H. and Wery, M. (1995). An etiologic study of hemoglobinuria and blackwater fever in the Kivu Mountains, Zaire. . Ann Soc Belg Med
Trop. 75(1): 51-63.
Dietze, R., Perkins, M., Boulos, F., Luz, B., Reller, G. and Corey, G. R. (1995). The diagnosis of P. falciparum infection using a new antigen detection system. American
Journal of Tropical Medicine and Hygiene, 52:45-49.
68 Dion, R. B. (1995). Lecture notes on Tropical Medicine, 4th ed.,, Section 1, Chapter 1, p.31 , Oxford: Blackwell Sciences, UK.
Dorsey, G., Gandhi, M. and Oyugi, J. (2000). Difficulties in the Prevention,
Diagnosis and Treatment of Imported Malaria. Arch Int Med.160 (16): 2505-2510.
Eisa, H. B. (1985). Laboratory Manual for Parasitological techniques, first ed., P; 23-
46. Khartoum University Press.
El Sayed, B. B., Arnot, D. E., Mukhtar, M. M., Baraka, O. Z., Dafalla, A. A.,
Elnaiem, D. E. and Nugud, A. H. (2000). A study of the urban malaria transmission problem in Khartoum. Acta Tropica, 75(2): 163-71.
Ferreira, A. W. (1990) Immunodiagnosis of malaria. In Diagnosis of Malaria (Lopez-
Antunano F. J. & Schmunis G. eds), Pan American Health Organization, Washington:
61- 71
Field, J. W. and Sandosham, A. A. (1964). The Romanowsky stain-Aqueous or
Methanolic? . Transactions of the Royal Society of Tropical Medicine and Hygiene,
58:164-72.
Foo, L. C., Rekhraj, V., Chiang, G. L. and Mak, L. W. (1992). Ovalocytosis protects against severe malaria parasitemia in the Malayan aborigines. American
Journal of Tropical Medicine and Hygiene, 47(3): 271-5.
69
Foote, S. J. and Cowman, A. F. (1994). The mode of action and the mechanism of resistance to antimalarial drugs. Acta Tropica, 56: 157 -171.
Freedman, D.O. (1992). Imported malaria here to stay. American Journal of Medicine,
93: 239-242.
Genton, B., Smith, T., Baea, K., Narara, A., Al Yaman, F., Beck, H. P., Hii, J. and
Alpers, M. (1994). Malaria: how useful are clinical criteria for improving the diagnosis in a highly endemic area? . Transactions of the Royal Society of Tropical Medicine and
Hygiene, 88: 537– 541.
Gilles, H. M. (1984). Infection & Tropical Disease. Medicine International, 4:141-4.
Gleeson, R. M. (1997) . An improved method for thick smear preparation using saponin as a lysing agent. Clinical Laboratory Haematology, 19(4): 249-51.
Gordon, R. M. and Lavoipierre, R. B. (1969). Entomology for Students of Medicine.
Second ed, P; 95-107. Blackwell Scientific Publication Ltd.
Greenwood, B. and Mutabingwa, T. (2002). Malaria in 2002. Nature, 415: 670-672.
70 Guyatt, H. L. and Snow, R. W. (2000). The epidemiology and burden of Plasmodium. falciparum malaria– related anemia among pregnant women in sub- Saharan Africa.
American Journal of Tropical Medicine and Hygiene, 64(1): 36– 44.
Haiana, C. S., Lian, M., Rosana, M., Alexander, B. and Frederica, G. (2001).
Development and optimization of polymerase chainreaction-based malaria diagnostic methods and their comparison with quantitative buffy coat assay. American Journal of
Tropical Medicine and Hygiene, 65(4): 355– 361.
Hamad, A. A., Nugud, A. H., Arnot, D. E., Giha, H. A., Abdel-Muhsin, A. M., Satti,
G. M., Theander, T. G., Creasey, A. M., Babiker, H. A. and Elnaiem, D. E. (2002).
A marked seasonality of malaria transmission in two rural sites in eastern Sudan. Acta
Tropica, 83(1): 71-82.
Hang, V. T., Tran, P. N., Thanh, L. T., Hien, L. V., O'Brien, E. and Morris. G. E.
(1995). Screening donor blood for malaria by polymerase chain reaction. Transactions of the Royal Society of Tropical Medicine and Hygiene, 89: 44-47.
Hanscheid, T. (2003). Current strategies to avoid misdiagnosis of malaria. Clinical
Microbiology Infection, 9(6): 497-504.
Houwen, M. (2002). Blood smear preparation and staining procedures. Clinical
Laboratory Medicine, 22(1): 1-14.
71 Htut, Y., Aye, K. H., Han, K. T., Kyaw, M. P., Shimono, K. and Okada, S. (2002).
Feasibility and limitations of acridine orange fluorescence technique using a Malaria
Diagnosis Microscope in Myanmar. Acta Medicine Okayama, 56(5): 219-22.
Ibrahim, S., M. (1995). Decisive assessment of diagnostic staining methods of malaria in eight public and private laboratories, Khartoum area. TDR/EMRO Final Report
Summary 1995-200, Small Grants Scheme (SGS) 1995 no 12
Igbal, J., Khalid, N. and Hira, P. R. (2002). Comparison of two commercial assays with expert microscopy for confirmation of symptomatically diagnosed malaria. Journal of Clinical Microbiology, 40(12): 4675-8.
Iqbal, J., Hira, P., Al-Ali, F., Khalid, N. and Sher, A. (2003). Modified Giemsa staining for rapid diagnosis of malaria infection. Medical Principal Practice, 12(3): 156-
9.
Jennifer, F. W. (2003). Advancing the war on malaria. Annals of International
Medicine, 138(8): 693-696.
Joklik, W. K., Willitt, H. P. and Amos, D. B. (1980). Zinsser Microbiology, 17th ed., P; 1445 - 1448, Appleton Century- A publishing Division of Prentice – Hall, Inc.
Joss, K. L. (1999). Successful blood draws techniques. Nurse Spectrum (Wash D C).
9(6): 9-10.
72 Kawamoto, F. (1991). Rapid diagnosis of malaria by fluorescence microscopy with light microscope and interference filter. Lancet, 337(8735): 200-2.
Keiser, J., Utzinger, J., Premji, Z., Yamagata,Y. and Singer, B. H. (2002). Acridine
Orange for malaria diagnosis: its diagnostic performance, its promotion and implementation in Tanzania, and the implications for malaria control. Annals of Tropical
Medicine and Parasitology, 96(7): 643-54.
Knight, R. (1985) Parasitic Diseases in Man, First ed., P; 199-205, Longman group Ltd,
Singapore.
Kochhar, R., Goenka, M. K., Mehta, S. and Mehta, S. K. (1990). Gastrointestinal bleeding in malaria. Indian Journal of Gastroenterology, 9(4): 295-6.
Lema, O. E., Carter, J. Y., Nagelkerke, N., Wangai, M. W., Kitenge, P., Gikunda,
S. M., Arube, P. A., Munafu, C. G., Materu, S. F., Adhiambo, C. A. and Mukunza.
M. G. (1999). Comparison of five methods of malaria detection in the outpatient setting .American Journal of Tropical Medicine and Hygiene, 60(2): 177-82.
Lennette, E. H., Houster, W. J., Balows, A. C. and Shadonny, JR. H. (1985).
Manual Clinical Microbiology, 4th ed., P; 612-620, American Society for Microbiology,
Washington- USA.
73 Lowe, B. S., Jeffa, N. K., New, L., Pedersen, C., Engbaek, K. and Marsh, K. (1996).
Acridine orange fluorescence techniques as alternatives to traditional Giemsa staining for the diagnosis of malaria in developing countries. Transactions of the Royal Society of Tropical Medicine and Hygiene, 90(1): 34-6.
Manson - Bahr, P. E. C. and Wilcocks, C. (1978). Manson's Tropical Disease, 17th ed., P; 49 - 86. ELBS & Baillier Tindall- London – UK.
Manson - Bahr, P. E. C. and Apted, F. I. C. (1985). Manson's Tropical Disease, 18th ed., P; 38 - 45. ELBS & Baillier Tindall- London - UK.
Makler, M. T.,Palmer, C. J. and Ager A.L. (1998). Areview of practical techniques for the diagnosis of malaria. Annals of Tropical Medicine and Parasitology, 92(4): 419-
33.
Markell, E. K., John, D. T. and Krotoski, W. A. (1999). Markell & Voge’s Medical
Parasitology. 8th ed. P. 90-121. W.B Sanders Co. Philadelphia – USA.
Marshall, P. N., Bentley, S.A. and Lewis, S.M. (1978). Staining properties and stability of a standardized Romanowsky stain. Journal of Clinical Pathology, 31:280-
282
74 Mason, D., Kawamoto, F. Lin, K., Laoboonchai, A. and Wongsrichanalai, C.
(2002). Comparison of two rapid field microscopy in the tests to expert immunochromatographi diagnosis of malaria. Acta Tropica, 82(1): 51-9.
Mehta, K. S., Halankar, A. R., Makwana, P. D., Torane, P. P., Satija, P . and
Shah, V. B. (2001). Severe acute renal failure in malaria. Postgraduate Medical
Journal, 47(1): 24.
Miller, L. H., Mason, S. J., Clyde, D. F. and Mc Ginniss, M. H. (1976). The resistance factor to Plasmodium vivax in blacks. The Duffy-blood-group genotype,
FyFy. New England Medicine Journal, 295(6): 302-4.
Miller, L H. (2002). The pathogenic basis of malaria. Nature, 415: 673-679.
Milne, L. M., Kyi, M. S., Chiodini, P. L. and Warhurst, D. C. (1994). Accuracy of routine laboratory diagnosis of malaria in the United Kingdom. Journal of Clinical
Pathology. 47:740-742
Mohamed-Nour, B. Y. (1998). Invitro study to asses the response of P.falciparum malaria to chloroquine, sulphadoxine, pyrimethamine, quinine and mefloquine inWad
Medani central state - Sudan. Thesis submitted to master degree. Faculty of Medicine,
University of Gazira- Sudan.
75 Moody, A. (2002). Rapid diagnostic tests for malaria parasite .Clinical Microbiology
Reviews, 15(1): 66-78.
Morassin, B., Fabre, R., Berry, A. and Magnaval, J. (2002). One year’s experience with the polymerase chain reaction as a routine method for diagnosis of imported malaria. American Journal of Tropical Medicine and Hygiene, 66(5): 503– 508.
Murphy, G. S. (1993). Vivax malaria resistant to treatment and prophylaxis with chloroquine. Lancet 341:96-100.
Nunn, J. F. and Tapp, E. (2000). Tropical diseases in Ancient Egypt. Transactions of the Royal Society of Tropical Medicine and Hygiene, 94:147-153.
Oh, M. D., Shin H., Shin, D., Kim, U., Lee, S., Kim, N., Choi, M. H., Chai, J. Y. and
Choe, K. (2001). Clinical features of vivax malaria. American Journal of Tropical
Medicine and Hygiene, 65(2): 143-6.
Ohrt, C., Purnomo, S., Sutamihardja, M. A., Tang, D. and Kain, K. C. (2002).
Impact of microscopy error on estimates of protective efficacy in malaria-prevention trials. Journal of Infectious Diseases, 186(4): 540-6.
Orjih, A. U. (1994). Saponin haemolysis for increasing concentration of Plasmodium falciparum infected erythrocytes. Lancet, 343(8892): 294-298.
76 Pandey, J., Talib, V. H., Ranga, S. and Gulati, I. (1995). Diagnosis of malaria--an overview. Indian Journal of Pathology & Microbiology, 38(4): 441-7.
Patsoula, E., Spanakos, G., Sofianatou, D., Parara, M. and Vakalis, N. C. (2003). A single step, PCR- based method for the detection and diVerentiation of P. vivax and P. falciparum. Annals of Tropical Medicine and Parasitology, 97(1): 15– 21.
Philip, J. R. (2003) . Antimalarial drug discovery: old and new approaches. Journal of
Experimental Biology, 206: 3735-3744.
Piper, R., Lebras, J., Wentworth, L., Hunt- Cooke, A., Houze, S., Chiodini, P. and
Makler, M. (1999). Immunocapture diagnostic assays for malaria using
Plasmodium falciparum lactate dehydrogenase (pLDH). American Journal of Tropical
Medicine Hygiene, 60: 109– 118.
Playford, E. and Walker, J. (2002). Evaluation of the ICT Malaria P.f/P.v and the
OptiMAL Rapid Diagnostic Tests for Malaria in Febrile Returned Travelers. Journal of
Clinical Microbiology, 40(11): 4166- 4171.
Ponnuduari, T., Billingsely P. F. and Rudin, W. (1988). Differential infectivity plasmodium for mosquitoes. Parasitology Today, 4:319-321.
Power, K., T. (1982) The Romanowsky stains: a review. American Journal of Medical
Technology 48: 519-523.
77 Rajakumar, K. and Weisse, M. (1999). Centennial year of Ronald Ross’ epic discovery of malaria transmission: an essay and tribute .South Medical Journal 92(6):
567-71.
Ray, K. (1984) Immunodiagnosis of malaria. Journal of Community Diseases, 16(1):
24-9.
Reed, M. B., Saliba, K. J., Caruana, S. R., Kirk, K. and Cowman, A. F. (2000). Pgh1 modulates sensitivity and resistance to multiple antimalarials in Plasmodium falciparum. Nature, 403: 906 -909.
Rubio, J. M., Buhigas, I., Subirats, M., Baquero, S., Puente, N. and Bento, A.
(2001). Limited level of accuracy provided by available rapid diagnostic tests for malaria enhances the need for PCR-based reference laboratories. Journal of Clinical
Microbiology, 39:2736- 2737.
Sacks, J. and Malaney, P. (2002). The economic and social burden of malaria. Nature,
415:680-685.
Schapira, A., Fogh, S., Jepsen, S. and Pedersen, N. S. (1994). Detection of antibodies to malaria: comparison of results with ELISA, IFAT, and crossed immunoelectrphoresis.
Acta Pathology Microbiology Immunology Scand, 92(6): 299-304.
78 Schmidt, G. D. and Robert, C. S. (1989). Foundation of Parasitology 4Th ed, P; 139-56,
Times Mirror/ Mosby College Publishing-USA.
Schulte, E. (1987). Standardization of the Romanowsky-Giemsa stain: the influence of staining time on the Romanowsky Giemsa-staining pattern. Acta Histochemistry,
34:153-62.
Shalev, O., Tsur, A. and Rahav, G. (1992). Falciparum malaria- induced hypoglycaemia in a diabetic patient. Postgraduate Medical Journal, 68(798): 281-2.
Shute, P.G. and Margon, M .G. (1966). Laboratory techniques for the study of malaria.
2end ed., P; 12-26. J &A Churchill Ltd- London-UK.
Sidhu, A. B., Verdier - Pinard, D. and Fidock, D.A. (2002). Chloroquine resistance in
Plasmodium falciparum malaria parasites conferred by pfcrt mutations. Science,
298:210 –213.
Smith, T. G., Ayi, K., Serghides, L., Mcallister, C. D. and Kain, K. C. (2002). Innate immunity to malaria caused by Plasmodium. falciparum. Clinical Investigation Medical
Journal, 25(6): 262-72.
Smith, T., Lines, J. and Marsh, K. (1993) Absence of seasonal variation in malaria parasitaemia in an area of intense seasonal transmission. Acta Tropica, (54): 55–72.
79 Snounou, G., Viriyyakosol, S., Jarra, W. and Thaithon, W. 1993). Identification of the four human malaria parasite in field samples by the polymerase chain reaction and detection of high prevalence of mixed infection. Molecular and Biochemical
Parasitology, 58:283-292.
Song, L. H., Goh, E. S., Phang, L. C., Poh, W. T. and Tay, S. K. (2000). Technical aspect of Thin Preparation. Singapore Medical Journal, 41(12): 575-8.
Srinivasan, S., Moody, A. and Chiodini, P. (2000). Comparison of blood- smears microscopy, the OptiMAL dipstick, rhodamine- 123 fluorescence staining and PCR, for monitoring antimalarial treatment. Annals of Tropical Medicine and Parasitology,
94:227–232.
Sudan Federal Ministry of Health, (2003). Options against malaria in sudan, conference report14- 15 October Khartoum. Sudan Federal Ministry of Health, World
Health Organization and Médecins Sans Frontières.
Thakor, H. G. (2000). Laboratory diagnosis of malaria. Indian Medical Association
Journal, 98(10): 623-7.
Thellier, M., Datry, A., Alfa, O., San, C., Biligui, S., Silvie, O. and Danis, M. (2002).
Diagnosis of malaria using thick bloodsmears: definition and evaluation of a faster protocol with improved readability. Annals of Tropical Medicine Parasitology, 96(2):
115-24.
80 United Nations (2003). Human Development Reports. Consolidated Inter-Agency
Appeal for Sudan.
Van den Ende, J., Vervoort, T., Van Gompel, A. and Lynen, L. (1998). Evaluation of two tests based on the detection of histidine rich protein 2 for the diagnosis of imported
Plasmodium falciparum malaria. Transactions of the Royal Society of Tropical Medicine and Hygiene, 92(3): 285-8.
Van der Hoek, W., Premasiri, D. A. and Wickremasinghe A. R. (1998). Clinical diagnosis of uncomplicated malaria in Seri Lanka. Southeast Asian Journal of Tropical
Medicine & Public Health. 29(2): 242-5.
Weatherall , DJ. And Abdalla, S. (1982). The anaemia of plasmodium falciparum malaria. British Medical Bulletin, 38(2): 147-51.
Wongsrichanalai, C., Arevalo, I., Laoboonchai, A., Yingyuen, K., Miller, R. S.,
Magill, A. J., Forney, J. R. and Gasser, R. A. (2003). Rapid diagnostic devices for malaria: field evaluation of a new prototype immunochromatographic assay for the detection of Plasmodium falciparum and non-falciparum Plasmodium. American
Journal of Tropical Medicine and Hygiene, 69(1): 26-30.
World Health Organization (1973). Chemotherapy of Malaria and Resistance to
Antimalarials. WHO Technical Report. 529 Series. Geneva: World Health Organization.
81 World Health Organization. (1980). Manual of basic technique for a health laboratory. P; 196-203.
World Health Organization. (1989). Malaria control as part of primary health care,
Technical Report Series, No 712.
World Health Organization. (1993). A global strategy for malaria control., Geneva:
World Health Organization
World Health Organization. (1996). World malaria situation. Part 1. WHO Weakly –
Epidemiological Record.71: 17-22,
World Health Organization (1996). Assessment of therapeutic efficacy of antimalarial drugs for uncomplicated falciparum malaria in areas with intense transmission. Geneva, World Health Organization,(WHO/MAL/96.1077).
World Health Organization (2000). Severe falciparum malaria. Transactions of the
Royal Society of Tropical Medicine and Hygiene, 94: 1–90.
World Health Organization (2004). Malaria / Diagnostic Method. Operational
Research in Tropical and Communicable Diseases / World Health Organization,
Regional Office for the Eastern Mediterranean (EMRO).
82 APPENDICES
Appendix 1. Relationship between detection of malaria parasites according to level of parasiteamia in blood smears stained with different stains:
Giemsa Saponin Field
No examined 203 203 203
No of positive smears 67 61 55
Smears with <1000 parasites/µl of blood 25 23 19
Smears with 1000-4000 parasites/µl of blood 11 11 10
Smears with 4001-7000 parasites/µl of blood 6 6 6
Smears with 7001-10000 parasites/µl of blood 7 6 6
Smears with 10001-13000 parasites/µl of blood 6 6 6
Smears with 13001-16000 parasites/µl of blood 4 4 3
Smears with 16001-19000 parasites/µl of blood 3 2 2
Smears with 19001-22000 parasites/µl of blood 3 2 2
Smears with >22000 parasites/µl of blood 2 1 1
83 Appendix 2. Comparison between the quality of Giemsa stain working solution and
time after preparation of the working solution:
Time of staining after preparation of Giemsa stain working solution
Blood smears 0 h 1 h 2 h 3h 4 h 6 h 8 h 10h 12 h
Number examined 120 120 120 120 120 120 120 120 120
Quality of smear staining (Distinguishable 96.7 92.5 90.8 89.6 52.5 39.2 20 10 2.5 components) (%)
P-value* (Compare to immediately .180 .065 .058 .000 .000 .000 .000 .000 after preparation)
* McNemar test Exact Sig. (2 sided).
84 Appendix 3. Preparation of stock Giemsa stain (Stock I):
One liter of stock stain prepared by the following method:
Giemsa powder ((Nen Tech.Ltd-UK) 7.6 grams
Absolute Methanol (BDH Chemical Ltd) 500 ml
Pure glycerol (BDH Chemical Ltd) 500 ml
1- The powder weighted on a piece of clean paper and transferred to a dry brown
bottle of one liter capacity which contains a few glass beads.
2- A dry cylinder used to measure the methanol and added to powder and well
mixed.
3- The same cylinder used to measure the glycerol and added to stain and well
mixed.
4- The bottle shake for five minutes and this shaking repeated for next 7 days after
which the stain ready for used.
85 Appendix 3a. Preparation of stock Giemsa stain (Stock II):
One liter of stock stain prepared by the following method:
Giemsa powder ((Nen Tech.Ltd-UK) 7.6 grams
Absolute Methanol (BDH Chemical Ltd) 667 ml
Pure glycerol (BDH Chemical Ltd) 333 ml
1- The powder weighted on a piece of clean paper and transferred to a dry brown
bottle of one liter capacity which contains a few glass beads.
2- A dry cylinder used to measure the methanol and added to powder and well
mixed.
3- The same cylinder used to measure the glycerol and added to stain and well
mixed.
4- The bottle shake for five minutes and this shaking repeated for next 7 days after
which the stain ready for used.
86 Appendix 3b. Preparation of stock Giemsa stain (Stock III):
One liter of stock stain prepared by the following method:
Giemsa powder (Nen Tech.Ltd-UK) 7.6 grams
Absolute Methanol (BDH Chemical Ltd) 750 ml
Pure glycerol (BDH Chemical Ltd) 250 ml
1- The powder weighted on a piece of clean paper and transferred to a dry brown
bottle of one liter capacity which contains a few glass beads.
2- A dry cylinder used to measure the methanol and added to powder and well
mixed.
3- The same cylinder used to measure the glycerol and added to stain and well
mixed.
4- The bottle shake for five minutes and this shaking repeated for next 7 days after
which the stain ready for used.
87 Appendix 4. Preparation of Field’s stain solution A:
500 ml of stain was prepared by the following method:
Field’s stain A powder (BDH Chemical Ltd) 5 grams
Distilled water (hot) 500 ml
1- The powder weighted on a piece of clean paper and transferred to a dry large
Pyrex beaker .
2- A dry cylinder used to measure water and heat to boiling.
3- The hot water added to stain and mixed to dissolve the powder.
4- When the stain was filtered into a clean-labeled bottle.
88 Appendix 4a. Preparation of Field’s stain solution B:
500 ml of stain was prepared by the following method:
Field’s stain B powder (BDH Chemical Ltd) 5 grams
Distilled water (hot) 500 ml
1- The powder weighted on a piece of clean paper and transferred to a dry large
Pyrex beaker.
2- A dry cylinder used to measure water and heat to boiling.
3- The hot water added to stain and mixed to dissolve the powder.
4- When the stain was filtered into a clean-labeled bottle.
89 Appendix 5. Preparation of Saponin lyzing solution:
Three liter of Saponin lyzing solution was prepared by the following method:
Saponin powder (Seelze Hanover- Germany) 0.45 grams
Normal Saline * 3 L
1- The powder weighted on a piece of clean paper and transferred to a dry
brown bottle of three liters capacity.
2- The Normal Saline added to the powder and mixed to dissolve the
powder.
* Normal saline prepared by dissolving 25.5 grams of Sodium Chloride in 3 liters of distilled water.
90
Appendix 6. Preparation of phosphate buffered solution pH 7.2:
Disodium hydrogen phosphate 1 gm
Potassium dihydrogen phosphate 0.7 gm
Distilled water 1 Litre
91 Appendix 7. Questionnaire :
NO……………
Name………………………………………Sex …………………… Age……………….
Result:
Positive
Negative Species Stage Density
Giemsa stained thick smear
Saponin smear
Field thick stained smear
92 Appendix 8. Form used to assess the blood smears of study techniques:
No. Parasite Parasite WBC Background WBCs RBCs Score Assessment nucleus cytoplasm nuclei 1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
93