MANUAL FOR THE LABORATORY DIAGNOSIS OF MALARIA

2nd Edition

MANUAL

FOR THE LABORATORY

DIAGNOSIS OF MALARIA

Ethiopian Public Health Institute

Federal Ministry of Health

January, 2020

Contents Foreword ...................................................................................................................... i

Acknowledgment....................................................................................................... iii

I. Acronyms ................................................................................................................ v

II. Glossary ................................................................................................................ viii

III. Scope and Purpose of the Manual ..................................................................... xv

A. Purpose ............................................................................................................. xv

B. Target Audience ................................................................................................ xv

CHAPTER ONE: Introduction to Malaria ......... 1 1.1. Malaria Etiology .................................................................................................. 1

1.2. Life Cycle and Mode of Transmission of Plasmodium ..................................... 2

1.3 Malaria Parasites ................................................................................................ 3

1.3.1 Incubation Period ................................................................................................ 5

1.3.2 Uncomplicated Malaria ...................................................................................... 5

1.3.3 Complicated (Severe Malaria) .......................................................................... 6

1.3.4. Malaria Relapses ................................................................................................. 8

1.3.5. Recrudescence ..................................................................................................... 8

1.3.6. Malaria Reinfection ............................................................................................. 9

1.3.7. Other Manifestations of Malaria ...................................................................... 9

1.4. Human Factors Resistance to Malaria ............................................................... 9

1.4.1. Genetic Factors .................................................................................................... 9

1.4.2. Acquired Immunity ........................................................................................... 10

1.5. Methods for Malaria Diagnosis........................................................................ 11

1.5.1. Clinical Diagnosis of Malaria .......................................................................... 11

1.5.2. Laboratory Diagnosis of Malaria .................................................................... 12

1.6. Global and Regional Burden of Malaria .......................................................... 19

1.6.1. Global Burden .................................................................................................... 19

1.6.2. Malaria Situation in Ethiopia .......................................................................... 20

1.6.3. National Malaria Control Strategy ................................................................. 23

1.6.4. Levels of Health Facilities and Types of Diagnostic Tests in Ethiopia .... 25

CHAPTER TWO: Microscope: Types, Parts,

Care and Handling .......................................... 27 2.1. Microscope ........................................................................................................ 27

2.2. Types of Microscope ........................................................................................ 27

2.2.1. Simple Microscope ........................................................................................... 27

2.2.2. Compound Microscope .................................................................................... 28

2.2.3. Electron Microscopy ......................................................................................... 35

2.3. Care and Handling of Microscope ................................................................... 35

2.4. Microscope Maintenance and Storage Conditions ........................................ 36

2.5. Log Book ........................................................................................................... 41

CHAPTER THREE: Laboratory Safety ............ 42 3.1 Introduction ....................................................................................................... 42

3.2 General Safety Guidelines ............................................................................... 43

3.3 Safety and Exposure Control Measures .......................................................... 45

3.4 Testing Infrastructure and Equipment Management ..................................... 48

3.5. Waste Disposal ................................................................................................. 50

CHAPTER FOUR: Specimen Collection, Smear

Preparation, Fixation and Staining (Pre-

Examination Process) .................................... 52 4.1. Blood Sample Collection .................................................................................. 52

4.1.1. Capillary Blood Collection ............................................................................... 52

4.1.2. Venous Blood..................................................................................................... 52

4.2. Blood Film Preparation..................................................................................... 53

4.2.1. Types of Blood Films ........................................................................................ 53

4.2.2. Common Mistakes in Making Blood Films .................................................. 55

4.3. Fixation of Blood Film ...................................................................................... 58

4.4. Staining of Blood Film ...................................................................................... 58

4.4.1. Principles of Romanowsky Stains .................................................................. 58

4.4.2. Giemsa Stain ...................................................................................................... 58

4.4.3. Field’s Stain ........................................................................................................ 59

4.5. Buffer Solution for Malaria Staining ..................................................................... 60

4.5.1. Buffer Tablets..................................................................................................... 60

4.5.2. Quality Control of Buffered Water ................................................................. 60

CHAPTER FIVE: Microscopic Examination and

Species Identification ............................................. 61 5.1. Examining Blood Films for Malaria Parasites ................................................. 61

5.2. Systematic Approach of Examining Thick and Thin Blood Films ................. 61

5.2.1 Examining the Thick Film ................................................................................. 61

5.2.2 Examining the Thin Film ................................................................................... 63

5.3. Identification of Malaria Parasite Species, Other Blood Parasites and

Artifacts ..................................................................................................................... 63

5.4. Microscopic Differentiation .............................................................................. 68

5.5. Artefacts and Contaminants Confusing Malaria Parasites............................. 78

5.6. Malaria Parasite Counting Methods ................................................................ 79

5.6.1. Number of Parasites/µl Of Blood (thick film): .............................................. 79

5.6.2. Proportion of Parasitized Erythrocytes/Total RBC Counts (thin film): ..... 80

5.6.3 Number of Parasites/µl Of Blood (thin film):................................................. 81

5.6.4 Semi Quantitative Count (thick film) .............................................................. 81

5.7. Reporting Blood Film Results .......................................................................... 81

CHAPTER SIX: Parasitological Diagnosis of Malaria

using Rapid Diagnostic Tests (RDTs) ..................... 83 6.1. RDTs and Their Significance ............................................................................. 83

6.2. RDT Versus Microscopy ................................................................................... 84

6.3. Malaria RDT Formats ........................................................................................ 85

6.4. Types of Malaria RDTs ..................................................................................... 86

6.5. Basic Principles of RDTs ................................................................................... 86

6.6. RDTs Mechanism of Action .............................................................................. 87

6.7. General Procedures of Malaria RDTs .............................................................. 88

6.8. Test Procedure .................................................................................................. 90

6.9. Strengths and Challenges of RDT .................................................................... 91

6.9.1 Strengths ............................................................................................................. 91

6.9.2 Challenges ........................................................................................................... 92

6.10. RDT Kit Selection and Handling ..................................................................... 92

6.10.1. The Plasmodium Species to Be Detected ................................................... 92

6.10.2. Accuracy (Sensitivity and Specificity) .......................................................... 92

6.10.3. Shelf Life and Stability .................................................................................... 93

6.10.4. Ease of Use ....................................................................................................... 93

6.10.5. Cost .................................................................................................................... 94

CHAPTER SEVEN: Document and Record

Keeping ........................................................... 95 7.1 Essential Elements of Recording and Reporting .............................................. 95

7.2 Laboratory Request and Report Forms ............................................................. 95

7.3 Entry of Data into the Laboratory Register ....................................................... 96

7.4 Consequences of Incorrect Reporting ............................................................... 97

7.5 Importance of Malaria Data ............................................................................... 97

7.6. Laboratory Confirmed Malaria Case Report Form ......................................... 97

7.7 Malaria Laboratory Performance Report Form ................................................ 98

CHAPTER EIGHT: Supply and Logistics

Management in Malaria Laboratory

Diagnosis...... .................................................. 99 8.1. Logistics Management ..................................................................................... 99

8.2. Supply List for Malaria Diagnosis .................................................................... 99

8.3. Logistics Management Information System (LMIS) ..................................... 101

8.4. Stock Management ......................................................................................... 102

8.4.1 Inventory Control ............................................................................................. 102

8.4.2 Assessing Malaria Stock Status .................................................................... 103

8.4.3 Conducting a Physical Count ......................................................................... 104

8.4.4 Conducting a Visual Inspection ..................................................................... 104

8.4.5 Record Keeping ................................................................................................ 105

8.4.6 Calculation of Required Supplies .................................................................. 105

8.5. How to Calculate Required Supply Levels: ................................................... 105

8.6. Storage of Malaria Laboratory Commodities ............................................... 108

8.6.1. Guideline for Malaria Laboratory Diagnosis Supply Storage ................. 108

8.6.2. Handling Damaged or Expired Stocks ........................................................ 109

CHAPTER NINE: Quality Assurance of Malaria

Laboratory Diagnosis ................................... 110

9.1. What is Quality Assurance? ........................................................................... 110

9.2. The Need for Accurate Malaria Laboratory Diagnosis ................................. 111

9.3. Errors Compromising Quality Laboratory Diagnosis ................................... 112

9.4. Objectives of Quality Assurance Programs .................................................. 112

9.5. Challenges in Malaria Laboratory Diagnosis ................................................ 113

9.6. Setting up a QA System ................................................................................. 113

9.7. Principles of QA in Malaria Laboratory Diagnosis........................................ 114

9.8. Components of Quality Assurance in Malaria Microscopy.......................... 114

9.8.1. Quality Control (QC) ....................................................................................... 114

9.8.2. External Quality Assessment (EQA) ............................................................ 116

9.8.3. Quality Improvement ..................................................................................... 122

9.9. Quality Assurance (QA) of Malaria RDTs ...................................................... 123

9.9.1. Planning for RDT Introduction ...................................................................... 123

9.9.2. Procurement ..................................................................................................... 124

9.9.3. Lot Testing: Pre- and Post-Market ................................................................ 124

9.9.4. Monitoring Performance in the Field .......................................................... 125

9.9.5. Training and Instructions for Users ............................................................. 125

9.9.6. Use of Results and Community Education ................................................. 125

9.9.7. Storage and Transport ................................................................................... 126

9.10. Quality Assurance of Malaria RDTs in Remote Areas ................................. 126

9.10.1. Ensuring Quality of RDTs ........................................................................... 126

9.10.2. External Quality Assessment of Malaria RDTs........................................ 127

9.10.3. Quality Indicators of Malaria RDT ............................................................. 128

CHAPTER TEN: Professional Ethics and Good

Laboratory Practices .................................... 129 10.1 What Is Ethics ............................................................................................... 130

10.2. Why is Ethics Important ............................................................................... 131

10.3. Types of Ethics .............................................................................................. 131

10.4. Elements of A Strong Work Ethics .............................................................. 133

10.5. Principle of Ethics ......................................................................................... 134

10.6. Core Values of Ethics.................................................................................... 135

10.7. Confidentiality and informed consent. ....................................................... 135

10.8. Right and Obligations of Medical Laboratory Professionals ..................... 136

10.9. Professional Malpractice ............................................................................. 138

10.10. Ethics and Law ............................................................................................. 140

10.11. Good Laboratory Practices (GLPs) .............................................................. 141

References .................................................... 143

Annexes ........................................................ 145 Annex 1: Microscope: Types, Parts, Care and Handling ...................................... 145

Annex 2: SOP for Care and Preventive Maintenance of Microscopes ................ 151

Annex 3: SOP for Capillary Blood Collection and Preparation of Malaria Blood

Films ........................................................................................................................ 153

Annex 4: SOP Preparation of Giemsa Stock Solution .......................................... 158

Annex 5: SOP Preparation of Giemsa Working Solution ..................................... 161

Annex 6: SOP for Preparation of Buffered Water ................................................. 162

Annex 7: SOP for Examination of Malaria Blood Films and Estimation of

Parasitemia ............................................................................................................. 163

Annex 8: SOP for Recording and Reporting of Malaria Blood Film Results ....... 166

Annex 9: Monthly Malaria Case Report Format ................................................... 167

Annex 10: SOP for Malaria Blood Film Slide Storage and Selection for Blinded

Rechecking .............................................................................................................. 168

Annex 11. Blinded Rechecking Result Recording and Feedback Forms ............. 169

11.1. Selected Slide Result Recording Form for Rechecking ........................... 169

11.2. Slide Reader Result Record Form for Rechecking (2nd Reader) ........... 169

11.3. Slide Reader Result Record Form for Rechecking (3rd Reader for

Discordant Result) .................................................................................................... 170

11.4. Performance Notification Form ................................................................... 171

Annex 12: Exposure Reporting Form .................................................................... 173

Annex 13. List of Contributors ............................................................................... 174

List of Tables

Table 3. 1 Safety Precautions for Chemicals Used in Malaria Microscopy. ............. 44

Table 4. 1 Most Common Technical Mistakes in Collection and Preparation of

Blood Smears ............................................................................................................... 57

Table 5. 1 Characteristics of Thick and Thin Blood Films.......................................... 63

Table 5. 2 Species Differentiation on Thin and Thick Films ...................................... 67

Table 5. 3 Species Differentiation of Malaria Parasites by Cytoplasmic Pattern of

Trophozoites in Giemsa-Stained Thick Blood Films Species .................................... 78

Table 6. 1 Comparison of RDTs Versus Microscopy ................................................. 84

Table 6. 2. Comparison of Rapid Diagnostic Tests for Malaria Antigens ................. 87

Table 6. 3 Limitations of RDT Results ......................................................................... 90

Table 8. 1 Example of A Stock Book ..........................................................................105

Table 8. 2 Example of Stock Card ............................................................................. 105

Table 8. 3 Example of A Quarterly Supplies Request and Report, Requirement

Form ........................................................................................................................... 107

Table 8. 4 Example of A Quarterly RDT Supplies Requirement Form .................... 107

List of Figures

Figure 1. 1. Life cycle of Plasmodium; Source: http://www.dpd.cdc.gov/dpdx ......... 2

Figure 1. 2. Quantitative buffy coat of packed blood ................................................. 15

Figure 1. 3. Malaria parasite with AO stained and examined under fluorescence

microscopy ................................................................................................................... 15

Figure 1. 4. Indirect immune fluorescent test for malaria parasite ........................... 17

Figure 1. 5. Malaria risk map of districts by annual parasite incidence, Ethiopia,

2017 .............................................................................................................................. 23

Figure 2. 1. Simple microscope .................................................................................. 28

Figure 2. 2. Compound microscope ............................................................................ 29

Figure 2. 3. Cabinet boxes ........................................................................................... 37

Figure 3. 1. Hazard safety signs .................................................................................. 47

Figure 3. 2. Some important Infrastructures for safe working area ......................... 48

Figure 3. 3. General safety equipment ....................................................................... 49

Figure 4. 1. Unstained and stained blood films. ........................................................ 55

Figure 4. 2. Badly positioned blood film .................................................................... 55

Figure 4. 3. Too much blood for both thin and thick films ........................................ 56

Figure 4. 4. Too small blood for both thin and thick film .......................................... 56

Figure 4. 5. A film made on a very greasy slide. ........................................................ 56

Figure 4. 6. The effect of chipped edge spreader on thin and thick films ................ 57

Figure 5. 1. Systematic approach of examining thick blood film ............................. 62

Figure 5. 2. Systematic approach of examining thin blood film ............................... 63

Figure 5. 3. Basic components of a malaria parasite inside a red blood cell ........... 64

Figure 5. 4. Trophozoites stage of the malaria parasite ............................................ 65

Figure 5. 5. Stages of schizonts growth ..................................................................... 65

Figure 5. 6. Gametocytes of Plasmodium falciparum and Plasmodium malariae .. 66

Figure 5. 7. The appearance of different stages of Knowlesi compared to P.

falciparum and P. malariae stages in thin blood film ................................................ 70

Figure 5. 8. Appearance of different species of Plasmodium in a thin blood film ... 71

Figure 5. 9. Appearance of different species of Plasmodium in a thick blood film

same as above ............................................................................................................. 71

Figure 5. 10. Blood elements, artefacts and contaminants that cause confusion. .. 79

Figure 6. 1. Different formats of malaria RDT: A-cassette; B-dipsticks; and C-card

test ................................................................................................................................ 85

Figure 6. 2. Mode of action of antigen-detecting malaria rapid diagnostic tests

(RDTs). .......................................................................................................................... 88

Figure 9. 1. The quality assurance cycle source: ..................................................... 111

i

Foreword

Malaria is one of the public health diseases in Ethiopia with predominant unstable

transmission. In 2017, the FMOH updated the country’s malaria risk strata based

upon malaria annual parasite incidence (API). Based on NMSP is updated plan for

the years 2017-2020 new epidemiological stratification,75% of the land mass is

malarious and the proportion of the population at risk of malaria is about 60% with

54 (6.4%) woredas having high transmission, chiefly at altitudes below 2,000 meters.

Malaria is mainly seasonal in the highland fringe areas and of relatively longer

transmission duration in lowland areas, river basins and valleys. The Ministry has

officially declared malaria elimination efforts for selected 239 low malaria burden

districts. Ongoing discussions are occurring with the FMOH to coordinate pre-

elimination activities together with other donor-supported projects that continue to

help shrink the malaria transmission map in Ethiopia.

According to WHO 2018 malaria report, Ethiopia marked decreases over 240 000

fewer cases in 2017 than in 2016.This still requires improving diagnosis of malaria

cases using microscopy or using multi-species RDTs, and providing prompt and

effective malaria case management at all health facilities in the country.

This manual is developed based on the recommendations of experts working in

Malaria Programs at the Federal Ministry of Health, Regional Health Bureaus,

National and Regional Reference Laboratories, and partners with the aim of

standardizing malaria laboratory diagnosis and strengthening the quality of

laboratory testing procedures for the diagnosis of malaria in the health facilities in

Ethiopia.

The manual is divided into nine chapters: Scope and purpose of the manual,

introduction to malaria ethology, global burden of malaria and malaria situation in

Ethiopia, parasitological diagnosis of malaria using microscopy, parasitological

diagnosis of malaria using RDTs, quality assurance of malaria laboratory diagnosis,

laboratory safety, supply and logistics management in malaria laboratory diagnosis,

and annexes of formats, registers and Standard operating procedures.

ii

EPHI believes that this manual will be useful for laboratory personnel and other

health workers during routine laboratory work and as a reference material for

trainers and supervisors on laboratory diagnosis of malaria during pre-service and

in-service trainings, practical attachments, during mentorship and supportive

supervisions and for quality control and quality assurance purposes. The manual

could be useful as a reference material for clinicians too, mainly to understand the

use and interpretation of laboratory tests for malaria case management. The manual

is also helpful for health facility managers to enable them in determining essential

laboratory commodity requirements for malaria laboratory diagnosis and the need

for their timely availability to ensure uninterrupted laboratory diagnostic services.

This manual should also be of interest to those non-governmental organizations and

funding agencies that are involved in the support for malaria laboratory diagnosis

improvement and quality assurance programs.

Finally, I would like to express my sincere appreciation and thanks to all

professionals and organizations who have contributed their expertise and resources

for the preparation of this manual.

Eba Abate (PhD)

Director General,

Ethiopian Public Health Institute

iii

Acknowledgment

The development of this Manual was made possible through the contribution of the

professionals and institutions listed below:

Name Organization

Abnet Abebe EPHI

Adisu Kebede EPHI

Gonfa Ayana EPHI

Wondwossen Kassa EPHI

Wondimeneh Liknaw EPHI

Adugna Abera EPHI

Desalegn Nega EPHI

Geremew Tasew EPHI

Bokretsiyon Gidey EPHI

Feven Girmachew EPHI

Foziya Mohammed EPHI

Shemsu Kedir EPHI

Asmare Mekonnen EPHI

Feleke Belachew ICAP

Mekonnen Tadesse ICAP

Leyikun Demeke ICAP

Afework Tamiru ICAP

Kinde Mulatu ICAP

Kefeni Kelbecha ICAP

Dr. Bereket Alemayehu ICAP

iv

The development of first edition of this Manual was made possible through the

contribution of the professionals and institutions listed below:

Contributors: Organization

Gudeta Tibesso EHNRI

Gonfa Ayana EHNRI

Ashenafi Assefa EHNRI

Abinet Abebe EHNRI

Yenew Kebede CDC Ethiopia

Zenebe Melaku CU-ICAP Ethiopia

Abebe Tadesse CU-ICAP Ethiopia

Fanuel Zewdu CU-ICAP Ethiopia

Meseret Habtamu CU-ICAP Ethiopia

Mekonnen Tadesse CU-ICAP Ethiopia

Joseph Malone CDC/PMI Ethiopia

Richard Reithinger USAID/PMI Ethiopia

Hiwot Teka USAID/PMI Ethiopia

Institutions

• Federal Ministry of Health

• Federal Hospitals

• Regional Health Bureaus

• Regional Reference Laboratories

• I-TECH

• The Carter Center

• Malaria Consortium

Core Group Members: Organization

Getachew Belay EHNRI

Habtamu Asrat EHNRI

Markos Sileshi EHNRI

Hussien Mohammed EHNRI

Sindew Mekasha EHNRI

Moges Kassa EHNRI

Bereket Hailegiorgis CU-ICAP New York

Tesfay Abreha CU-ICAP Ethiopia

Sintayehu G/Sellasie CU-ICAP Ethiopia

Leykun Demeke CU-ICAP Ethiopia

Samuel Girma CU-ICAP Ethiopia

Micheal Aidoo CDC Atlanta

v

I. Acronyms

Asl Above Sea Level

ARDS Acute Respiratory Distress Syndrome

AQ Amodiaquine

API Annual Parasite Incidence

Ab Antibody

Ag Antigen

ACT Artemisinin-based Combination Therapy

AL Arthemisisin-Lumefantrine

AMU Average Monthly Usage

QC Chloroquine

DNA Deoxyribose Nucleic Acid

ELISA Enzyme linked Immune-sorbent Assay

EPHI Ethiopian Public Health Institute

EDTA Ethylene Diamine Tetra Acetic Acid

ECP Exposure Control Plan

EQA External Quality Assessment

FMOH Federal Ministry of Health

GNFM Global New Funding Model

HC Health Center

HEWs Health Extension Workers

H.F Health Facility

HMIS Health Management Information System

HP Health Post

HSDP Health Sector Development Programme

HSTP Health Sector Transformation Plan

Hgbs Hemoglobin s

HRp2 Histidine Rich Protein 2

HO Hospital

HIV Human Immunodeficiency Virus

HLA Human Leukocyte Antigen

IRS Indoor Residual Spray

vi

IEC /BCC Information Education Communication / Behavioral Change

Communication

ITN Insecticide Treated Net

ITNs Insecticide Treated Nets

IRT Integrated Refresher Training

IQC Internal Quality Control

IM Intra Muscular

IV Intravenous

LMIS Logistic Management Information System

LLIN Long Lasting Insecticide Nets

MIS Malaria Indicator Survey

M&E

µl

Monitoring and Evaluation

Micro Liter

NEQAS National External Quality Assessment Scheme

NPSP National Malaria Strategic Plan

PMA Pan-Malaria Antigen

PPE Personal Protective Equipment

Pf Plasmodium Falciparum

PfHRP2 Plasmodium Falciparum Histidine Rich Protein

PLDH Plasmodium Lactate Dehydrogenase

Pm Plasmodium Malaria

Po Plasmodium Ovale

Pv Plasmodium Vivax

PCR Polymerase Chain Reaction

PMI President’s Malaria Initiative

PT Proficiency Test

PHL Public Health Laboratory

QA Quality Assurance

QC Quality Control

QBC Quantitative Buffy Coat

RDTs Rapid Diagnostic Tests

RBC Red Blood Cell

REQAS Regional External Quality Assessment Scheme

vii

RHB Regional Health Bureau

RL Regional Laboratory

SEM Scanning Electron Microscope

SDPs Service Delivery Points

SCM Severe and Complicated Malaria

SOP Standard Operational Procedure

SP Sulphadoxine Pyrimethamine

USAID The United States Agency for International Development

Rx Treatment

WBC White Blood Cell

WHO World Health Organization

viii

II. Glossary

Anopheles, infected: - Female Anopheles mosquitoes with detectable malaria

parasites.

Anopheles, infective: - Female Anopheles mosquitoes with sporozoites in the

salivary glands

Antibody: - A specialized serum protein (immunoglobulin or gamma globulin)

produced by B lymphocytes in the blood in response to an exposure to foreign

proteins (antigens). The antibodies specifically bind to the antigens that induced the

immune response. Antibodies help defend the body against infectious agents,

including bacteria, viruses, or parasites.

Antigen: - Any substance that stimulates the immune system to produce antibodies.

Antigens are often foreign substances: invading bacteria, viruses, or parasites

Artemisinin-based combination therapy (ACT): - A combination of an Artemisinin

derivative with a longer-acting antimalarial drug that has a different mode of action.

Asexual cycle: -The life-cycle of the malaria parasite in host from merozoite invasion

of red blood cells to schizonts rupture (merozoite → ring stage → trophozoites →

schizonts → merozoites). Duration approximately 48 h in Plasmodium falciparum,

P. ovale and P. vivax; 72 h in P. malariae.

Asexual parasitemia: - The presence in host red blood cells of asexual parasites. The

level of asexual parasitaemia can be expressed in several different ways: the

percentage of infected red blood cells, the number of infected cells per unit volume

of blood, the number of parasites seen in one microscopic field in a high-power

examination of a thick blood film, or the number of parasites seen per 200– 1000

white blood cells in a high power examination of a thick blood film.

Case confirmed: - Malaria case (or infection) in which the parasite has been detected

in a diagnostic test, i.e. microscopy, a rapid diagnostic test or a molecular diagnostic

test Note: On rare occasions, the presence of occult malaria infection in a blood or

organ donor is confirmed retrospectively by the demonstration of malaria parasites

in the recipient of the blood or organ.

Case imported: - Malaria case or infection in which the infection was acquired

outside the area in which it is diagnosed.

Case, indigenous: - A case contracted locally with no evidence of importation and

no direct link to transmission from an imported case

Case relapsing: - Malaria case attributed to activation of hypnozoites of P. vivax or

P. ovale acquired previously

ix

Note: The latency of a relapsing case can be > 6–12 months. The

occurrence of relapsing cases is not an indication of operational failure,

but their existence should lead to evaluation of the possibility of ongoing

transmission

Case suspected malaria: - Illness suspected by a health worker to be due to malaria,

generally on the basis of the presence of fever with or without other symptoms.

Cerebral malaria: - Severe P. falciparum malaria with impaired consciousness

(Glasgow coma scale < 11, Blantyre coma scale < 3) persisting for > 1 hour after a

seizure Note: The initial neurological symptoms are often drowsiness, confusion,

failure to eat or drink or convulsions (see current WHO definition of severe malaria

in the Guidelines for the treatment of malaria. 2015, Third edition).

Control: - Reduction of disease incidence, prevalence, morbidity or mortality to a

locally acceptable level as a result of deliberate efforts.

Drug efficacy: - Capacity of an antimalarial medicine to achieve the therapeutic

objective when administered at a recommended dose, which is well tolerated and

has minimal toxicity

Drug resistance: - The ability of a parasite strain to survive and/or to multiply despite

the administration and absorption of a medicine given in doses equal to or higher

than those usually recommended but within the tolerance of the subject, provided

drug exposure at the site of action is adequate. Resistance to anti malarias arises

because of the selection of parasites with genetic mutations or gene amplifications

that confer reduced susceptibility (WHO).

Efficacy: - The power or capacity to produce a desired effect

Elimination: - The interruption of local mosquito-borne malaria transmission in a

defined geographical area, creating a zero incidence of locally contracted cases.

Imported cases will continue to occur and continued intervention measures are

required.

Elimination of disease: - Reduction to zero of the incidence of a specified disease in

a defined geographical area as a result of deliberate efforts.

Elimination of infection: - Reduction to zero of the incidence of infection caused by

a specified agent in a defined geographical area as a result of deliberate efforts.

Endemic: - Where disease occurs consistently.

Epidemic: - The occurrence of more cases of disease than expected in a given area

or among a specific group of people over a particular period of time.

x

Epidemiology: - The study of the distribution and determinants of health-related

states or events in specified populations; the application of this study to control

health problems.

Eradication: - Permanent reduction to zero of the worldwide incidence of infection

caused by a specific agent as a result of deliberate efforts;

Erythrocytic stage: - A stage in the life cycle of the malaria parasite found in the red

blood cells. Erythrocytic stage parasites cause the symptoms of malaria.

Exoerythrocytic stage: - A stage in the life cycle of the malaria parasite found in liver

cells (hepatocytes). Exoerythrocytic stage parasites do not cause symptoms.

External quality assessment: - A system whereby a reference laboratory sends

stained blood films to a laboratory for examination. The laboratory receiving the

slides is not informed of the correct result of the slides until the laboratory has

reported their findings back to the reference laboratory.

False negative slide: - A positive smear that is misread as negative.

False positive slide: - A negative smear that is misread as positive.

Feedback: - The process of communicating results of external quality control to the

original laboratory, including identification of errors and recommendations for

remedial action.

G6PD deficiency: - An inherited abnormality that causes the loss of a red blood cell

enzyme. People who are G6PD deficient should not take the antimalarial drug

primaquine.

Gametocyte: - The sexual stage of malaria parasites. Male gametocytes

(microgametocytes) and female gametocytes (macro gametocytes) are inside red

blood cells in the circulation. If a female Anopheles mosquito ingests them, they

undergo sexual reproduction, which starts the extrinsic (sporogonic) cycle of the

parasite in the mosquito. Gametocytes of Plasmodium falciparum are typically

banana or crescent-shaped (from the Latin falcis = sickle).

Hypnozoite: - Dormant form of malaria parasites found in liver cells. Hypnozoite

occur only with Plasmodium vivax and P. ovale. After sporozoites (inoculated by the

mosquito) invade liver cells, some sporozoites develop into dormant forms (the

Hypnozoite), which do not cause any symptoms. Hypnozoite can become activated

months or years after the initial infection, producing a relapse.

Hypoglycemia: - Low blood glucose; can occur with malaria. In addition, treatment

with quinine and quinidine stimulate insulin secretion, reducing blood glucose.

xi

Immune system: - The cells, tissues, and organs that help the body resist infection

and disease by producing antibodies and/or cells that inhibit the multiplication of

the infectious agent.

Immunity: - Protection generated by the body’s immune system, in response to

previous malaria attacks, resulting in the ability to control or lessen a malaria attack.

Incubation period: - The interval of time between infection by a microorganism and

the onset of the illness or the first symptoms of the illness. With malaria, the

incubation is between the mosquito bite and the first symptoms. Incubation periods

range from 7 to 40 days, depending on the species.

Indigenous malaria: - Mosquito-borne transmission of malaria in a geographic area

where malaria occurs regularly.

Infection: - The invasion of an organism by a pathogen, such as bacteria, viruses, or

parasites. Some, but not all, infections lead to disease.

Introduced malaria: - Mosquito-borne transmission of malaria from an imported

case in a geographic area where malaria does not regularly occur.

Malaria pigment (haemozoin): - A dark brown granular pigment formed by malaria

parasites as a by-product of haemoglobin catabolism. The pigment is evident in

mature trophozoites and schizonts. They may also be present in white blood cells

(peripheral monocytes and polymorph nuclear neutrophils) and in the placenta.

Merozoite: - A daughter-cell formed by asexual development in the life cycle of

malaria parasites. Liver-stage and blood-stage malaria parasites develop into

schizonts, which contain many merozoites. When the schizonts are mature, they

(and their host cells!) rupture, the merozoites are released and infect red blood cells.

Microscopists: - A person who uses a microscope to read blood films to aid or

confirm the diagnosis of malaria and reports on their findings. The term is used in

this manual to include personnel at all levels of a malaria programme involved in

this work, from professors involved in teaching and research to village health

volunteers specifically trained in malaria microscopy.

Oocyst: - A stage in the life cycle of malaria parasites, oocysts are rounded cysts

located in the outer wall of the stomach of mosquitoes. Sporozoites develop inside

the oocysts. When mature, the oocysts rupture and release the sporozoites, which

then migrate into the mosquito’s salivary glands, ready for injection into the human

host.

Outbreak: - An epidemic limited to a localized increase in disease incidence, e.g. in

a village, town or closed institution.

xii

Pandemic: - An epidemic occurring over a very wide area, crossing international

boundaries and usually affecting a large number of people.

Parasite: - Any organism that lives in or on another organism without benefiting the

host organism; commonly refers to pathogens, most commonly to protozoans and

helminthes.

Parasitemia: - The presence of parasites in the blood. The term can also be used to

express the quantity of parasites in the blood (for example, a parasitemia of 2

percent).

Paroxysm: - A sudden attack or increase in intensity of a symptom, usually occurring

at intervals

Pathogen: - Bacteria, viruses, parasites, or fungi that can cause disease.

Plasmodium: - The genus of the parasite that causes malaria. The genus includes

four species that infect humans: Plasmodium falciparum, Plasmodium vivax,

Plasmodium ovale, and Plasmodium malariae.

Pre-erythrocytic development: - The life-cycle of the malaria parasite when it first

enters the host. Following inoculation into a human by the female anopheline

mosquito, sporozoites invade parenchyma cells in the host liver and multiply within

the hepatocytes for 5–12 days, forming hepatic schizonts. These then burst

liberating merozoites into the bloodstream, which subsequently invade red blood

cells

Presumptive treatment: - Treatment of clinically suspected cases without, or prior

to, results from confirmatory laboratory tests

Panel testing: - The process by which laboratories (known as the “test laboratories”)

performs malaria microscopy on a set of prepared slides received from the National

and Regional Laboratories. This exercise can check both the laboratories’ staining

quality as well as the ability of technicians to recognize and identify malaria

parasites present.

Quality assurance: - The monitoring and maintenance of high accuracy, reliability

and efficiency of laboratory services. Quality assurance addresses all factors that

affect laboratory performance including test performance (quality control, internal

and external) equipment and reagent quality, workload, workplace conditions,

training and laboratory staff support.

Quality control: - Measures the quality of a test or a reagent. For malaria microscopy,

the most common form of quality control (QC) is the cross-checking of routine blood

slides to monitor the accuracy of examination. Quality control also encompasses

external quality control and reagent quality control. Crosschecking QC is a system

xiii

whereby a sample of routine blood slides are crosschecked for accuracy by a

supervisor or the regional/national laboratory. Reagent QC is a system of formally

monitoring the quality of the reagents used in the laboratory.

Quality Improvement: - A process by which the components of microscopy and RDT

diagnostic services are analyzed with the aim of identifying and permanently

correcting any deficiencies. Data collection, data analysis, and creative problem

solving are skills used in this process.

Radical cure (also radical treatment): - Complete elimination of malaria parasites

from the body; the term applies specifically to elimination of dormant liver stage

parasites (Hypnozoite) found in Plasmodium vivax and P. ovale.

Recrudescence: - A repeated attack of malaria (short-term relapse or delayed), due

to the survival of malaria parasites in red blood cells. It is recurrence of asexual

parasitaemia of the same genotype(s) that caused the original illness, due to

incomplete clearance of asexual parasites after antimalarial treatment.

Note: - Recrudescence is different from reinfection with a parasite of the

same or different genotype(s) and relapse in P. vivax and P. ovale

infections.

Recurrence: - Reappearance of asexual parasitaemia after treatment, due to

recrudescence, relapse (in P. vivax and P. ovale infections only) or a new infection

Relapse: - Recurrence of disease after it has been apparently cured. In malaria, true

relapses are caused by reactivation of dormant liver stage parasites (Hypnozoite)

found in Plasmodium vivax and P. ovale.

Residual insecticide spraying: - Spraying insecticides that have residual efficacy

(that continue to affect mosquitoes for several months) against houses where

people spend nighttime hours. Residual insecticide spraying is done to kill

mosquitoes when they come to rest on the walls, usually after a blood meal.

Resistance: - The ability of an organism to develop strains that are impervious to

specific threats to their existence. The malaria parasite has developed strains that

are resistant to drugs, such as chloroquine. The Anopheles mosquito has developed

strains that are resistant to DDT and other insecticides.

Ring stage: - Young usually ring-shaped intra-erythrocytic malaria parasites, before

malaria pigment is evident under microscopy

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Schizogony: - Asexual reproductive stage of malaria parasites. In red blood cells,

schizogony entails development of a single trophozoites into numerous merozoites;

a similar process happens in infected liver cells.

Schizonts: - A developmental form of the malaria parasite that contains many

merozoites. Schizonts are seen in the liver-stage and blood-stage parasites.

Serology: - The branch of science dealing with the measurement and

characterization of antibodies and other immunological substances in body fluids,

particularly serum.

Slide positivity rate: - The proportion of positive slides, detected by microscopy,

among all those examined within a laboratory over a defined period of time.

Sporozoites: - A stage in the life cycle of the malaria parasite. Sporozoites, produced

in the mosquito, migrate to the mosquito's salivary glands. They can be inoculated

into a human host when the mosquito takes a blood meal on the human. In the

human, the sporozoites enter liver cells where they develop into the next stage of

the malaria parasite life cycle (the liver stage or exo-erythrocytic stage).

Trophozoites: - A developmental form during the blood stage of malaria parasites.

After merozoites have invaded the red blood cell, they develop into trophozoites

(sometimes, early trophozoites are called rings or ring stage parasites); trophozoites

develop into schizonts.

Vector: - An organism (for example, Anopheles mosquitoes) that transmits an

infectious agent (for example, malaria parasites) from one host to the other (for

example, humans)

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III. Scope and Purpose of the Manual

A. Purpose

The purpose of this manual is to guide professionals and stakeholders responsible

for malaria control and prevention programs on the best ways of ensuring quality

laboratory diagnosis and Treatment. The manual describes overview of malaria

epidemiology, laboratory procedure, quality assurance and supply management;

laboratory safety and outlines the technical knowledge needed for laboratory

diagnosis of malaria. The aim of this manual is to help to ensure that malaria

diagnosis at national, regional, district and community levels are efficiently and

effectively organized to allow early diagnosis and prompt, effective treatment.

The manual provides basic information for the successful operation of malaria

laboratory diagnosis and defines the skills required in the following areas:

• Implementation of quality assured malaria laboratory diagnosis through

standard procedure

• Planning training and conducting quality assurance program

• Planning effective lab diagnosis and identifying the technical and managerial

elements that require revision

• Logistical organization to ensure regular supplies

• Planning supervision, monitoring and evaluation

• Coordinating and integrating malaria diagnosis with other laboratory programs

B. Target Audience

The manual is intended for use in particular by health professionals and

stakeholders working on malaria laboratory diagnosis program, and in general for

multidisciplinary teams involved in managing national malaria control program,

including program managers, epidemiologists, program supervisors, health

educators, logistics officers and trainers. Health project managers dealing with

malaria at national, district and community levels, including those responsible for

private health services, will also find this manual useful. The manual will be a useful

resource in Ministry of Health or in projects supported by international and

multilateral cooperation agencies or nongovernmental organizations, in medical,

nursing, laboratory and public health schools for training in effective malaria case

management and helping researchers to guide and follow standard malaria

laboratory diagnosis techniques and procedures.

1

CHAPTER ONE: Introduction to Malaria

Chapter Description:

This chapter describes Malaria etiology, transmission and Lifecycle, Malaria

diagnostic methods, Malaria global burden, Malaria situation in Ethiopia and

National Strategic Plan for Malaria Prevention, Control and Elimination in Ethiopia.

1.1. Malaria Etiology

Malaria is a disease caused by obligate intra-cellular protozoan blood parasites of

the Plasmodium species and transmitted to humans by the bite of infected female

Anopheles mosquitoes. Malaria trophozoites may also be transmitted through

blood transfusion and trans-parentally (congenital malaria). The life cycle follows

three stages: the exoerythrocytic, erythrocytic and sporogonic cycle. There are

approximately 156 named species of Plasmodium which infect various species of

vertebrates. There are four different human malaria species (P. falciparum, P. vivax,

P. malariae and P. ovale), of which P. falciparum and P. vivax are the most prevalent

and P. falciparum the most dangerous. Recently, a new malaria parasite species

named P. knowlesi is identified in Asia affecting both humans and animals. Malaria

can be very severe and can lead to death if left untreated. Malaria parasite is

transmitted from an infected person to another by the bite of a female anopheline

mosquito. This can occur only after the parasite has been inside the mosquito for at

least a week.

Female mosquitoes take blood meals to carry out egg production, and such blood

meals are the link between the human and the mosquito hosts in the parasite life

cycle. The successful development of the malaria parasite in the mosquito (from the

“gametocyte” stage to the “sporozoites” stage) depends on several factors. The

most important is ambient temperature and humidity (higher temperatures

accelerate the parasite growth in the mosquito) and whether

the Anopheles survives long enough to allow the parasite to complete its cycle in

the mosquito host (“sporogonic” or “extrinsic” cycle, duration 10 to 18 days).

Differently from the human mosquito host does not suffer noticeably from the

presence of the parasites.

2

1.2. Life Cycle and Mode of Transmission of Plasmodium

The natural ecology of malaria involves malaria parasites infecting successively two

types of hosts: humans (intermediate host) and female Anopheles mosquitoes

(definitive host). In humans, the parasites grow and multiply first in the liver cells

and then in the red cells of the blood. In the blood, successive broods of parasites

grow inside the red cells and destroy them, releasing daughter parasites

(“merozoite”) that continue the cycle by invading other red cells.

The blood stage parasites are those that cause the symptoms of malaria. When

certain forms of blood stage parasites (“gametocytes”) infective stage to mosquito

are picked up by a female Anopheles mosquito during a blood meal, they start

another, different cycle of growth and multiplication in the mosquito. After 10-18

days, the parasites are found (as “sporozoites”) in the mosquito’s salivary glands.

When the Anopheles mosquito takes a blood meal on another human, the

sporozoites (infective stage to human being) are injected with the mosquito’s saliva

and start another human infection when they parasitize the liver cells.

Thus, the mosquito carries the disease from one human to another (acting as a

“vector”). Differently from the human host, the mosquito vector does not suffer

from the presence of the parasites.

Figure 1. 1. Life cycle of plasmodium; Source: http://www.dpd.cdc.gov/dpdx

3

The malaria parasite life cycle involves two hosts. During a blood meal, a malaria-

infected female Anopheles mosquito inoculates sporozoites into the human

host . Sporozoites infect liver cells and mature into schizonts, which rupture

and release merozoite . (Of note, in P. vivax and P. ovale a dormant stage

(Hypnozoite) can persist in the liver and cause relapses by invading the bloodstream

weeks, or even years later.) After this initial replication in the liver (exo-erythrocytic

schizogony ), the parasites undergo asexual multiplication in the erythrocytes

(erythrocytic schizogony ). Merozoites infect red blood cells . The ring stage

trophozoites mature into schizonts, which rupture releasing merozoites . Some

parasites differentiate into sexual erythrocytic stages (gametocytes) . Blood stage

parasites are responsible for the clinical manifestations of the disease. The

gametocytes, male (microgametocytes) and female (macro gametocytes), are

ingested by an Anopheles mosquito during a blood meal . The parasites’

multiplication in the mosquito is known as the sporogonic cycle . While in the

mosquito’s stomach, the microgametes penetrate the macrogametes generating

zygotes . The zygotes in turn become motile and elongated (ookinetes) which

invade the midget wall of the mosquito where they develop into oocysts . The

oocysts grow, rupture, and release sporozoites , which make their way to the

mosquito’s salivary glands. Inoculation of the sporozoites into a new human host

perpetuates the malaria life cycle.

1.3 Malaria Parasites

The species infecting humans are:

• P. falciparum, which is found worldwide in tropical and subtropical areas, and

especially in Africa where this species predominates. P. falciparum can cause

severe malaria because it multiples rapidly in the blood, and can thus cause

severe blood loss (anemia). In addition, the infected parasites can clog small

blood vessels. When this occurs in the brain, cerebral malaria results, a

complication that can be fatal.

• P. vivax, which is found mostly in Asia, Latin America, and in some parts of

Africa. Because of the population densities especially in Asia it is probably

the most prevalent human malaria parasite. P. vivax (as well as P. ovale) has

dormant liver stages (“Hypnozoite”) that can activate and invade the blood

(“relapse”) several months or years after the infecting mosquito bite.

4

• P. ovale is found mostly in Africa (especially West Africa) and the islands of

the western Pacific. It is biologically and morphologically very similar to P.

vivax. However, differently from P. vivax, it can infect individuals who are

negative for the Duffy blood group, which is the case for many residents of

sub-Saharan Africa. This explains the greater prevalence of P. ovale (rather

than P. vivax) in most of Africa.

• P. malariae, found worldwide, is the only human malaria parasite species

that has a quartan cycle (three-day cycle). (The three other species have a

tertian, two-day cycle.) If untreated, P. malariae causes a long-lasting,

chronic infection that in some cases can last a lifetime. In some chronically

infected patients P. malariae can cause serious complications such as the

nephrotic syndrome.

• P. knowlesi is found throughout Southeast Asia as a natural pathogen of

long-tailed and pig-tailed macaques. It has recently been shown to be a

significant cause of zoonotic malaria in that region, particularly in

Malaysia. P. knowlesi has a 24-hour replication cycle and so can rapidly

progress from an uncomplicated to a severe infection; fatal cases have been

reported.

Infection with malaria parasites may result in a wide variety of symptoms, ranging

from absent or very mild symptoms to severe disease and even death. Malaria

disease can be categorized as uncomplicated or severe (complicated). In general,

malaria is a curable disease if diagnosed and treated promptly and correctly.

Despite being preventable and treatable, malaria continues to have a devastating

impact on people’s health and livelihoods around the world.

All the clinical symptoms associated with malaria are caused by the asexual

erythrocytic or blood stage parasites. When the parasite develops in the erythrocyte,

numerous known and unknown waste substances such as hemozoin pigment and

other toxic factors accumulate in the infected red blood cell. These are dumped into

the bloodstream when the infected cells lyse and release invasive merozoites. The

hemozoin and other toxic factors such as glucose phosphate isomerase (GPI)

stimulate macrophages and other cells to produce cytokines and other soluble

factors which act to produce fever and rigors and probably influence other severe

pathophysiology associated with malaria.

5

Plasmodium falciparum-infected erythrocytes, particularly those with mature

trophozoites, adhere to the vascular endothelium of venular blood vessel walls and

do not freely circulate in the blood. When this sequestration of infected erythrocytes

occurs in the vessels of the brain it is believed to be a factor in causing the severe

disease syndrome known as cerebral malaria, which is associated with high

mortality.

1.3.1 Incubation Period

Following the infective bite by the Anopheles mosquito, a period of time (the

“incubation period”) goes by before the first symptoms appear. The incubation

period in most cases varies from 7 to 30 days. The shorter periods are observed

most frequently with P. falciparum and the longer ones with P. malariae.

Anti-malarial drugs taken for prophylaxis by travelers can delay the appearance of

malaria symptoms by weeks or months, long after the traveler has left the malaria-

endemic area. (This can happen particularly with P. vivax and P. ovale, both of

which can produce dormant liver stage parasites; the liver stages may reactivate

and cause disease months after the infective mosquito bite.)

Such long delays between exposure and development of symptoms can result in

misdiagnosis or delayed diagnosis because of reduced clinical suspicion by the

health-care provider. Returned travelers should always remind their health-care

providers of any travel in areas where malaria occurs during the past 12 months.

1.3.2 Uncomplicated Malaria

Uncomplicated malaria: - is characterized by fever and other features including

chills, profuse sweating, muscle pains, joint pains, headache, abdominal pain,

diarrhea, nausea, vomiting, loss of appetite, irritability, and refusal to feed (in

infants). These features may occur singly or in combination and are due to the

presence of asexual forms of the parasites in the blood. Classically (but infrequently

observed) the attacks occur every second day with the “tertian” parasites (P.

falciparum, P. vivax, and P. ovale) and every third day with the “quartan” parasite

(P. malariae).

More commonly, the patient presents with a combination of the following

symptoms:

In countries where cases of malaria are infrequent, these symptoms may be

attributed to influenza, a cold, or other common infections, especially if malaria is

6

not suspected. Conversely, in countries where malaria is frequent, residents often

recognize the symptoms as malaria and treat themselves without seeking diagnostic

confirmation (“presumptive treatment”).

The classical (but rarely observed) malaria attack lasts 6-10 hours. It consists of:

• A cold stage (sensation of cold, shivering)

• A hot stage (fever, headaches, vomiting; seizures in young children) and

finally

• A sweating stage (sweats, return to normal temperature, tiredness).

Physical findings may include:

• Elevated temperatures

• Perspiration

• Weakness

• Enlarged spleen

• Mild jaundice

• Enlargement of the liver

• Increased respiratory rate

1.3.3 Complicated (Severe Malaria)

Severe and complicated malaria is a life threatening condition, defined as the

detection of P. falciparum in peripheral blood together with any of the following

clinical or laboratory features (singly or in combination):

• Inability to or difficulty in sitting upright; standing or walking without support; or

inability to feed (in an infant)

• Alteration in the level of consciousness (ranging from drowsiness to deep coma)

• Cerebral malaria (unarousable coma not attributable to any other cause, other

neurological signs)

• Respiratory distress

• Multiple generalized convulsions (2 or more episodes within a 24-hour period)

• Circulatory collapse (shock, septicemia)

• Pulmonary oedema

• Abnormal bleeding (Disseminated Intravascular Coagulation – DIC)

• Jaundice

• Hemoglobinuria (black water fever)

7

• Acute renal failure – presenting as oliguria (passing scanty urine) or anuria (not

passing urine)

• Severe anemia (haemoglobin <5g/dl or hematocrit < 15%)

• Hypoglycemia (blood glucose level < 2.2 mmol/l)

• Hyperparasitaemia (parasitaemia of >200,000/μl - in patients from high

transmission areas; or 100,000/μl in patients from low transmission areas)

• hyperlactatemia (whole blood lactate >5 mmol/l)

Examples of illnesses that may present with symptoms and signs similar to malaria

include:

• Meningitis

• Otitis media

• Pharyngo-tonsillitis

• Pneumonia

• Acute gastroenteritis

• Typhoid fever

• Urinary tract infection

• Viral infections (e.g. mumps, measles)

• Hepatitis

Complicated malaria occurs when infections are complicated by serious organ

failures or abnormalities in the patient’s blood or metabolism. The manifestations

of severe malaria include.

• Cerebral malaria, with abnormal behavior, impairment of consciousness,

seizures, coma, or other neurologic abnormalities

• Severe anemia due to hemolysis (destruction of the red blood cells)

• Hemoglobinuria (hemoglobin in the urine) due to hemolysis

• Acute respiratory distress syndrome (ARDS), an inflammatory reaction in the

lungs that inhibits oxygen exchange, which may occur even after the parasite

counts have decreased in response to treatment

• Abnormalities in blood coagulation

• Low blood pressure caused by cardiovascular collapse

• Acute kidney failure

• Hyper parasitemia, where more than 5% of the red blood cells are infected by

malaria parasites

8

• Metabolic acidosis (excessive acidity in the blood and tissue fluids), often in

association with hypoglycemia

• Hypoglycemia (low blood glucose). Hypoglycemia may also occur in

pregnant women with uncomplicated malaria, or after treatment with

quinine.

Severe malaria is a medical emergency and should be treated urgently and

aggressively.

1.3.4. Malaria Relapses

In P. vivax and P. ovale infections, patients having recovered from the first episode

of illness may suffer several additional attacks (“relapses”) after months or even

years without symptoms. Relapses occur because P. vivax and P. ovale have

dormant liver stage parasites (“Hypnozoite”) that may reactivate.

Relapse: Recurrence of disease after it has been apparently cured. In malaria, true

relapses are caused by reactivation of dormant liver stage parasites (hypnozoites)

found in Plasmodium vivax and P. ovale. Treatment to reduce the chance of such

relapses is available and should follow treatment of the first attack

1.3.5. Recrudescence

Plasmodium falciparum malaria recurrences after a complete treatment can occur

by two different mechanisms, reinfection or recrudescence.

Recrudescence: A repeated attack of malaria due to the survival of malaria parasites

in red blood cells. It can be due to a) incomplete or inadequate treatment as a result

of drug resistance or improper choice of medication b) an antigenic variation c)

infection by different strains. Recrudescence result from persistent erythrocytic

infection, which re-emerges within a defined period following antimalarial

treatment. Symptoms of malaria can recur after varying symptom-free periods.

Depending upon the cause, recurrence can be classified as

either recrudescence, relapse, or reinfection. Recrudescence is when symptoms

return after a symptom-free period. It is caused by parasites surviving in the blood

as a result of inadequate or ineffective treatment. Recurrence of asexual

parasitaemia of the same genotype(s) that caused the original illness, due to

incomplete clearance of asexual parasites after antimalarial treatment.

9

Note: Recrudescence is different from reinfection with a parasite of the

same or different genotype(s) and relapse in P. vivax and P. ovale

infections

1.3.6. Malaria Reinfection

is a new infection that follows a primary infection; can be distinguished from

recrudescence by the parasite genotype, which is often (but not always) different

from that which caused the initial infection

1.3.7. Other Manifestations of Malaria

• Neurologic defects may occasionally persist following cerebral malaria, especially

in children. Such defects include trouble with movements (ataxia), palsies, speech

difficulties, deafness, and blindness.

• Recurrent infections with P. falciparum may result in severe anemia. This occurs

especially in young children in tropical Africa with frequent infections that are

inadequately treated.

• Malaria during pregnancy (especially P. falciparum) may cause severe disease in

the mother, and may lead to premature delivery or delivery of a low-birth-weight

baby.

• On rare occasions, P. vivax malaria can cause rupture of the spleen.

• Nephrotic syndrome (a chronic, severe kidney disease) can result from chronic or

repeated infections with P. malariae.

• Hyper reactive malarial splenomegaly (also called “tropical splenomegaly

syndrome”) occurs infrequently and is attributed to an abnormal immune

response to repeated malarial infections. The disease is marked by a very

enlarged spleen and liver, abnormal immunologic findings, anemia, and a

susceptibility to other infections (such as skin or respiratory infections.

1.4. Human Factors Resistance to Malaria

1.4.1. Genetic Factors

Biologic characteristics present from birth can protect against certain types of

malaria. Two genetic factors, both associated with human red blood cells, have been

shown to be epidemiologically important. Persons who have the sickle cell trait

(heterozygotes for the abnormal hemoglobin gene HbS) are relatively protected

against P. falciparum malaria and thus enjoy a biologic advantage. Because P.

falciparum malaria has been a leading cause of death in Africa since remote times,

the sickle cell trait is now more frequently found in Africa and in persons of African

10

ancestry than in other population groups. In general, the prevalence of hemoglobin-

related disorders and other blood cell dyscrasias, such as Hemoglobin C, the

thalassemia’s and G6PD deficiency, are more prevalent in malaria endemic areas

and are thought to provide protection from malarial disease.

Persons who are negative for the Duffy blood group have red blood cells that are

resistant to infection by P. vivax. Since the majority of Africans are Duffy negative, P.

vivax is rare in Africa south of the Sahara, especially West Africa. In that area, the

niche of P. vivax has been taken over by P. ovale, a very similar parasite that does

infect Duffy-negative persons.

Other genetic factors related to red blood cells also influence malaria, but to a lesser

extent. Various genetic determinants (such as the Human leukocyte antigen “HLA

complex,” which plays a role in control of immune responses) may equally influence

an individual’s risk of developing severe malaria.

1.4.2. Acquired Immunity

Acquired immunity greatly influences how malaria affects an individual and a

community. After repeated attacks of malaria a person may develop a partially

protective immunity. Such “semi-immune” persons often can still be infected by

malaria parasites but may not develop severe disease, and, in fact, frequently lack

any typical malaria symptoms.

In areas with high P. falciparum transmission (most of Africa south of the Sahara),

newborns will be protected during the first few months of life presumably by

maternal antibodies transferred to them through the placenta. As these antibodies

decrease with time, these young children become vulnerable to disease and death

by malaria. If they survive repeated infections to an older age (2-5 years) they will

have reached a protective semi-immune status. Thus in high transmission areas,

young children are a major risk group and are targeted preferentially by malaria

control interventions.

In areas with lower transmission (such as Asia and Latin America), infections are

less frequent and a larger proportion of the older children and adults have no

protective immunity. In such areas, malaria disease can be found in all age groups,

and epidemics can occur.

11

1.5. Methods for Malaria Diagnosis

Diagnosis of malaria depends on the demonstration of parasites in the blood,

usually by microscopy. Additional laboratory findings may include mild anemia,

mild decrease in blood platelets (thrombocytopenia), elevation of bilirubin, and

elevation of aminotransferases.

Once malaria is suspected on clinical grounds, it is mandatory to obtain the

laboratory confirmation of the presence of malaria parasites. Clinicians could

request for diagnostic test for malaria to confirm the diagnosis of malaria in a patient

with symptoms and signs suggestive of malaria disease; to rule out malaria

infection in a patient with other known causes of fever; to confirm malaria in febrile

infants under 3 months of age; to look for treatment failure; and to investigate

causes of anemia, jaundice or splenomegaly.

1.5.1. Clinical Diagnosis of Malaria

Clinical diagnosis is based on the patient’s symptoms and physical findings at

examination, and the only approach where laboratory support doesn't exist. The

first symptoms of malaria (most often fever, chills, sweats, headaches, muscle

pains, nausea and vomiting) are often not specific and are also found in other

diseases (such as the “flu” and common viral infections). Likewise, the physical

findings are often not specific (elevated temperature, perspiration, tiredness).

In severe malaria (caused by Plasmodium falciparum), clinical findings (confusion,

coma, neurologic focal signs, severe anemia, respiratory difficulties) are more

striking and may increase the index of suspicion for malaria. Basically, clinical

finding alone is unreliable and should always be confirmed by a laboratory test for

malaria. In addition to ordering the malaria specific diagnostic tests described

below, the health-care provider should conduct an initial workup and request a

complete blood count and a routine chemistry panel. In the event that the person

does have a positive malaria test, these additional tests will be useful in determining

whether the patient has uncomplicated or severe manifestations of the malaria

infection. Specifically, these tests can detect severe anemia, hypoglycemia, renal

failure, hyperbilirubinemia, and acid-base disturbances.

12

1.5.2. Laboratory Diagnosis of Malaria

Once malaria is suspected on clinical grounds, it is mandatory to obtain the

laboratory confirmation of the presence of malaria parasites. Clinicians could

request for diagnostic test for malaria to confirm the diagnosis of malaria in a patient

with symptoms and signs suggestive of malaria disease; to rule out malaria

infection in a patient with other known causes of fever; to confirm malaria in febrile

infants under 3 months of age; to look for treatment failure; and to investigate

causes of anemia, jaundice or splenomegaly.

1.5.2.1. Common Diagnostic Methods

The two laboratory diagnostic methods or tools most often used for confirming a

diagnosis of malaria are:

Microscopy

Malaria parasites can be detected, identified and quantified by examining stained

blood film under the microscope. This technique remains the gold standard for

laboratory confirmation of malaria. However, it depends on the quality of the

reagents, of the microscope, and on the experience of the laboratory personnel.

When used stained thick and thin blood film, Microscopy remains the “gold

standard” for laboratory confirmation of malaria parasites. It can detect to the

lowest 5 parasites per micro liter of blood. This test should be performed

immediately when ordered by a health-care provider. They should not be saved for

the most qualified staff to perform or batched for convenience. In addition, these

tests should not be sent out to reference laboratories with results available only days

to weeks later. It is vital that health-care providers receive results from these tests

within hours in order to appropriately treat their patients infected with malaria.

Advantages

Microscopy is an established, relatively simple technique that is familiar to most

laboratorians. Any laboratory that can perform routine hematology tests is

equipped to perform a thin and thick malaria smear. Within a few hours of collecting

the blood, the microscopy test can provide valuable information. First and foremost,

it can determine that malaria parasites are present in the patient’s blood. Once the

diagnosis is established – usually by detecting parasites in the thick smear – the

laboratorian can examine the thin smear to determine the malaria species and the

parasitemia, or the percentage of the patient’s red blood cells that are infected with

13

malaria parasites. The thin and thick smears are able to provide all 3 of these vital

pieces of information to the doctor to guide the initial treatment decisions that need

to be made acutely.

Disadvantages

Microscopy results are only as reliable as the laboratories performing the tests.

Those laboratorians who does not perform this test regularly, and may not be

maintaining optimal proficiency.

Rapid Diagnostic Test

RDTs: RDTs detect antigens (proteins produced by malaria parasite) in the blood of

a patient with malaria. Various test kits are available to detect antigens derived from

malaria parasites. Such immunologic (“immunochromatographic”) tests most often

use a dipstick or cassette format, and provide results in 2-15 minutes. These “Rapid

Diagnostic Tests” (RDTs) offer a useful alternative to microscopy in situations where

reliable microscopic diagnosis is not available. Malaria RDTs are currently used in

some clinical settings and programs.

A Rapid Diagnostic Test (RDT) is an alternate way of quickly establishing the

diagnosis of malaria infection by detecting specific malaria antigens in a person’s

blood.

Technique

A blood specimen collected from the patient is applied to the sample pad on the test

card along with certain reagents. After 15 minutes, the presence of specific bands in

the test card window indicate whether the patient is infected with Plasmodium

falciparum or one of the other 3 species of human malaria. It is recommended that

the laboratory maintain a supply of blood containing P. falciparum for use as a

positive control.

Advantages

High-quality malaria microscopy is not always immediately available in every

clinical setting where patients might seek medical attention. Although this practice

is discouraged, many healthcare settings either save blood samples for malaria

microscopy until a qualified person is available to perform the test, or send the

blood samples to commercial or reference laboratories. These practices have

14

resulted in long delays in diagnosis. The laboratories associated with these health-

care settings may now use an RDT to more rapidly determine if their patients are

infected with malaria.

Disadvantages

The use of the RDT does not eliminate the need for malaria microscopy. The RDT

may not be able to detect some infections with lower numbers of malaria parasites

circulating in the patient’s bloodstream. Also, there is insufficient data available to

determine the ability of this test to detect the 2 less common species of malaria, P.

ovale and P. malariae. Therefore, all negative RDTs must be followed by microscopy

to confirm the result.

In addition, all positive RDTs also should be followed by microscopy. The currently

approved RDT detects 2 different malaria antigens; one is specific for P. falciparum

and the other is found in all 4 human species of malaria. Thus, microscopy is needed

to determine the species of malaria that was detected by the RDT. In addition,

microscopy is needed to quantify the proportion of red blood cells that are infected,

which is an important prognostic indicator.

1.5.2.2. Advanced Laboratory Diagnostic Methods

Although advanced malaria diagnostic methods exist, they are not as suitable for

wide application in the field as microscopy or RDTs. They are unsuitable for use in

routine disease management in resource-limited settings and are often used for

research purposes. These are:

A. Quantitative Buffy Coat (QBC)

This technique is a qualitative method for rapidly detecting malaria parasites in

centrifuged capillary or venous blood. QBC utilizes density gradient layering of

stained blood cells, together with mechanical expansion of the hematocrit buffy

coat.

15

Figure 1. 2. Quantitative buffy coat of packed blood

The parasites are detected by fluorescent microscopy using acridine orange stain. It

is fast, easy and may be more sensitive than the traditional thick film examination.

Its main advantages are faster result delivery within 15-30 minutes, and a potential

for accidentally detection of filarial worms. However, it may provide false positive

results due to artifacts, species differentiation can be difficult, and per test cost is

expensive.

Figure 1. 3. Malaria parasite with AO stained and examined under Fluorescence

microscopy

Advantage

• Quick

• Can accidentally detect filarial worms

• Sensitive (at least as good as thick blood film)

Challenges od QBC

• May provide false positive results/Artifacts may be reported as positive

• Species differentiation can be difficult

• Expensive

16

B. Thin film acridine orange technique/ Microscopy using Kawamoto’s

fluorochrome technique

Fluorescence microscopy combined with fluorochrome staining of thin blood films

with acridine orange (AO) has been reported to be more sensitive than the

Romanowsky technique for the detection of malaria parasites and emits two

fluorescence colors, green (530 nm) and red (650 nm) when excited at 430 nm and

492~495 nm, respectively. Therefore, AO staining permits differential coloration of

green (nuclei) and red (cytoplasm) in stained parasites; the outlines of the parasites

stained by these dyes are well preserved and the general morphology is comparable

to specimens stained by Giemsa.

C. Immunological Tests (Anti-malarial Antibody Test)

Antibodies to the asexual blood stages appear a few days after malarial infection,

increase in titer over the next few weeks, and persist for months or years in semi-

immune patients in endemic areas, where re-infection is frequent. The antibody

tests can be done using either indirect immune fluorescence (IFA) tests or an

enzyme-linked immune sorbent assay (ELISA). Because of the time required for

development of antibodies and also the persistence of antibodies, serologic testing

is not practical for routine diagnosis of acute malaria but instead used to determine

past exposure.

Indirect Fluorescent Antibody Test

Malaria antibody detection is performed using the indirect fluorescent antibody

(IFA) test. The IFA procedure can be used to determine if a patient has been infected

with Plasmodium. Because of the time required for development of antibody and

also the persistence of antibodies, serologic testing is not practical for routine

diagnosis of acute malaria. However, antibody detection may be useful for:

• Screening blood donors involved in cases of transfusion-induced malaria

when the donor’s parasitemia may be below the detectable level of blood film

examination

• Testing a patient, usually from an endemic area, who has had repeated or

chronic malaria infections for a condition known as tropical splenomegaly

syndrome

• Testing a patient who has been recently treated for malaria but in whom the

diagnosis is questioned.

17

Species-specific testing is available for three of the four human species: P.

falciparum, P. vivax, and P. malariae. P. ovale antigens are not always readily

available and so antibody testing is not performed routinely. Cross reactions often

occur between Plasmodium species and Babesia species. Blood stage Plasmodium

species schizonts (merozoites) are used as antigen. The patient’s serum is exposed

to the organisms; homologous antibody, if present, attaches to the antigen, forming

an antigen-antibody (Ag-Ab) complex. Fluorescein-labeled anti-human antibody is

then added, which attaches to the patient’s malaria-specific antibodies. When the

slide is examined with a fluorescence microscope, if parasites fluoresce an apple

green color, a positive reaction has occurred.

Enzyme immunoassays have also been employed as a tool to screen blood donors,

but have limited sensitivity due to use of only Plasmodium falciparum antigen

instead of antigens of all four human species.

Figure 1. 4. Indirect immune fluorescent test for malaria parasite

D. Polymerase Chain Reaction (PCR)

This technique is used to detect parasite nucleic acids. Parasite nucleic acids are

detected using polymerase chain reaction (PCR). The principle is based on the

extraction of parasite DNA and amplification by polymerase chain reaction using

specific primers to yield a product that can easily be visualized in ethidium bromide

stained agarose gel. As little as one parasite per microliter of blood can be detected

by this method. It is highly specific and sensitive (10 times more sensitive than

microscopy) in detecting the plasmodium species, particularly in cases of low level

parasitemia and mixed infections, with a sensitivity of 1.35 to 0.38 parasites/μl for P.

falciparum and 0.12 parasites/μl for P. vivax. However, it requires expensive

laboratory equipment in specialized laboratory settings and often used in reference

laboratories to confirm malaria parasite species (if in doubt); to validate Rapid

Diagnostic Tests (RDTs) as part of planned quality assurance programmers’; and for

research purposes.

18

It is of limited utility for the diagnosis of acutely ill patients in the standard healthcare

setting. PCR results are often not available quickly enough to be of value in

establishing the diagnosis of malaria infection. It is most useful for confirming the

species of malaria parasite after the diagnosis has been established by either smear

microscopy or RDT.

E. Flowcytometry

Flowcytometry and automated hematology analyzers have been found to be useful

in indicating diagnosis of malaria during routine blood counts. In cases of malaria,

abnormal cell clusters and small particles with DNA fluorescence, probably free

malarial parasites, have been seen on automated hematology analyzers and it is

suggested that malaria can be suspected based on the scatter plots produced on the

analyzer. Automated detection of malaria pigment in white blood cells may also

suggest a possibility of malaria with a sensitivity of 95% and a specificity of 88%.

Reference

1. Centers for Disease Control and Prevention Division of Parasitic Diseases and

Malaria 1600 Clifton Road MS A-06Atlanta, GA 30329-4027 [email protected]

https://www.cdc.gov/malaria/about/biology/index.html

2. Global Technical Strategy for malaria 2016–2030 WHO

3. https://www.cdc.gov/malaria/references_resources/mmwr.html

4. Rougemont M, Van Saanen M, Sahli R, Hinrikson HP, Bille J, Jaton K. Detection

of Four Plasmodium Species in Blood from Humans by 18S rRNA Gene Subunit-

Based and Species-Specific Real-Time PCR Assays. J Clin Microbiol 2004,

42(12):5636.

5. Snounou G, Viriyakosol S, Zhu XP, Jarra W, Pinheiro L, do Rosario VE, et al. High

sensitivity detection of human malaria parasites by the use of nested

polymerase chain reaction. Mol Biochem Parastiol 1993; 61:315–320.

6. Hojgaard A, Lukacik G, Piesman J. Detection of Borrelia burgdorferi, Anaplasma

phagocytophilum and Babesia microti, with two different multiplex PCR assays.

Ticks and Tick-borne Diseases 2014 (5):349–351.

7. Bonnet S, Jouglin M, Malandrin L, Becker C, A. Agoulon A, L’Hostis M, Chauvin

A. Transstadial and transovarial persistence of Babesia divergens DNA in Ixodes

ricinus ticks fed on infected blood in a new skin-feeding technique. Parasitol

2007; 134:197–207

19

1.6. Global and Regional Burden of Malaria

1.6.1. Global Burden

Malaria is a serious public health problem in many parts of the world, exacting an

unacceptable toll on the health and economic welfare of the world’s poorest

communities. According to WHO 2018 report, in 2017, an estimated 219 million

cases of malaria occurred worldwide compared with 216 million cases in 2016

1.6.1.1. Regional and Global Trends in Malaria Cases

Most of the cases in 2017 were in the WHO African Region (92%), followed by the

WHO South-East Asia Region (5%) and the WHO Eastern Mediterranean Region

(2%).

The 10 highest burden countries in Africa reported increases in cases of malaria in

2017 compared with 2016. Of these, Nigeria, Madagascar and the Democratic

Republic of the Congo had the highest estimated increases, all greater than half a

million cases.

Rwanda has noted a reduction in its malaria burden, with 430 000 fewer cases in

2017 than in 2016, and Ethiopia and Pakistan marked decreases of over 240 000

cases over the same period.

The incidence rate of malaria declined globally between 2010 and 2017, from 72 to

59 cases per 1000 population at risk

Plasmodium falciparum is the most prevalent malaria parasite in the WHO African

Region, accounting for 99.7% of estimated malaria cases in 2017.

1.6.1.2. Malaria Deaths

According to WHO 2018 report, in 2017, there were an estimated 435 000 deaths

from malaria globally, compared with 451 000 estimated deaths in 2016, Children

aged under 5 years are the most vulnerable group affected by malaria. In 2017, they

accounted for 61% (266 000) of all malaria deaths worldwide

The WHO African Region accounted for 194 million cases and 407,000 of all malaria

deaths in 2017. Nearly 80% of global malaria deaths in 2017 were concentrated in

17 countries in the WHO African Region and India; seven of these countries

accounted for 53% of all global malaria deaths: Nigeria (19%), Democratic Republic

of the Congo (11%), Burkina Faso (6%), United Republic of Tanzania (5%), Sierra

Leone (4%), Niger (4%) and India (4%).

20

1.6.1.3. Malaria Elimination

Globally, the elimination net is widening, with more countries moving towards zero

indigenous cases: in 2017, 46 countries reported fewer than 10 000 such cases, up

from 44 countries in 2016 and 37 countries in 2010. The number of countries with

less than 100 indigenous cases – a strong indicator that elimination is within reach

– increased from 15 countries in 2010 to 24 countries in 2016 and 26 countries in

2017.

• Paraguay was certified by WHO as malaria free in 2018, while Algeria,

Argentina and Uzbekistan have made formal requests to WHO for

certification

• In 2016, WHO identified 21 countries with the potential to eliminate malaria

by the year 2020. WHO is working with the governments in these countries –

known as “E-2020 countries” – to support their elimination acceleration

goals.

• Although 11 E-2020 countries remain on track to achieve their elimination

goals, 10 have reported increases in indigenous malaria cases in 2017

compared with 2016.

1.6.2. Malaria Situation in Ethiopia

1.6.2.1. Seasonality, Weather, Geography and Climate

In Ethiopia, malaria is the most severe public health problem, and among the

leading causes of morbidity and mortality. Approximately 75% of the country land

mass is malarious, and 60% of the total population is at risk of malaria parasite

infection.

Altitude and climate (rainfall and temperature) are the most important determinants

of malaria transmission in the country. The transmission is seasonal and largely

unstable, where the peak malaria transmission occurs between September and

December in most parts of Ethiopia, after the main rainy season from June to

August. In addition, some areas experience a second minor malaria transmission

period from April to June, following a short rainy season from February to March.

January and July typically represent low malaria transmission seasons in most

communities. Since peak malaria transmission often coincides with the planting and

harvesting season. Although all age groups are at risk of malaria, the majority of

malaria burden is among older children and working adults in rural agricultural

areas, there is a resultant heavy economic burden in Ethiopia.

21

1.6.2.2 Vector Species and Abundance A member of the An. gambiae complex,

• Anopheles (A.) arabiensis, is the primary malaria vector in Ethiopia, with

• A. funestus,

• A. pharoensis, and

• A. nili as secondary vectors.

The sporozoites rate for A. arabiensis has been recorded to be as high as 5.4%. The

host-seeking behavior of A. arabiensis varies with the human blood index collected

from different areas ranging between 7.7% and 100%. A. funestus, a mosquito that

prefers to feed exclusively on humans, can be found along the swamps of the Baro

and rivers and shores of lakes in Tana in the North and the Rift Valley areas. A.

pharoensis is widely distributed in Ethiopia and has shown high levels of insecticide

resistance, but its role in malaria transmission is unclear. An. nili can be an

important vector for malaria, particularly in Gambella Regional State. Insecticide

resistance among these vectors has become an important issue, with implications

for vector control strategies.

1.6.2.3 Parasite Prevalence, Altitude Strata and Annual Parasite Incidence

(API):

According to HMIS 2017, the two most dominant parasitic species responsible to

cause malaria in Ethiopia are P. falciparum and P. vivax with proportion of about

70% and 30% respectively. Typical human and mosquito behavior results in most

malaria parasite transmission occurring indoors during nighttime hours within rural

households in lowlands and middle elevations, and only occasionally in the

highland fringe areas of Ethiopia greater than 2,000 meters above sea level (asl).

Malaria transmission may also sometimes occur outdoors during nighttime work or

social activities, or may be associated with temporary overnight travel to other

malarious areas. Recent published and unpublished reports indicate an increased

malaria incidence among migrant laborers in various parts of the country, most

importantly in the northwest development corridors of the country bordering Sudan

and South Sudan

Many Ethiopian communities have low and unstable malaria transmission patterns

that result in low host immunity and significant clinical malaria illness risk after

malaria infections, increased tendency for rapid progression to severe malaria, and

propensity for malaria epidemics affecting all age groups.

22

The epidemiology of malaria in Ethiopia, therefore, contrasts with that of many

other countries in Africa with high malaria transmission where malaria morbidity

and mortality mainly affect young children. Emerging data from episodic special

outbreak investigations and unpublished anecdotes from Ethiopian malaria partners

suggest that older boys and men may be at special risk for malaria from

occupational and travel-related factors such as engaging in seasonal migrant farm

work.

Malaria parasite prevalence (as measured by microscopy) in Ethiopia was 0.5%

(Malaria Indicator Survey (MIS) 2015), while slide positivity rate approximately 25%

(HMIS 2017), and annual incidence rate was 18% (HMIS 2017).

The 2011 MIS indicated that 1.3% of individuals were positive for malaria using

microscopy and 4.5% were positive for malaria using RDTs below 2,000 meters, with

only 0.1% prevalence above 2,000-meter elevation. Plasmodium falciparum

constituted 77% of infections detected below 2,000 meters. There was essentially no

P. falciparum detected by microscopy among persons surveyed within households

having measured elevations above 2,000 meters in the 2011 MIS.

In 2017, the FMOH updated the country’s malaria risk strata based upon malaria

annual parasite incidence (API), calculated from micro-plan data from more than 800

districts.

A malaria risk map from this API analysis, areas with malaria transmission risk by

API classified as:

1. High (≥100 cases/1,000 population/year),

2. Moderate (≥5 <100),

3. low (>0 -<5), and

4. Malaria-free (~0). Areas with the highest malaria transmission risk as

stratified by district API appear to be largely in the lowlands and midlands of

the western border with South Sudan and Sudan. Many densely populated

highland areas were newly classified as malaria-free (API=0), including the

capital city of Addis Ababa. Based on the current stratification,75% of the land

mass is malarious and the proportion of the population at risk of malaria is

about 60% with 54 (6.4%) woredas having high transmission.

23

Figure 1. 5 Malaria risk map of districts by annual parasite incidence, Ethiopia,

2017

N.B. Both global and national malaria information on epidemiology could be

changed from time to time as per WHO annual report. So, readers are advised

to refer updated information from current WHO reports.

1.6.3. National Malaria Control Strategy

The National Malaria Strategic plan (NMSP) for the years 2014-2020 was finalized in

August 2014, which was envisioned to be aligned with the next five-year health

sector transformation plan (HSTP) 2015/16–2019/20 and submitted along with the

concept note for the Global Fund New Funding Model (NFM) application.

1.6.3.1. Updates in the National Malaria Control Strategy of the Year 2017–2020

The main components of the Strategy

• Early diagnosis and effective treatment

• Access to laboratory Diagnosis and treatment of febrile patients within 24 hrs.

• Selective vector control measures: ITNs and IRS

• Environmental control – eliminating insect breeding sites

• Epidemic prevention and control

• Early detection and containment of epidemics

Support strategies:

Human Resources (HR) development

o Information education communication (IEC) /Behavioral change

communication (BCC)

o Monitoring and Evaluation (M& E)

• operational research

The proposed goals and objectives for the2017-2020 NMSP includes: al

24

• By 2020, to achieve near zero malaria deaths in Ethiopia (near zero malaria

death is defined as no more than 1 confirmed malaria death per 100,000

populations at risk)/year.

• By 2020, to reduce malaria cases by 40% from baseline of 2016.

• By 2030, to eliminate malaria from Ethiopia.

Strategic Objectives

1. By 2020, all households living in malaria endemic areas will have the

knowledge, attitudes and practice to adopt appropriate health-seeking

behavior for malaria prevention and control.

2. By 2017 and beyond, 100% of suspected malaria cases are diagnosed using

RDTs or

microscopy within 24 hours of fever onset.

3. By 2017 and beyond, 100% of confirmed malaria cases are treated according

to the national guidelines.

4. By 2017 and beyond, ensure that the population at risk of malaria has

universal access to one type of globally recommended vector control

intervention.

5. By 2020, malaria elimination program will be implemented in 239 districts.

6. By 2020, 100% complete data and evidence will be generated at all levels

within the nationally designated time periods to facilitate appropriate

decision-making

The National strategic plan provides a detailed account on the status and direction

of the major malaria prevention and control strategies which includes:

N.B: Note that any data in this chapter may be updated annually as necessary.

A. Community Empowerment and Mobilization

Community empowerment and mobilization are central to malaria prevention and

control. Ethiopia’s Health Extension Program educates, mobilizes and involves the

community in all aspects and stages of malaria control and leads to increased

ownership of the program.

B. Diagnosis and Case Management

Since 2005, there has been a major shift from clinical diagnosis to confirmatory

diagnosis following the wide-scale use of RDTs in peripheral health facilities. To

improve the quality of malaria diagnosis and treatment at peripheral health facilities

(health posts) pan specific RDTs are now being introduced. HEWs will be trained on

the use of multi-species RDTs in the integrated refresher training (IRT).

25

C. Prevention

The main major vector control activities implemented in the country include IRS,

LLINs and environmental control.

D. Active Surveillance and Epidemic Control

Aims to achieve a high quality, broadly based malaria infection detection,

investigation and response ‘Surveillance System’ to further reduce malaria

transmission and improve the detection and timely response to malaria epidemics.

Malaria detection, investigation, response and elimination activities will achieve a

high quality, broadly based malaria infection detection, investigation and response

surveillance System to further reduce malaria transmission, prevent and stop

epidemics and eliminate malaria especially in targeted areas that are prone to

outbreaks. There will be a transition from epidemic detection and response to

surveillance and infection response as transmission declines to near zero.

E. Health system strengthening and capacity building

The health system strengthening and capacity building includes monitoring and

Evaluation activities and development of Human Resources.

1.6.3.2. Updates in the Strategy Section

The Ministry has officially declared malaria elimination efforts for selected 239 low

malaria burden districts. Ongoing discussions are occurring with the FMOH to

coordinate pre-elimination activities together with other donor-supported projects

that continue to help shrink the malaria transmission map in Ethiopia.

Current control interventions will be strengthened, with emphasis given to: -

• Improving access to ITNs,

• case management and

• case detection services,

• surveillance,

• stock management and

• capacity building activities at the district level to further reduce cases

1.6.4. Levels of Health Facilities and Types of Diagnostic Tests in Ethiopia

1.6.4.1. National and Regional Reference Laboratories

The national and regional reference laboratories are performing specialized

laboratory diagnostic tests mainly for operational researches and trainings. Malaria

26

parasite molecular, serological tests, drug level determinations and RDT evaluations

are conducted at the national reference laboratory. Malaria microscopy is mainly

used at the national level for research, large surveys, quality control and training

purposes. At the regional reference laboratories, malaria microscopy is mainly

conducted for the purposes of training and external quality assessment schemes.

In accordance with the National Malaria guidelines of 2012, malaria microscopy is

the sole technique employed in hospital and health center levels. Therefore, it is

critical that these facilities are equipped with standard microscopes, have adequate

supplies and skilled microscopists.

1.6.4.2. Health Posts

The basis of suspicion for malaria infection is fever (rise in body temperature) from

the patient’s history and verified by touching or recording the temperature with a

thermometer. Rapid diagnostic tests (RDTs) shall be used by the health extension

workers

References

1. World Malaria Report 2018

2. World Malaria Report 2017

3. President’s Malaria Initiative, Ethiopia Malaria Operational Plan FY

2018

4. National Malaria Indicator Survey 2017

5. National Malaria Program Monitoring and Evaluation Plan 2014 – 2020

6. The FMOH’S NMSP (2017-2020)

7. Global Technical Strategy for Malaria 2016–2030

8. WHO Malaria Terminology

27

CHAPTER TWO: Microscope: Types, Parts, Care

and Handling

Chapter Description:

This chapter provides the learners with the knowledge, skills and right attitudes to

set up a light microscope for optimum resolution, to prepare routine samples and

to observe, identify and report sample characteristics.

2.1. Microscope

Microscope is an instrument used to see objects that are too small to be seen by the

naked eye. Microscopy is the science of investigating small objects using

microscope. Microscopic means invisible to the eye unless aided by a microscope.

Microscope has an essential role for the diagnosis and management of many

infectious diseases such as malaria, tuberculosis, intestinal parasites, etc. through

examination of clinical specimen.

2.2. Types of Microscope

There are three types of microscopes (See the detail in Annex 1):

1. Simple Microscope.

2. Compound Microscope.

3. Electron Microscope

2.2.1. Simple Microscope

Simple microscope is generally considered to be the first microscope. It is an

ordinary magnifying glass which may have a magnification of 5x, 10x, 20x or more.

It was created in the 17th century by Antony van Leeuwenhoek, who combined a

convex lens with a holder for specimens. Magnifying between 200 and 300 times, it

was essentially a magnifying glass. While this microscope was simple, it was still

powerful enough to provide van Leeuwenhoek information about biological

specimens, including the difference in shapes between red blood cells. Today,

simple microscopes are not used often because the introduction of a second lens

led to the more powerful compound microscope.

28

Figure 2. 1. simple microscope

2.2.2. Compound Microscope

A compound microscope has a much higher magnification than the simple

microscope. The typical compound light microscope is capable of increasing our

ability to see details 1000 times enlarged, so that objects as small as 0.1 micrometer

(µm) or 100 nanometers (nm) can be seen. This microscope uses at least two lenses

positioned at different places. A magnified image of the object is first produced by

one lens and this image is further enlarged by a second lens to give a more highly

magnified object. These two lenses are placed one at the end of each tube. The first

lens which is near to the object is known as the objective lens. While the second lens

which is near the eye is known as the eyepiece lens.

29

Figure 2. 2. compound microscope

2.2.2.1. Types of Compound Microscope

Based on the available number of eyepieces, we can have at least two types of

compound microscopes:

a. Monocular microscopes

• Have a single eyepiece

• Are convenient for use by beginners, for field work where there is no

electricity and for photographing clinical specimens.

b. Binocular microscopes

• Have two eyepieces

• Are recommended where much microscopic work has to be done, i.e. in

routine examinations.

The total magnification power of a microscope is the magnification of its objective

multiplied by that of its eyepiece. For example, using a 10x objective and 10x

eyepiece, the total magnification of microscope is 100x.

The Resolving Power of a Microscope.

• The resolving power of a microscope is described as the ability of the

microscope to: A. separate clearly two objects that are very close together.

The ability of an objective to distinguish the dots separately and distinctly.

30

• The limit of usable magnification. The actual power or magnification of a

compound optical microscope is the product of the powers of the ocular

(eyepiece) and the objective lens. The maximum normal magnifications of

the ocular and objective are 10× and 100× respectively, giving a final

magnification of 1,000×.

o The human eye can separate dots that are 0.25 mm in diameter.

o A light microscope can separate dots that are 0.25µm apart.

o The electron microscope can separate dots that are 0.5 nm apart.

Based on the type of illumination system, different types of compound microscopes

are available:

1. Light microscope

2. Fluorescent microscope

3. Dark field microscope

4. Phase contrast microscope

I. Light microscope.

A light microscope (LM) is an instrument that uses visible light and magnifying

lenses to examine small objects not visible to the naked eye or in finer detail than

the naked eye allows. Light from a mirror is reflected up through the specimen, or

object to be viewed, into the powerful objective lens, which produces the first

magnification. The image produced by the objective lens is then magnified again by

the eyepiece lens, which acts as a simple magnifying glass.

II. Fluorescent microscope

A fluorescence microscope is an optical microscope that uses fluorescence and

phosphorescence instead of, or in addition to, reflection and absorption to study

properties of organic or inorganic substances. The conventional microscope uses

visible light (400-700 nanometers) to illuminate and produce a magnified image of

a sample. A fluorescence microscope, on the other hand, uses a much higher

intensity light source which excites a fluorescent species in a sample of interest

III. Dark field microscope.

Dark Field illumination is a technique used to observe unstained samples causing

them to appear brightly lit against a dark, almost purely black, background. When

light hits an object, rays are scattered in all azimuths or directions.

31

IV. Phase contrast microscope.

Phase-contrast microscopy is an optical microscopy technique that converts phase

shifts in light passing through a transparent specimen to brightness changes in the

image. Phase shifts themselves are invisible, but become visible when shown as

brightness variations.

2.2.2.2. Parts of Compound Microscope and their Use

Microscope stand - the stand of a basic microscope includes

Tube - Holds the eyepiece and objectives in line and at the correct distance

Stage - Is a flat surface where the specimen to be examined is placed. - In the center

of the stage there is circular hole that allows the light from the mirror or lamp to

pass through

Mechanical stage - This enables the slide on which the specimen is mounted to be

moved in a controlled way, vertically or horizontally.

Sub stage - Immediately below the stage is the sub stage which holds a condenser

lens with an iris diagraph and a holder for light filters and stops.

Foot/Base - This ensures microscope stability on the laboratory bench.

The Mechanical Adjustment System

Coarse adjustment

• Usually used to focus using low-power objectives

• Controlled by a pair of large knobs positioned one on each end of the body

• Rotation of these knobs moves the tube with its lenses or, in some

microscopes, the stage up or down fairly rapidly.

Fine adjustment

• Use to focus objectives for high-power objectives because they require a fine

adjustment

• Moves the objectives or stage up or down very slowly.

• Controlled/moved by two smaller knobs on each side of the microscope.

Condenser adjustment

• The condenser has an adjustment system for its focusing light onto the

specimen on the stage. This is done by opening and closing of its aperture.

• It can also be swung aside to remove it or to exchange it with another.

• The condenser is usually focused by rotating a knob to one side of it.

32

Optics of a Light Microscope

Objectives

• Objectives are the most important parts of a microscope because the quality

and most of the magnification of the image depend on them.

• Modern objectives are described according to their magnification and older

objectives are often described according to their equivalent focal length (EFL)

The focal length of the lens is the distance between the lens and the image sensor

when the subject is in focus, usually stated in millimeters (e.g., 28 mm, 50 mm, or

100 mm). In the case of zoom lenses, both the minimum and maximum focal lengths

are stated, for example 18–55 mm.

Description in

Objective Diameter Equivalent focal length (EFL)

10x 16mm Or 2/3inch

40x 4mm Or 1/6 inch

100x 2mm Or 1/12 inch

- For most routine medical laboratory work, 10x, 40x and 100x objectives are

required.

Source of description:

http://www.microscope-microscope.org/basic/microscope-parts.htm

The low power objective: 10x

• Used for initial scanning and observation in most microscopic works.

• Used for initial focusing and light adjustment of the microscope.

The high power objective: 40x

• Used for more detailed study as the total magnification with 10x eyepiece is

400.

• Used for the diagnosis of intestinal protozoa parasites, urine sediments/cells,

casts crystals, and histological sections

The oil immersion objectives: 100x

• This lens has a very short focal length and working distance.

• The objective lens rests almost on a microscopic slide when in use.

• Known as oil immersion objective since a special grade oil must be placed

between the objective and the slide.

33

• Oil is used (with refractive index of 1.515) to increase the numerical aperture

and the resolving power of the objective.

Ocular (Eyepiece)

• A lens that magnifies the image formed by the objectives.

• The usual magnification of the ocular is 10x, others are 4x, 6x, 7x, 15x and

sometimes as high as 20x.

• The higher the power, the greater the total magnification of the microscope.

The lower the power of the eyepiece, however, the brighter and sharper is

the image.

Condenser

• A large lens with an iris diaphragm placed below the stage.

• It directs and focuses the beam of light from the light source, lamp or mirror,

to the specimen under examination. - Usually consists of two or sometimes

three lenses

• The lenses are curved so that the light can pass to the objectives at a

sufficiently wide angle.

• The condenser position is adjustable; it can be raised and lowered beneath

the stage and the light must be correctly focused on the material to be

examined.

Iris diaphragm

• It controls the amount of light passing through the specimen under

examination.

• Located at the bottom of the condenser, under the lenses, but within the

condenser body.

• It can be opened or closed as necessary to adjust the light intensity.

Köhler Adjustment

• August Köhler invented the procedure for optimum illumination of an object

in a light microscope. Köhler illumination is also known as double diaphragm

illumination because it employs both a field and an aperture iris diaphragm

to set up the illumination. If the light path is set up properly, you will have the

advantages of an evenly illuminated field, a bright image without glare and

minimum heating of the specimen. Refer to the appendix for instructions on

how to adjust the Köhler illumination.

34

Note: In certain microscopes, the field diaphragm is usually not

present and the Köhler adjustment does not apply.

Mirror

• Used in the microscope without built in illumination

• It reflects the beam of light from the light source upwards through the iris in

to the condenser.

The illumination system

• The modern compound microscope most often has a built-in illumination system

with a controller to adjust the amount of light comfortable for the microscopists.

Light source (Power switch)

The light source in your microscope is a lamp that you turn on and off using a

switch. It is the main power switch that turns the illumination on or off.

2.2.2.3. Routine Use of Basic Microscope: Steps

1. Place the microscope on a firm bench and not exposed to direct sunlight.

2. Switch on the light.

3. Place the specimen to be examined on the stage.

4. Select the objective to be used.

• Its recommended to begin examination with10x objectives. Once in focus, all

the other objectives also will be in focus.

5. Focus the objectives

• Move the objectives carefully downwards using the coarse adjustment knob

and locking at it from the side until the lens is near the specimen but not

touching it.

• Move the objectives slowly upwards, until the image comes into view and is

sharply focused.

6. Focus the condenser

• Open the iris of the condenser fully and, focus the condenser on the detail of

the light source until the image appears sharp.

7. Adjust the opening of the condenser iris according to the specimen examined

• Stained smears the condenser iris should be opened more widely giving a

well-illuminated image with fine details.

35

• Unstained specimen the condenser iris should be opened in reduced manner

to increase the contrast.

8. Examine the specimen the specimen using the mechanical stage to move it

9. For a higher magnification, swing the 40x into place. Focus the 40x objectives

using the fine adjustment.

10. For the highest magnification, add a drop of immersion oil to the specimen and

swing the 100x oil immersion objectives in to place. Open the iris fully to fill the

objectives with light.

Note: If examining a stained smear directly with the oil immersion lens

and it is not possible to focus it, remove the slide and check that the oil

has been placed on the smear side of the slide.

2.2.3. Electron Microscopy

Electron microscopes use a beam of electrons rather than visible light to illuminate

the sample. They focus the electron beam using electromagnetic coils instead of

glass lenses (as a light microscope does) because electrons can’t pass through

glass.

A. The transmission electron microscope (TEM) was the first electron

microscope to be developed. It works by shooting a beam of electrons at a

thin slice of a sample and detecting those electrons that make it through to

the other side. The TEM lets us look in very high resolution at a thin section

of a sample (and is therefore analogous to the compound light microscope).

This makes it particularly good for learning about how components inside a

cell, such as organelles, are structured.

B. The scanning electron microscope (SEM) lets us see the surface of three-

dimensional objects in high resolution. It works by scanning the surface of an

object with a focused beam of electrons and detecting electrons that are

reflected from and knocked off the sample surface. At low magnifications,

entire objects (such as insects) viewed on the SEM can be in focus at the same

time. That’s why the SEM is so good at generating three-dimensional images

of lice, flies, snowflakes and so on.

2.3. Care and Handling of Microscope

Good working knowledge and proper care of the microscope are critical to good

diagnostic work. There are only a few absolute rules to observe in caring for the

36

microscopes you will use. Taken care of, these instruments will last many decades

and continue to work well. Please report any malfunctions immediately.

• Always use two hands to carry the microscope - one on the arm and one

under the base. Never carry the microscope upside down, for the ocular can

and will fall out.

• Never expose it to sharp knocks, vibrations, moisture, dust or direct sunlight.

• Use lens paper to clean all lenses before and after using the oil immersion

lens. Other papers are too impure and will scratch the optical coating on the

lenses. Also, do not use any liquids when cleaning the lenses – use lens paper

only!

• Always use the proper focusing technique to avoid ramming the objective

lens into a slide - this can break the objective lens and/or ruin an expensive

slide.

• Always turn off the light when not in use.

• Always carefully place the wire out of harm’s way. Wires looped in the leg

spaces invite a major microscope disaster. Try sliding the wire down through

the drawer handles beside your bench space.

• Always replace the cover on the microscope when you put it away

2.4. Microscope Maintenance and Storage Conditions

(see the detail in annex 2)

Routine optical and mechanical maintenance of compound microscopes can ensure

that your microscope works well for years. Periodic microscope servicing by a

qualified microscope technician is recommended. Compound microscopes should

generally be serviced after about 200 hours of use. For most schools, this would be

about every three years; possibly more frequent if the microscope is used multiple

times each day.

Never attempt to disassemble any part of the microscope for repair. If there is any

problem with the microscope, contact the microscope company’s technical support

unit or their local agents, or consult with a qualified technician, around.

Humidity causes fungal growth on the surface of lenses and prisms. This can cause

cloudiness of the view field and rusting of metal parts of the microscope. To protect

the microscope from fungus, always keep the glass surface as clean as possible and

37

free of dirt and fingerprints. Reduce the growth of fungus by continuously using an

air conditioner to lower humidity. The use of air-conditioning in the daytime only

will lead to condensation on the microscope once it is turned off, again favoring

growth of fungus. Alternatively, drying the microscope within a temperature-

controlled cabinet, silica gel (desiccant), or anti-mold strips may be useful.

Cabinet Box (for humidity and temperature control) (see Figure 4.4) Store a

microscope in a cabinet box with air inlets and outlets for air circulation and a 20-

watt bulb for keeping a dry, stable environment

Figure 2. 3 Cabinet Boxes

Silica Gel Place dry blue silica gel (about 250 g) in a shallow plate and place it in the

bottom of the sealed microscope box. Silica gel is blue when it is dry, but turns

pinkish when it becomes wet. As soon as the silica gel becomes pink, replace it.

Alternatively, heat the gel in crucibles until I the absorbed water evaporates & turns

blue again before using it.

SOP:

Changing a Microscope Bulb or Fuse Replacing the Microscope Bulb

• Before replacing the lamp bulb or fuse, be sure to turn off the power switch

and disconnect the power cord from the socket.

• Carefully lie the microscope on its side.

• Carefully unscrew the base plate underneath the microscope stand and open

it.

• Remove the bulb by pulling gently. Check to see if the wire in the bulb is

broken or 'burnt out'.

• Replace with new bulb of the same wattage.

• Replace the base plate.

38

• When replacing a fuse, keep the microscope in an upright position.

• Look at the rear of the base stand, where you will see a small plastic cover

with 'fuse' written on it.

• Unscrew the cover carefully, remove it and check whether the fuse is broken

with a magnifying device. Replace this if it is broken.

• Replace the bulb with the same type of fuse. Screw it carefully into place.

• Plug the microscope into the socket and turn the microscope on to check the

light.

• Find the location of the bulb

• Follow manufacturer’s instructions to remove the bulb

• Use tissue paper or an appropriate device to remove the bulb from the

microscope

• Check the model number on the bulb to ensure the use of a correct

replacement bulb

• Replace the bulb by holding it with lens paper or an appropriate device

NB: Never touch the bulb with your fingers.

Microscope Repair

• Never disassemble the microscope

• Optics: eyepieces and objectives

• Mechanics: stage and focus adjustments

• Repair of these items requires a service engineer

Cleaning a Microscope

Anti-mold strips

Anti-mold strips can be also applied to prevent mold. Replace these strips every 3

years. Always keep the four optical parts of the microscope clean (see figure 11.1of

the original note). Remove dust attached to the microscope with a blower or other

towels/tissue paper.

Use only immersion oil with the proper clearness, viscosity, and refractive index for

the immersion lens. Cedar oil and other types of oil such as baby oil, cooking oil and

liquid paraffin are not acceptable for this purpose as they will damage the lens.

Before putting the microscope away, wipe off the immersion oil by rubbing the

surface of the immersion (100x objective) lens gently with a washed soft gauze or

39

lens paper which is lightly moistened with ethyl ether/alcohol (80/20 vol/vol). This

can also be used to remove fingerprints or grease. Remove dust by softly brushing

the surfaces. For cleaning lenses and filters, wipe the object from the center, winding

a spiral to the periphery.

Microscope Cleaning Process

• Cleaning the eye piece

• Cleaning the objectives

• Cleaning the microscope stage

• Cleaning the microscope body

• Cleaning the condenser

• Cleaning Microscope Eyepiece Lenses

Steps in Cleaning Eye Piece

• Blow to remove dust before wiping lens

• Clean the eyepieces with a cotton swab moistened with lens cleaning solution

• Clean in a circular motion inside out

• Wipe the eyepieces dry with lens paper

• Repeat cleaning and drying if required

Cleaning Objectives

• Objectives are cleaned while attached to the microscope

• Moisten the lens paper with the cleaning solution

• Wipe gently the objective in a circular motion from the inside out

• Wipe with a dry tissue or lens cleaning paper

• Objectives should never be removed from the nosepiece

Cleaning the Microscope Stage

• Wipe the microscope stage using the cleaning solution on a soft cloth

• Thoroughly dry the stage

• Repeat the above steps, if required

Cleaning the Microscope Body

• Unplug the microscope from the power source

• Moisten the cotton pad with a mild cleaning agent (please give examples)

40

• Wipe the microscope body to remove dust, dirt and oil

• Repeat steps1–3, if required

Cleaning the Condenser

• Unplug the microscope from the power source

• Clean the condenser lens and auxiliary lens using lint-free cotton swabs

moistened with lens cleaning solution

• Wipe with dry swabs

Troubleshooting

• There are several conditions that can affect the proper functioning of the

microscope. Review these problems and their solutions.

1. The brightness of the viewing field is poor

Problem Solution

The condenser is too low Raise the condenser to correct its position

The condenser iris diaphragm is

closed

Open the diaphragm properly

2. Dark shadows in the field which move as you turn around the eye piece

Problem Solution

The surface of the eye piece has

scratches

Replace the eye piece

The eye piece is dirty Clean the eye piece

3. the image with the high power objective is not clear

Problem Solution

The slide is upside down Turn the slide over

There is an air bubble in the oil Move 100x lense quickly from side to side

The dirt in the objective Clean the lense

The oil is too sticky Use thinner immersion oil/specified immersion oil

4. The image is not clear with low power objective

Problem Solution

There is oil on the lense Clean the lense

There is a layer of dust in the upper

surface of the objective

Clean the lense

41

If the view field is still dim and cloudy, consider the following possible causes:

• Massive growth of fungus on the lenses or prisms due to storage in a high

humidity environment

• Penetration of immersion oil between the lenses of the objective through

damaged lens cement (due to use of poor-quality oil such as cedar oil or

misuse of xylene). This is likely the cause if a completely hazy field becomes

clear after changing the objective.

• A damaged objective (due to careless focusing, dropping, rough changing of

slides)

Frequently-encountered operational errors include the following:

• Focusing the first slide using the 100x immersion objective without passing

through a low power objective.

• Changing slides from under the immersion objective without turning it away

first.

• Wiping lenses without first blowing away dust and sand.

• Cleaning lenses or other parts with xylene.

• Using cedar wood oil, liquid paraffin, or xylene-diluted oil instead of pure

synthetic immersion oil.

• Keeping the microscope in a confined space without ventilation in a humid

climate.

2.5. Log Book

A microscope log book should be maintained to enter problems encountered in the

operation of the microscope, maintenance schedule, repairs done on the

microscope and availability of spares like bulbs, fuses, anti-mold strips etc.

42

CHAPTER THREE: Laboratory Safety

Chapter Descriptions:

This chapter is intended to provide general guidelines for laboratory safety as the

basis for maintaining a safe healthy working environment for laboratory users and

responsible for adhering to all safety guidelines and regulations for demonstrating

competency in implementing laboratory safety techniques.

3.1 Introduction

Safety is one of the 12 quality system essentials which is important in order to

protect the lives of employees and patients, to protect laboratory equipment and

facilities, and to protect the environment.

Why Safety is Important?

▪ Coming in contact with human blood or blood products is potentially

hazardous

▪ Good Safety practice protectant us Safety involves taking precautions to

protect you and the client against infection.

What Else Needs protection?

▪ Good safety practice also protects Other people who may come in contact

with testing by-products

▪ It Protect integrity of test products Protect environment from hazardous

materials

43

Laboratory Safety Policy

The management of the health facility is responsible for providing a safe healthy

working environment and to proactively maintain a well-documented and safe

workplace. Employees are responsible for adhering to all safety guidelines and

regulations for demonstrating competency in implementing laboratory safety

techniques. Each blood sample drawn or handled in a health facility carries the risk

of occupational exposure to HIV and other blood-borne infectious agents and other

biological samples are also potentially infectious. Not adhering to laboratory Safety

cause laboratory accident which result in loss of reputation, loss of customers / loss

of income, negative effet on staff rétention, increased costs and litigation, insurance.

3.2 General Safety Guidelines

Standard operating procedures (SOPs) that cover all steps should be clearly written

and carried out which also ensures safety measures. Generally, the following safety

precautions should be implemented at all times.

• Wear a laboratory coat when in the working room and remove any protective

clothing before leaving the laboratory.

• Wear gloves when taking and handling blood specimens.

• Do not touch your eyes, nose or other exposed membranes or the skin with

gloved hands.

• Change gloves between patients and remove the gloves before touching

objects and surfaces e.g. door handles and other objects not usually touched

by gloved hands; wash your hands and put on new gloves.

• Cover broken skin with water resistant wound covers before wearing gloves

• Wash your hands with soap and water immediately after any contamination

and after the work is completed. If gloves are worn, wash your hands with soap

and water after removing gloves.

• Puncture wounds, cuts and skin contaminated by spills or splashes of blood

should be thoroughly washed with soap and water. Bleeding from the wound

should be encouraged.

• Dispose of used lancets in a sharps container.

• Disinfect work surface areas when blood collection procedures are completed

and at the end of each working day.

• Do not eat, drink or smoke in the working area.

44

• For use on all surfaces, use 0.5% solution of bleach.

• Prepare fresh working solutions of bleach daily.

• Carefully handle all chemicals and reagents according to accepted standards

(refer the table below)

Table 3. 1 Safety Precautions for Chemicals Used in Malaria Microscopy.

Chemical Main hazard Safety precautions Giemsa

stock

Stain

• Highly flammable with flash

point12ºC

• Keep away from sources of ignition

• Avoid inhaling fumes and contact

with skin

Giemsa

Powder • Harmful if inhaled or

swallowed

• May cause irritation to

respiratory tract

• Contact with strong oxidizers

may cause fire or explosion.

• Fire or excessive heat may

produce hazardous

decomposition products.

• Keep container tightly closed in a

cool, well ventilated place.

45

Methanol • Highly flammable with

flashpoint 12ºC

• Volatile and hygroscopic

• Toxic if ingested or inhaled

• Can cause dermatitis and

damage to the optic nerve

and central nervous

system

• Keep away from sources of

ignition, sodium hypochlorite,

nitric acid chloroform, hydrogen

peroxide

• Avoid breathing vapor, protect skin

and eyes

• Use in a well-ventilated area

or preferable in a fume hood

Xylene • Harmful if inhaled, may cause

dermatitis if in contact with

skin

• Flammable with flashpoint

12ºC

• Protect from skin contact and use

in a well-ventilated area

• Do not keep in plastic containers

unless they are made of

polypropylene

• Do not use caps with rubber liners

3.3 Safety and Exposure Control Measures

Application of safety procedures in the laboratory is crucial to minimize accidental

exposure to infectious agents achieved by

• Applying universal safety precautions: Treat all biological samples as

infectious

• Wearing PPE

• Training Procedures to limit risk of infection should be instituted during blood

collection, sample handling, testing and disposal. Even though there are a

variety of different microorganisms that may put laboratory personnel at risk

while doing their job, the most important microorganisms to consider are

hepatitis B and C, and HIV.

There are 3 main routes of pathogen entry into the body:

• Non-intact skin: Naturally intact skin provides a good barrier; this barrier is

lost when skin is not intact.

• Mucous membrane exposure in eyes, nose and mouth

• Percutaneous injury (through the skin): Needle sticks, cuts and punctures

The transmission of HIV through needle stick injury ranges from 0.01-0.06%. If in

case needle-stick injury occurs, the following procedures help to avoid immediate

infection:

Personal hygiene and preventive measures: wash, wash, wash

Mucous membranes: flush thoroughly with plenty of clean water

Skin: apply soap and water for 5 minutes

Percutaneous: apply soap and water for 5 minutes

46

• Notify supervisor immediately

• Get completed exposure Report and

• Consult with local senior management in the health facility regarding

possible treatment and follow-up

The major elements of safety measures are:

A. Hand washing and First Aid

B. Exposure Control Plan

C. Hazard Communication

D. Exposure Determination

E. Methods of Compliance

F. Review of Safety Procedures and Training

A. Hand Washing and First Aid

Needle stick injury is the most common injury faced by laboratory personnel during

blood draws. Use the following preventive measures:

• Wash the punctured hand with running water and soap

• Encourage bleeding but don’t apply excess pressure

• Notify and consult senior staff at the facility regarding possible treatment and

follow-up.

Hand washing is the number one preventive measure against the spread of

infection. Therefore, wash hands before and after handling patients and after

handling all materials known or suspected to be contaminated.

B. Exposure Control Plan (ECP)

As part of the Laboratory Safety Manual, an Exposure Control Plan (ECP) addresses

blood-borne pathogen exposure and should include procedures for:

• Hazard communication

• Exposure determination

• Methods of compliance

• Exposure evaluation

• Post-evaluation for exposure occurrences

• Annual review of procedures and training

• Conduct regular safety audit

Along with a laboratory safety manual, a laboratory manager should have policies

and procedures for an exposure control plan.

47

C. Hazard Communication

The laboratory manager should provide graphics, warning signs and labels for

general hazard safety and bio-safety issues, Personal Protective Equipment (PPE)

and practices which are not allowed in the laboratory. General hazard safety

information such as toxic or carcinogenic reagents, poisons, flammable,

combustible or radioactive reagents, and volatile solvents should be provided. In

addition, a file of material safety data sheets (MSDS) specific for all chemicals used

in the laboratory should be available for reference.

Figure 3. 1. Hazard safety signs

D. Exposure Determination

The laboratory manager or Safety Officer should identify the laboratory working

areas which are at risk of exposure, and should those in the ECP.

There are various levels of exposure

• Those working directly with blood borne pathogens are at increased risk.

• Those working in the laboratory but not necessarily with the blood borne

pathogens are at secondary risk.

The laboratory manager needs to determine and document the risk level of

potential occupational exposure for all laboratory personnel.

E. Methods of Compliance

Employers need to implement administrative, engineering and PPE controls

as a means to protect against employee exposure to bio-hazardous blood

borne pathogens.

• Administrative Control

o Use SOP’s to limit employee exposure to blood borne pathogens

o Usage of appropriate safety signs

48

• Engineering Controls

o Employ procedures for use of safety devices used in the laboratory

(safety cabinets, safety needle devices, sharps containers, etc.)

• Personal Protective Equipment (PPE)

o Train laboratory personnel on the use of PPEs.

F. Review of Procedures and Training

All procedures and policies should be reviewed annually. Procedures for training

laboratory personnel on potential hazards and exposure precautions should be

established. Post evaluation procedures for treatment, counseling and follow-up, if

exposure occurs, should be established. Training of laboratory personnel should be

done at commencement of employment and on a yearly basis thereafter. All training

should be documented for each employee.

3.4 Testing Infrastructure and Equipment Management

Applying safety practices in the laboratory requires both infrastructure and trained

human resources. The following are infrastructure requirements:

• Waste disposal facilities

• Adequate light, water, sewage, ventilation and electrical facilities

• Appropriate laboratory design (superstructure, furniture and space)

• Appropriate storage facilities

• Restricted access to the laboratory.

Specimen collection area Hand washing area

Figure 3. 2 Some important Infrastructures for safe working area

Safety devices and facilities are important to operate malaria diagnosis in

compliance with general safety standards and universal precautions. Some of

the safety facilities are personal protective equipment’s (PPE), sharp containers,

49

hand washing and eye wash stations, emergency showers, incinerators and

others.

A. Personal Protective Equipment (PPE)

During phlebotomy, exposure to contaminated sharps presents the major risk.

During biological sample preparation in the laboratory, exposure to the skin or

mucous membranes presents the major risk. Perform the following during sample

preparation:

• Wear gloves and a laboratory coat

o Do not wear gloves or a laboratory coat in common areas or at home

o Change gloves regularly and never re-use gloves (change between

patients)

o Change gloves whenever it is contaminated

• Don’t wear open shoes or slipper

• Draw blood only in dedicated areas

• Avoid crowded areas or pathways

• Cover broken skin with water resistant wound covers even when wearing

gloves

Figure 3. 3. General safety equipment

B. Sharp Container

Sharps injury is the riskiest route of exposure. Sharps include: needles, blood

lancets, pipettes, broken test tubes,

• Use biohazard-labeled sharps containers that are leak proof.

• Use single-use disposable lancets, needles and scalpels.

• Use phlebotomy equipment with built-in safety features.

50

C. Eye wash

• Use eye washes for splashes to the eye

o Flush for 5 minutes for pathogens

o Flush at least 15 minutes for most chemicals.

3.5. Waste Disposal

Wastes should be segregated according to their types. Usually solid and liquid

wastes are collected and separately. Additionally, segregation can be made for

infectious and noninfectious wastes. The solid wastes should further be segregated

into sharp and non-sharp wastes. Liquid wastes can be classified into chemical and

biological types.

In general, to protect the surrounding population, the laboratory must dispose of

wastes safely. Burning waste in an incinerator is usually the most practical way for

safe destruction of laboratory waste. If safe burning cannot be arranged, discard the

waste in a deep pit of at least 1.5 meters’ depth. Access to the disposal site should

be restricted by building a fence around the site to keep animals and children away.

The burial site should be lined with a material of low permeability (e.g., clay), if

available and the location of the site should be selected at least 50 meters away from

any water source to prevent contamination of the water table. The site should have

proper drainage, be located downhill from any wells, free of standing water and not

in an area that floods.

If an autoclave is available, place infected materials inside and follow procedures

for safe and adequate sterilization.

• In addition, the underneath measures should be followed for waste disposal:

• Dispose of all biohazardous waste appropriately.

• Use dedicated leak-proof biohazard bags and bins for all potentially infectious

material.

• Discard biohazardous waste daily.

• Incinerate all solid waste after recommended disinfection.

• Liquid contaminated waste (e.g., human tissue, blood, feces, urine and other

body fluids) requires special handling. Carefully pour wastes down a utility

sink drain or into a flushable toilet and rinse the toilet or sink carefully and

thoroughly with water to remove residual wastes. If a sewage system doesn’t

exist, dispose of liquids in a deep, covered hole, not into open drains.

51

Decontaminate specimen containers by placing them in a 0.5% chlorine

solution for 10 minutes before washing them.

52

CHAPTER FOUR: Specimen Collection, Smear

Preparation, Fixation and Staining (Pre-

Examination Process)

Chapter Description

This chapter includes the pre-examination or pre-analytical procedures such as

blood specimen collection, smear preparation, fixation and staining of blood films

for detection and identification of malaria parasites.

4.1. Blood Sample Collection

Whenever possible, specimens should be collected for laboratory examination

before treatment is initiated. When malaria is suspected, blood smears should be

obtained and examined without delay. Blood sample for malaria parasite diagnosis

can be collected either from the capillaries or the veins (See the detail in Annex 3)

4.1.1. Capillary Blood Collection

Capillary blood is obtained by finger stick and is preferred to venous blood. Blood

obtained by pricking a fingertip is the ideal sample because the density of developed

trophozoites or schizonts is greater in blood from capillary-rich areas. For adults, the

best site to prick is the lateral side of the third or fourth finger of the non-dominant

hand (left hand unless the patient is left-handed) and the big toe and/or the heel is

preferred for infants. The skin area to be punctured should be warm so that blood

flow will be adequate. Depending on the physical settings and the patient’s

condition, warming the hand with warm water, covering the hand with a hot, wet

towel or briskly rubbing the hand may be used to warm the hands prior to the finger

prick. Label the frosted end of the slide with the patient ID number and date. Apply

gentle pressure again (do not squeeze the finger too tightly) to transfer more blood

and collect two or three larger drops (approximately 6 µl) on the slide, about 1 cm

from the drop intended for the thin film or 1 cm from the end of the slide.

4.1.2. Venous Blood

Venous blood is obtained by venipuncture. It is not collected for routine use in

malaria laboratory diagnosis. The venipuncture procedure is complex, requiring

both knowledge and skill to perform. Each phlebotomist generally establishes a

method that is comfortable for her or him. Several essential steps are required for

53

every successful collection procedure and venipuncture site selection. Although the

larger and fuller median cubital and cephalic veins of the arm are used most

frequently, the basilic vein on the dorsum of the arm or dorsal hand veins are also

acceptable for venipuncture. Palpate and trace the path of veins with the index

finger. Arteries pulsate, are most elastic, and have a thick wall. Thrombosed veins

lack resilience, feel cord-like, and roll easily.

If superficial veins are not readily apparent, you can force blood into the vein by

massaging the arm from wrist to elbow, tap the site with index and second finger,

apply a warm, damp wash cloth to the site for 5 minutes, or lower the extremity over

the bedside to allow the veins to fill. Foot veins are a last option because of the

higher probability of complications. One should recognize complications associated

with the phlebotomy procedure, assess the need for recollection and/or rejection of

sample and perform proper labeling of the specimen.

Venous blood samples provide sufficient material for performing a variety of

diagnostic tests, including concentration procedures (filariasis, trypanosomiasis).

However, in some parasitic diseases (e.g., for diagnosis of malaria in particular),

anticoagulants in the venous blood specimen can interfere with parasite

morphology and staining characteristics; this problem can be further compounded

by excessive delays prior to making the smears. In such cases, capillary blood

samples are preferable.

Note: Capillary blood is always preferred for malaria diagnosis than

venous blood

4.2. Blood Film Preparation

4.2.1. Types of Blood Films

Two types of blood films, thick and thin, are used in the microscopic diagnosis of

malaria. Both thick and thin films should be prepared and examined in all cases of

suspected malaria. For routine malaria microscopy, thin and thick blood films are

prepared on the same slide.

54

4.2.1.1. Thick Blood Film

Thick blood film consists of a thick layer of lysed erythrocytes. The blood elements

(including parasites, if any) are more concentrated (~30x) than in an equal area of a

thin smear, allowing a greater volume of blood to be examined. Because a larger

volume (6 μl) of blood is examined in the thick film, it is much better than the thin

film for detection of low levels of parasitemia and reappearance of circulating

parasites during infection, recrudescence or relapse. Thick film is therefore the most

suitable method for the rapid detection of the parasite, but it does not permit an

optimal review of parasite morphology for species identification. If the thick smear

is positive for malaria parasites, the thin smear should be used for species

identification. Thus, the thick films are performed to detect and quantify (parasite

density) malaria parasites in routine malaria microscopic diagnosis.

4.2.1.2. Thin Blood Film

Thin blood film consists of blood spread in a layer such that the thickness decreases

progressively toward the feathered edge of microscopic slide. In the feathered edge,

the red blood cells should be in a single layer, not touching one another. Thin blood

smear should be fixed with methanol so that the parasites are found intact inside

the RBCs. The morphological identification of the parasite to the species level is

much easier and provides greater specificity than the thick film examination.

However, low-density infections can be missed and require more time to read. Thin

blood film is used to assist in the identification of the malaria species after the

parasites have been seen in the thick film. Both thick and thin films must be

thoroughly dry. Allow the slide with the thin and thick films to dry inside a folder

rack in a flat, level position (which allows the thick film to dry with even thickness),

protected from flies, dust and extreme heat. Insufficiently dried blood film (and/or

blood films that are too thick) can detach from the slides during staining. Thin

smears will dry and be ready to fix and stain in about 15 minutes. Thick smears will

dry in a minimum of 30 minutes at room temperature. You can accelerate the drying

by using a fan or hair dryer (set on cool). Do not dry in an incubator or by exposure

to heat or sunlight as this will fix the blood cells and interfere with lysing the red

blood cells prior to staining.

55

Figure 4. 1 Unstained and stained blood films.

Qualities of good thick and thin films

A thin smear should

• Be uniformly spread over the slide

• Be thin enough so that it is tongue shaped

• Consist of a single layer of RBCs with a feathered end

A thick smear should

• Be 10 mm away from the edge of the slide

• Be round in shape with a diameter of about 10-12 mm

• Have a thickness containing 10 layers of RBCs

• Have 10-12 WBCs visible per oil immersion field of microscope

4.2.2. Common Mistakes in Making Blood Films

(a) Badly positioned blood films

Care should be taken that the blood films are correctly smeared on the slide. If they

are not, it may be difficult to examine the thick film. Also portions of the films may

even be rubbed off during the staining or drying process.

Figure 4. 2 .badly positioned blood film

56

(b) Too much volume of blood

After staining films made with too much blood, the background of the thick film will

be too blue. There will be too many white blood cells per thick film field, and these

could obscure or cover up any malaria parasites that are present. If the thin film is

too thick, red blood cells will be on top of one another and it will be impossible to

examine them properly after fixation.

Figure 4. 3. Too much blood for both thin and thick films

(c) Too little volume of blood:

If too little blood is used to make the films, there will not be enough white cells in

the thick film field and you will not be able to examine enough blood in the standard

examination. The thin film may be too small for use as a label for patient

identification.

Figure 4. 4. Too small blood for both thin and thick film

(d) Blood films spread on a greasy slide:

On a greasy slide blood films will spread unevenly, making the examination very

difficult. Some of the thick film will probably come off the slide during the staining

process

Figure 4. 5. A film made on a very greasy slide.

57

(e) Chipped edge of spreader slide

When the edge of the spreader slide is chipped, the thin film spreads unevenly, is

streaky and has many ‘tails.’ The spreading of the thick film may also be affected.

Figure 4. 6. the effect of chipped edge spreader on thin and thick films

(f) Thin film too large, thick film in the wrong place

If the thin film is too large, the thick film will be out of place and may be so near the

edge of the slide that it cannot be seen through the microscope. During staining or

drying, portions of the thick film will probably be scraped off by the edges of the

staining trough or drying rack. It may be very difficult or impossible to position the

thick film on the microscope stage for examination.

Table 4. 1 Most Common Technical Mistakes in Collection and Preparation of

Blood Smears

Mistake Effect

Pricking of non-dried finger The parasites and host cells may be

fixed by the alcoholic detergent

solution

Use of unclean or contaminated slides The blood smear will not be spread

evenly. Generates artifacts commonly

mistaken for malaria parasites,

including bacteria, fungi, stain

precipitation, and dirt and cell debris.

Delay in making blood film once you

transfer the drops of blood to the slide

The blood smear will not be spread

evenly due to the beginning of

coagulation process.

Too much blood for thin films

Erythrocytes are laid on multiple

layers. Observation is impossible.

Too little blood used for thin films Parasites may be virtually absent if

parasitaemia is low

Labeling the slide (Improper or no

labeling)

Error may happen- Confusion may

arise leading to slides that are

58

unidentifiable and cannot be linked to

a patient-

Slides are wrapped together before all

the thick films are properly dried

The slides stick to one another and

become unusable

Excessive time elapses between blood

collection and preparation of thick

films

Auto fixation occurs and hemolysis is

impossible.

Exposure of thick films to excessive

heat

Auto fixation occurs and hemolysis is

impossible

Thick films are dried too slowly P. falciparum gametocytes may

exflagellate

Inappropriate washing of stain from

the slide

Stain deposits may render the

observation difficult

4.3. Fixation of Blood Film

Fixation is used to preserve cellular and parasitic morphology. It is done when the

blood films are completely dried. Absolute methanol is a recommended fixative

solution.

Note: Auto-fixation may also occur spontaneously with time if thin films

not fixed immediately after being dried (7 to 15 days, varying with

humidity and temperature of the atmosphere)

4.4. Staining of Blood Film

4.4.1. Principles of Romanowsky Stains

Blood cells and parasites are stained by Romanowsky stains. Romanowsky stains

comprise two staining components: azures (oxidation products of methylene blue)

and eosin. Examples of Romanowsky stains include Field’s stain, Giemsa’s stain,

Leishman’s stain and Wright’s stain. Giemsa stain is regarded as the best stain for

malaria microscopy. Field’s stain is useful in health facilities with a low patient

workload as it is rapid, economical and easy to use. All Romanowsky stains can be

used to stain thick and thin blood films once the staining principles are understood.

4.4.2. Giemsa Stain

The Giemsa stain must be diluted for use with water buffered to a pH 7.2, depending

on the specific technique used. The stain should be tested for proper staining

reaction before use. The stock is stable, but it must be protected from moisture

because of the staining reaction. Giemsa stain will not function as expected if stock

59

is mixed with even small amounts of water or moisture solution during its

preparation or storage.

To control the quality of Giemsa stain for proper staining results, a known positive

smear should be included with each new batch of working Giemsa stain. Control

slides may be prepared from a patient’s blood and stored for future use. From a

patient known to have a malaria infection, a blood sample is collected in an EDTA

(ethylene diamine tetra acetic acid) or citrated blood tube if it requires multiple blood

film preparations or needs further diagnosis at a molecular level. An ideal quality

control blood film should have at least one parasite in every 2–3 fields on a thin

blood smear. Make as many thin smears as possible, preferably within one hour of

drawing the blood from the patient. Label the outside of the slide box with the

species, date and ‘Giemsa control slides.’ The slides can be stored at room

temperature but will last longer if stored at -20°C or -70°C. Just before use, remove

the slide from the box and allow the condensation to evaporate; label the slide with

the date and ‘+ control.’ The smear can then be stained and examined to check that

the working solution of Giemsa stain is of good quality.

4.4.2.1. Principle of Giemsa Stain

A properly stained blood film is critical for malaria diagnosis, especially for precise

identification of malaria species. Use of Giemsa stain is the recommended and most

reliable procedure for staining thick and thin blood films. Giemsa solution is

composed of eosin and methylene blue (azure). The eosin component stains the

parasite nucleus red, while the methylene blue component stains the cytoplasm

blue.

4.4.2.2. Preparation of Giemsa Stock Stain

To make about 500 ml of Geimsa stock solution, we need (Annex 4 and 5):

• Giemsa Powder ------------------------------------------------------------- 3.8g

• Absolute methanol---------------------------------------------------------- 250ml

• Glycerol---------------------------------------------------------------------- 250ml

4.4.3. Field’s Stain

Field’s stain is useful for rapid detection of malaria parasites particularly for thick

films. However, Schuffner’s dots are not always stained with this procedure. It is

60

made up of Field’s stain A and Field’s stain B as both are used in the staining

procedure.

4.5. Buffer Solution for Malaria Staining

A phosphate buffer solution, correctly balanced to pH 7.2, is essential for Giemsa

and Field’s staining for malaria parasites. Check the pH level using narrow-range pH

papers or a pH meter and store the buffer solution at room temperature. The buffer

is stable for several months. To check its quality, the pH of buffered water should be

checked, and appropriate correcting fluid should be added.

4.5.1. Buffer Tablets

Buffer tablets that produce a solution of pH 7.2 when dissolved are readily available

from laboratory suppliers but are rather expensive. Per the manufacturer’s

instruction, different grams of the buffer tablet are dissolved in a defined volume of

distilled water and used to prepare a Geimsa working solution (Annex 6).

4.5.2. Quality Control of Buffered Water

Prepare a buffer reagent carefully; weighing accurately the dry chemicals and

checking the pH level. Alternatively, use buffer tablets. Store buffered reagents at 2-

8ºC in a tightly stoppered (preferably plastic) bottle; when in use, avoid leaving the

reagents exposed to sunlight (which encourages the growth of algae) and check for

contamination (cloudiness) at regular intervals.

61

CHAPTER FIVE: Microscopic Examination and

Species Identification

Chapter Description

This chapter describes about the microscopic examination and species identification

of plasmodium species. Moreover, it describes quantification of the parasites.

Microscopy is the accepted standard method for detecting, identifying and

quantifying malaria parasites in the stained blood film. Microscopy requires trained

laboratory personnel and equipment (functional microscope) and other logistics.

Stained thick or thin blood films are examined for the presence of malaria parasites,

using an electric binocular microscope. In the absence of electricity, alternative

power sources must be used to ensure quality microscopy.

5.1. Examining Blood Films for Malaria Parasites

Microscopic examination of Giemsa stained thick and thin blood film is the most

acceptable method for detecting malaria parasites. This technique requires trained

and experienced laboratory personnel and equipment such as a microscope. The

thick film used for rapid detection of malaria parasites and enhances sensitivity for

the detection of low levels of parasitemia. On the other hand, the thin film is used

to confirm plasmodium species and also to assist in the identification of mixed

infections. Both thick and thin films should initially be examined completely with

low power magnification to avoid missing large organism such as Microfilaria and

Trypanosomes.

5.2. Systematic Approach of Examining Thick and Thin Blood Films

5.2.1 Examining the Thick Film

Thick films are performed to detect parasites and measure parasite density

(quantification), and can be used to monitor response to treatment. Parasites are

quantified by counting ring forms (trophozoites) against white blood cells. The

results are expressed as parasite count per 200 white blood cells (WBC) or parasite

count per microliter of blood, assuming a total white blood cell count of 8000/μl if a

measured white blood cell count is not available. This method of quantitation is

useful in low and moderate parasitaemia. Examination of a thick film requires

observation of 100 good fields. The blood film can be pronounced negative only

62

after no parasites have been found in at least 100 fields. If parasites are found, a

further 100 fields should be examined before a final species identification is made.

This ensures that there is little possibility of a mixed infection being overlooked.

Since the erythrocytes have been lysed and the parasites are more concentrated,

the thick film is useful for screening of parasites and detecting mixed infections.

Procedure for examination of thick blood film

1. Place the stained slide on the mechanical stage.

2. Scan the entire film at a low magnification.

3. Place a drop of immersion oil on the middle of the thick film.

4. Examine the film using the 100×.

5. Select an area that is well-stained, free of stain precipitate, and well-

populated WBCs (10-20 WBCs/field).

6. Move the blood film following the pattern shown in the diagram.

7. If you see parasites, make a tentative species determination on the

thick film and then examine the thin film to confirm the species

present.

Figure 5. 1 Systematic approach of examining thick blood film

Determination of "No Hemoparasites Found" (NHF)

According to the WHO malaria diagnosis, at least 100 fields, each containing

approximately 20WBCs, should be screened before reporting a thick blood film is

negative. Assuming an average WBC count of 8,000/μl of blood, this gives a

threshold of sensitivity of 4 parasites per microliter of blood and examination of the

above fields are sufficient. However, in non-immune or immunocompromised

patients, symptomatic malaria can occur at lower parasite densities, and screening

of more fields (e.g., 200, 300, or even the whole smear) might be necessary,

depending on the clinical context and the availability of laboratory personnel and

time.

63

5.2.2 Examining the Thin Film

Thin films are useful for species identification of plasmodium species. Examination

of thin film greatly assists in the identification of mixed infections and is also used

for quantification purposes.

Examination of thin films is recommended in the following circumstances:

• When the thick film is too small, has become auto fixed, or is not examinable

for other reasons.

• When it is necessary to confirm the identification of species.

Procedure for examination of thick blood film

1. Place a drop of immersion oil on the edge of the middle of the film.

2. Carefully examine the film using the 100×.

3. If doubtful diagnosis examine more fields

Figure 5. 2 Systematic approach of examining thin blood film

Thick blood film Thin blood film

-Lysed RBCs -Fixed RBCs

-Many layers -Single layer

-Large volume -Smaller volume

-Good screening test -Good species differentiation

-Low density infection can be detected -Low density infection can be missed

-More difficult to diagnose species -Requires more time to read

Table 5. 1 Characteristics of Thick and Thin Blood Films

5.3. Identification of Malaria Parasite Species, Other Blood Parasites and

Artifacts

A. Morphology, stages and distinctive diagnostic features of the plasmodium

species

The simplest guide of distinguishing between the four species of malaria is the effect

of the parasite on infected red blood cells. Diagnostic features to be considered

include the size of the red blood cell (whether it is enlarged or not) and presence or

absence of Schuffner’s dots or Maurer’s dots (also known as Maurer’s clefts) within

the cell. Schuffner’s dot refers to a hematological finding that is associated with

64

malaria, exclusively found in Plasmodium ovale and Plasmodium vivax.

Plasmodium vivax induces morphologic alterations in infected host erythrocytes

that are visible by light microscopy in Romanowsky-stained blood smears as

multiple brick-red dots. These morphologic changes, referred to as Schuffner’s dots,

are important in the identification of this species of malarial parasite. Maurer's dots

are any of the fine granular precipitates or irregular cytoplasmic particles usually

present in red blood cells infected with the trophozoites of Plasmodium falciparum.

Figure 5. 3 Basic components of a malaria parasite inside a red blood cell

Malaria parasites pass through a number of developmental stages. In all stages,

the structural features of the parasite stain the same color using Romanowsky

stains, as follows:

• Chromatin (part of the parasite nucleus) is usually round in shape and stains

red.

• Cytoplasm occurs in a number of forms, from a ring shape to a totally

irregular shape. It is always stained blue, although the shade of blue may

vary between malaria species.

• Malaria pigments stain in various shades from yellow-gold through brown

to black.

• Stippling stains in shades of pink which vary among different species.

B. Identification of Malaria Parasite Species and Stages

The trophozoites stage

This stage is the most commonly seen. It is often called the ring stage (although

sometimes it takes the form of an incomplete ring).

65

Figure 5. 4 Trophozoites stage of the malaria parasite

Because the trophozoites stage is a growing stage, the parasite within the red blood

cell may vary in size from small to quite large. Pigment appears as the parasite

grows. Malaria pigment is a byproduct of the growth or metabolism of the parasite.

It does not stain, but has a color of its own, which may range from pale yellow to

dark brown or black.

The schizonts stage

At the schizonts stage the malaria parasite starts to reproduce. This reproduction is

referred to as asexual because the parasite is neither male nor female but

reproduces itself by simple division. There are several obvious phases in this stage,

ranging from parasites with two chromatin pieces to parasites with a number of

chromatin dots and definite cytoplasm. These are seen clearly in the diagram below.

Figure 5. 5 Stages of schizonts growth

Red blood

Chromatin dot (red stain)

Cytoplasm (blue

Chromatin dot (red Cytoplasm (blue

Red blood

66

Note: The process of forming schizonts, which takes place in the liver and

in blood, is referred to as schizogony.

The gametocyte stage

The gametocyte stage is the sexual stage, in which parasites develop into either

male or female forms in humans for the preparation of the next stages, which takes

place in the mid-gut of the female Anopheles mosquito. Gametocytes may be either

banana-shaped (P. falciparum) or round (other species), and are either male

(microgametocytes) or female (macro gametocytes).

Figure 5. 6 Gametocytes of plasmodium falciparum and plasmodium malariae

C. Appearance of different malaria parasite species In thick blood films

In thick blood films, the staining process ruptures non-nucleated red cells but keeps

white cells, nucleated red cells and parasite structures intact. Parasites and white

blood cells can therefore be seen, although they may appear smaller and less well-

defined than in thin blood films.

In thick blood films, the fine rings of cytoplasm of trophozoites may appear

incomplete or broken. The absence of red blood cells makes Schuffner’s dots

(usually seen in Plasmodium ovale or vivax) more difficult to see. The “ghosts” of

red cells may be seen surrounding the parasites in the thinner part of the film.

In Thin Blood Films

Malaria parasites appear more well-defined in thin blood films, although it may be

more difficult to detect parasites in blood with low parasitaemia. Some species of

malaria parasites have an effect on the appearance of red blood cells in thin blood

films: for example, enlargement of red cells in P. vivax infection; or oval red cells in

P. ovale infection. Staining may also reveal Schüffner’s dots, or Maurer’s clefts

within the cells.

67

The tables below provide a reference for both thin and thick film species

differentiations.

Table 5. 2 Species Differentiation on Thin and Thick Films

Species differentiation on Thin films

Feature P.

falciparum

P. vivax P. ovale P. malariae

Enlarged infected

RBC

- + + -

Infected RBC

shape

Round Round,

distorted

Oval,

fimbriated

Round

Stippling infected

RBC

Mauer clefts Schuffner

spots

Schuffner

spots

None

Trophozoite shape Small ring,

Delicate

Large ring,

amoeboid

Large ring,

compact

Small ring,

compact

Chromatin dot Often

double

Single Single Single

Mature schizont Rare, 12-30

merozoites

12-24

merozoites

4-12

merozoites

6-12

merzoites

Gametocyte Crescent

shape

large, round large, round compact,

round

Species differentiation on Thick films

Feature P.

falciparum

P. vivax P. ovale P. malariae

Uniform

trophozoites

+

Fragmented

trophozoites

++ +

Compact

trophozoites

+ +

Pigmented

trophozoites

+

Irregular

cytoplasm

+ +

Stippling ("RBC

ghosts")

+ +

Schizonts visible Very rarely Often Often Often

Gametocytes

visible

occasionally Usually Usually Usually

68

5.4. Microscopic Differentiation

Microscopic differentiation of species depends on host cell and parasite

characteristics.

1. Features of infected red cells and ghosts

o Change in size, shape and color

o Presence of dots, Maurer's clefts (not on ghost cell) on infected

red cells

o Single or multiple infection of each cell

2. Parasite morphology at specific stages

o Number and size of chromatin beads

o Shape and size of cytoplasm

o Degree of pigmentation within cytoplasm

o Stages of parasite seen together

Diagnostic Points of Malaria Parasites

For P. falciparum

• Red Cells are not enlarged

• Maurer’s dots may be present

• No Schüffner’s dots

• Rings appear fine and delicate,

there may be several in one cell

• Some rings with 2 chromatin dots

• Presence of marginal or appliqué

forms

• Gametocytes have a

crescent/banana shape

• Usually trophozoites and

gametocytes are seen

• Schizonts are rarely seen

Plate 1 Diagnostic stages of P. falciparum

69

For P. vivax

• Red cells are usually enlarged

• Schüffner’s dots are frequently

present in the red cells as shown

above

• The mature ring forms tend to be

large and coarse

• Trophozoites schizonts and

gametocytes

• Developing forms are frequently

present

Plate 2 Diagnostic stages of P. vivax

For P. ovale

• Red cells are enlarged

• Comet forms are common (top

right)

• Rings are large and coarse

• Schüffner’s dots, when crescent,

may be prominent

• Mature schizonts similar to those

of P. malariae but larger and more

coarse

Plate 3 Diagnostic stages of P. ovale

For P. malariae

• RBCs are not enlarged

• Ring forms may have a square-like

appearance

• Band forms are a characteristic of

this species

• Mature schizonts may have a

typical daisy head appearance

with up to ten merozoites

Plate 4 Diagnostic stages of P. malariae

The appearance of Plasmodium stages in the different malaria species are shown

on Plates 1 – 4. Source: www.rph.wa.gov.au/malaria/diagnosis.html

70

Figure 5. 7 The appearance of different stages of Knowlesi compared to P.

falciparum and P. malariae stages in thin blood film

(source; https://cmr.asm.org/content/26/2/165#F4)

71

Figure 5. 8 Appearance of different species of plasmodium in a thin blood film

FIGURE 16 APPEARANCE OF DIFFERENT SPECIES OF PLASMO

Figure 5. 9 Appearance of Different Species of Plasmodium in A Thick Blood

Film Same as Above

72

Plate 1: Plasmodium falciparum stages in Giemsa-stained in thin and thick film

Source: WHO (1991). Malaria training module

73

Plate 2: Plasmodium malariae stages in Giemsa stained thin and thick film

Source: WHO (1991). Malaria training module

74

Plate 3: Plasmodium ovale stages in Giemsa-stained thin and thick film

Source: WHO (1991). Malaria training module

75

Plate 4: Plasmodium vivax stages in Giemsa stained thin and thick film

Source: WHO (1991). Malaria training module

76

Species Stage of parasite in peripheral blood

Trophozoites Schizonts Gametocyte

P. falciparum

Young, growing

trophozoites

and/or mature

gametocytes

usually seen

Size: small to medium

Number: often numerous

Shape: ring and comma

forms common

Chromatin: often two dots

Cytoplasm: regular, fine to

fleshy

Mature forms: sometimes

present in severe malaria,

compact with pigment as

few coarse grains or a

mass

Usually associated

with many young

ring forms.

Size: small, compact

Number: few,

uncommon, usually

in severe malaria

Mature forms: 12-30

or more merozoites

in compact cluster

Pigment: single dark

mass

Immature pointed-

end forms

uncommon.

Mature forms: banana-shaped or

rounded

Chromatin: single,

well defined

Pigment: scattered,

coarse, rice-grain-

like; pink extrusion

body sometimes

present. Eroded

forms with only

chromatin and

pigment often seen

P. vivax

All stages seen;

Schüffner’s

stippling in

‘ghost’ of host

red cells,

Size: small to large

Number: few to moderate

Shape: broken ring to

irregular forms common

Chromatin: single,

occasionally two

Size: large

Number: few to

moderate Mature forms: 12-24

merozoites, usually

16, in irregular

Immature forms

difficult to

distinguish from

mature trophozoites.

Mature forms: round, large

77

especially at

film edge

Cytoplasm: irregular or

fragmented

Mature forms: compact,

dense Pigment: scattered, fine

cluster Pigment: loose mass

Chromatin: single,

well defined

Pigment: scattered,

fine. Eroded forms

with scanty or no

cytoplasm and only

chromatin and

pigment present

P. ovale

All stages seen;

prominent

Schüffner’s

stippling in

“ghost” of host

red cells,

especially at

film edge.

Size: may be smaller than

P. Vivax Number: usually few

Shape: ring to rounded,

compact forms

Chromatin: single,

prominent

Cytoplasm: fairly regular,

fleshy

Pigment: scattered,

coarse

Size: rather like p. malariae Number: few

Mature forms: 4-12

merozoites, usually

8, in loose cluster

Pigment: concentrated mass

Immature forms

difficult to distinguish

from mature

trophozoites.

Mature forms: round,

may be smaller than

p. vivax Chromatin: single,

well defined

Pigment: scattered,

coarse. Eroded forms

with only chromatin

and pigment present.

P. malariae

All stages seen. Size: small

Number: usually few

Shape: ring to rounded,

compact forms

Chromatin: single, large

Cytoplasm: regular, dense

Size: small, compact

Number: usually

few; Mature forms: 6-12 merozoites,

usually 8, in loose

cluster, some

Immature and

certain mature forms

difficult to

distinguish from

mature trophozoites

Mature forms: round: compact

78

Pigment: scattered,

abundant, with yellow

tinge in older forms

apparently without

cytoplasm

Pigment: concentrated

Chromatin: single,

well defined

Pigment: scattered,

coarse, may be

peripherally

distributed. Eroded

forms with only

chromatin and

pigment present

Table 5. 3 Species Differentiation of Malaria Parasites by Cytoplasmic Pattern of

Trophozoites In Giemsa-Stained Thick Blood Films Species

5.5. Artefacts and Contaminants Confusing Malaria Parasites

Artifacts found on slides may include the following:

• Vegetable spores, yeast, pollen, algae and bacteria in the stain or on the slide

• Platelets

• Howell-jolly bodies in anaemic patients

• Ghosts of immature red cells mimicking Schüffner's stippling

79

Figure 5. 10 Blood elements, artefacts and contaminants that cause confusion.

5.6. Malaria Parasite Counting Methods

The determination of the number of circulating parasites is exceedingly important

for clinical purposes – to monitor the evolution of the disease and the efficacy of

therapy (See the detail in Annex 7).

Different methods have been used.

5.6.1. Number of Parasites/µl Of Blood (thick film):

Accurate parasite density estimation based on parasites per micro liter or white cell

count is necessary when parasite density determination is important for clinical

decision-making (for example in severe malaria or where monitoring of treatment

efficacy is required) and in clinical trials. It is recommended in routine practice that

parasite quantitation be performed against 200 or 500 WBCs. If, after counting 200

80

WBCs, 100 or more parasites are found, record the results in terms of the number

of parasites per 200 WBCs. If less than 100 parasites are found after counting 200

WBCs, parasite quantification should be continued until 500 WBCs are counted.

(This gives a probability <5% of chance variation greater than 25% of true parasite

density using a x100 oil immersion objective and an eyepiece with a field number

of 18). All parasites in the final field are counted, even if the count exceeds 500

WBCs. To determine parasite density, the parasite count is adjusted against the true

WBC count where available. If unavailable, a common practice is to assume a WBC

count of 8000/µL.

# Parasites/μl =# Parasites counted

WBCs countedxWBCs/μL

Example: Patient WBC = 8000/μL, and # of parasites counted against 200 WBCs =

650

Parasite count/μL =650

200x8000/μL

=26000 parasites/μl

5.6.2. Proportion of Parasitized Erythrocytes/Total RBC Counts (thin film):

This method will indicate the percentage of erythrocytes that are infected by malaria

parasites. The percentage of infected red cell is determined by counting the number

of red cells and that of parasitized red cells. This method of quantification is useful

in high parasitemia. Since it takes almost 10 times as long to examine a thin film as

to examine a thick film, routine examination of thin films is not recommended. The

number of parasitized erythrocytes (asexual forms) present in 25 microscopic fields

is counted and divided by the total number of erythrocytes present in these fields

(about 5,000), and multiplied by 100.

% Parasitemia =# of Parasitized RBCs

Total RBCs counted in 25 fieldsx100

Example: Average # of RBCs/25 field = 5000, and # of parasitized RBCs/25 fields =

100

% Parasitemia =100

5000x100

=2%

In this example 2% of the RBCs are infected with asexual forms of malaria parasites

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5.6.3 Number of Parasites/µl Of Blood (thin film):

This method requires the preliminary determination of the number of erythrocytes

present in the average microscopic field. The number of asexual parasites is counted

in at least 25 microscopic fields. The number of RBCs in the average microscopic

field is about 200, so total RBCs counted in 25 fields is roughly 200*25 = 5000. If the

hemogram is not available, RBCs/µL is assumed at 5,000,000 for males and

4,500,000 for females

Parasites/μl =# of Parasites

Total RBCs counted in 25 fieldsxRBCs/μl

Example: Parasites counted against 5000 RBCs (25 fields) = 50, i.e number of RBCs

in 25 fields is 5000, and # RBCs /μl = 5,000000 /male patient

Parasites/μl =50

5000x5000000

=50,000/μl of blood

5.6.4 Semi Quantitative Count (thick film)

This method has been used for a long period of time but currently it is not

recommended because it is less accurate.

Reporting:

• + 1-10 asexual parasites / 100 thick film fields

• ++ 11-100 asexual parasites / 100 thick film fields

• +++ 1-10 asexual parasites / single thick film field

• ++++ > 10 asexual parasites / single thick film field

5.7. Reporting Blood Film Results

Reporting of malaria positive blood film should include the species, stage and, if

necessary, the density of the parasite.

Species, stage and approximate number of parasites

1. P. falciparum

o Ring only + Number (per µl)

o Ring and gametocyte

o If schizonts seen, report immediately and mark with a red pen!

2. P. vivax, P. malariae, and P. ovale – all stages can be found.

3. Mixed infection – report appropriately

4. No hemoparasites found (NHPF) or Negative – if you do not find malaria and other

hemo-parasite after examination of a minimum of 100 oil immersion fields.

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Diagnostic Quality Control Depends Upon The: -

• Compliance with standards

• Availability of supplies, equipment, infrastructure

• Condition of the microscope

• Training of lab personnel and Regular supervision

• Quality of reagents and stains and Cleanliness

• Work load, Technical ability and type of techniques used

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CHAPTER SIX: Parasitological Diagnosis of

Malaria using Rapid Diagnostic Tests (RDTs)

Chapter Description

This chapter describes about malaria rapid diagnostic test (RDT) and its significance,

and the different formats and types of RDTs. Moreover, the principles, mechanism

of action and details about the procedures of RDTs are included in this chapter. The

strength and limitations of RDTs are also highlighted. RDTs are important alternative

method of malaria diagnosis in remote settings where laboratory equipment and

other facilities are not available.

6.1. RDTs and Their Significance

Rapid Diagnostic Tests (RDTs) or malaria rapid diagnostic devices detect malaria

parasite antigens in a small amount of blood by immunochromatography. In this

assay, monoclonal antibodies directed against the target parasite antigen and

impregnated on a test strip. The result, usually a colored test line, is obtained in 5

to 30 minutes. Malaria RDTs have the potential to greatly improve the quality of

management of malaria in areas where the gold standard method of diagnosis, high

quality microscopy, is not readily available.

In Ethiopia as in other malaria-endemic countries, there is an increasing support for

parasite-based rather than clinical diagnosis in places where microscopy is

unavailable, for instance at health posts. This is particularly the case due to the

implementation of Artemisinin-based Combination Therapies (ACT) and the

extension of health services to remote areas. RDTs are important for at least three

reasons to distinguish fever caused by malaria parasites from those caused by other

illnesses:

• Many life-threatening illnesses, such as meningitis and acute lower

respiratory infections, cause symptoms similar to malaria (fever, chills,

malaise, aches, etc.). Treating all febrile cases as malaria means that patients

with other conditions may not get the treatment they really need. When an

RDT shows that a febrile patient does not have malaria, the health worker will

manage the other febrile illness accordingly.

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• ACT is currently much more expensive than older antimalarial drugs such as

chloroquine (CQ), sulfadoxine-pyrimethamine (SP) and amodiaquine (AQ).

Rather than giving these expensive drugs to all patients with fever, RDTs can

help target ACTs to patients who really have malaria.

• Avoiding unnecessary use of ACTs on patients who do not have malaria may

help prevent or delay drug resistance; making ACTs effective for a longer

period.

6.2. RDT Versus Microscopy

Antigen-based RDTs can be performed by a wider range of health providers at all

levels of health care with minimal training. In addition, the results are available

immediately to the health provider. The tests have an important role at the periphery

of health services because none of the rural clinics has the ability to diagnose

malaria on-site due to a lack of microscopes and trained technicians to evaluate

blood films. However, RDTs are costlier per test than microscopy and should only

be used where microscopy is not available. It is not recommended to use RDTs

where there is a well-functioning laboratory, as microscopy provides additional

useful information such as parasite density, species identification, and support to

patient follow-up. Moreover, microscopy is more sensitive than RDTs.

Table 6. 1 Comparison of RDTs Versus Microscopy

Characteristics Microscopy RDTs

Format Slides with blood smear Test device

Equipment Microscope Kit only

Training Trained microscopists Anyone with a little training

Test duration 20-60 minutes or more 5-30 minutes

Test result Direct visualization of the

parasites

Color changes on antibody coated

lines

Capability Detects and differentiates

all Plasmodium species at

different stages

Detects malaria antigen

(pfHRP2/PMA/pLDH) from asexual

and/or sexual forms of the parasite,

but cannot differentiate the stages

of the parasites

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Detection

threshold

5-10 parasites/µl of blood 100-500/ µl for P. falciparum, higher

for non-falciparum

Species

differentiation

Possible Not possible by Pan specific RDTs

Quantification Possible Not possible

Differentiation

between

sexual and

asexual stages

Possible Not possible

Limitations Availability of equipment

and skilled microscopists,

particularly at remote

areas and odd hours

Unpredictable efficiency at low and

very high parasitemia; cross

reactions among Plasmodium

species and with auto antibodies;

persistence of antigens; HRP-2 gene

deletion

6.3. Malaria RDT Formats

RDTs are commonly prepared in three different formats: dipsticks, cassettes and

cards.

Dipsticks are impregnated nitrocellulose strips; it can be done by dipping in wells

containing blood. Cassettes are plastic cases containing nitrocellulose paper with a

test and control area. They are easy and safe to use and handle.

Cards -flaps containing nitrocellulose strips. Undergo sample wicking up the

nitrocellulose strip after application of the blood and reagent.

Figure 6. 1 Different formats of malaria RDT: A-cassette; B-dipsticks; and C-card

test

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6.4. Types of Malaria RDTs

Malaria RDTs can be categorized into two depending on how many species they can

detect. These are: Mono species (eg. Paracheck) and multispecies (eg. Carestart)

6.5. Basic Principles of RDTs

Rapid diagnostic tests are immune-chromatographic tests that detect specific

parasite antigens mainly Histidine Rich Protein II (HRP-2) or Plasmodium lactate

dehydrogenase (pLDH). Plasmodium aldolase is another antigen that is used in

some tests. Some RDTs can detect only one species (Plasmodium falciparum), while

others detect one or more species of malaria parasites that infect humans.

Antigens detected by currently used RDTs

1. Histidine Rich Protein II (HRP-2): is a protein produced by trophozoites and young

gametocytes of P. falciparum. A substantial amount of HRP-2 is secreted by the

parasite in to the host bloodstream and the antigen can be detected in erythrocytes,

serum, plasma, cerebrospinal fluid and even urine as a secreted water-soluble

protein. Tests based on HRP-2 detect only P. falciparum. HRP-2 has been shown to

persist and may be detectable for more than a month after clinical symptoms of

malaria have disappeared and parasites are cleared from the host. HRP-2 based tests

are relatively more stable at high ambient temperatures and humidity, and usually

less costly.

2. Plasmodium Lactic Acid (Lactate) Dehydrogenase (pLDH): is produced by both

trophozoites and gametocytes of malaria parasites. The pLDH antigen is present in

and released from parasite infected erythrocytes. pLDH is found in all four human

malaria species, and different isomers of pLDH for each of the four species exist.

Currently available pLDH RDTs detect pLDH specific to P. falciparum, P. vivax or are

pan-specific detecting all Plasmodium species that infect humans. Some pLDH RDTs

are specific for P. vivax. Since pLDH is disappeared from the circulation within five

days of successful antimalarial therapy, this test has the ability to differentiate

untreated from treated malaria, and may therefore be used for patient follow up,

although pLDH is also produced by gametocytes. Tests based on pLDH are less

stable at high ambient temperatures and humidity, and are more costly.

3. Plasmodium aldolase: is an enzyme produced in the glycolytic pathway by all

species of human Plasmodium parasites (pan-specific) and has been used in a

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combined P. falciparum/P. vivax immunochromatographic test. Tests that detect

aldolase appear to be less sensitive than tests that detect the other parasite

products. Aldolase behaves in much the same way as pLDH.

The various RDT’s appear to be similar; they vary considerably in their functioning

due to the intrinsic character of the critical components employed and their final

result.

Table 6. 2. Comparison of Rapid Diagnostic Tests for Malaria Antigens

6.6. RDTs Mechanism of Action

Though variations occur among malaria RDT products, RDTs are lateral flow

immune-chromatographic antigen-detection tests, which rely on the capture of dye-

labeled antibodies to produce a visible band on a strip of nitro-cellulose. With

malaria RDTs, the dye-labeled antibody first binds to a parasite antigen, and the

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resultant complex is captured on the strip by a band of bound antibody, forming a

visible line (test line). A control line gives information on the integrity of the

antibody-dye conjugate, but does not confirm the ability to detect parasite antigen.

The modes of action are indicated below:

• A dye-labeled antibody, specific for targeting antigen, is present on the lower

end of the nitrocellulose strip or in a plastic well provided with the strip.

Antibody, also specific for the target antigen, is bound to the strip in a thin

(test) line, and either antibody specific for the labeled antibody, or antigen, is

bound at the control line.

• Blood and buffer, which have been placed on the strip or in the well, are

mixed with labeled antibody and are drawn up on the strip across the lines

of bound antibody.

If antigens are present, some labeled antibody will be trapped on the test line.

Excess labeled antibody is trapped on the control line.

Figure 6. 2 Mode of action of antigen-detecting malaria rapid diagnostic tests

(RDTs).

source: who/RDT, ensuring quality and access for malaria diagnosis: how can it be achieved, www.nature. com/reviews/micro

6.7. General Procedures of Malaria RDTs

Malaria RDT's are designed and standardized primarily for testing specimen

obtained from fresh capillary whole blood and blood correctly collected through

vein puncture. Venous blood collected in appropriate anticoagulants, when stored

at 2-8oC may be stable for up to 72 hours provided they are not contaminated.

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The test procedure varies between the different test kits. Therefore, reference should

always be made to the specific manufacturer’s instruction. In general, the blood

specimen is mixed with a buffer solution that contains a hemolyzing compound and

a specific antibody that is labeled with a visually detectable marker such as colloidal

gold. In some kits, labeled antibody is pre-deposited during the manufacturing

process and only a lysing or washing buffer is added. If the target antigen is present

in the blood, a labeled antigen-antibody complex is formed and it migrates up the

test strip to be captured by the pre deposited capture antibodies specific for the

antigens and against the labeled antibody (as a procedural control). A washing

buffer is then added to remove the hemoglobin and permit visualization of any

colored lines formed by the immobilized antigen-antibody complexes.

A. For PfHRP-2 only test strips

• The PfHRP-2 test strips have 2 lines, one for the control and the other for the

PfHRP2 antigen.

• Invalid test result: no color change on the control line with or without a color

change on the test line.

• Negative: color change only on the control line and no color change on the

test lines.

• Positive: test for P. falciparum malaria with the PfHRP-2 test: color change on

both test and control lines. In certain situations, HRP2-detecting tests are less

sensitive, particularly for parasites that express little or no target antigen,

resulting in a false-negative result.

B. For PfHRP-2/pan-specific test strips

• The PfHRP-2/pan-specific pLDH or aldolase test strips have three lines, one

for control, and the other two for P. falciparum (PfHRP-2 or pLDH specific for

P. falciparum) and non-falciparum antigens (pan specific pLDH or aldolase),

respectively.

• With the PfHRP-2/ the pan-pLDH tests, a color change on the control line and

the pan-specific line indicates non-falciparum infection.

• A color change on all 3 lines indicates the presence of P. falciparum infection,

either as mono infection or as a mixed infection with non-falciparum species.

• If the PfHRP-2 line is visible when the pan-specific line is not, the test is

interpreted as positive for P. falciparum infection.

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Note: Mixed infections of P. falciparum with the non-falciparum species

cannot be differentiated from pure P. falciparum infections.

C. P. falciparum pLDH/Pan-specific pLDH or aldolase

• The P. falciparum pLDH/pan-specific PLDH or aldolase test strips have 3 lines,

one for control, and the other two for P. falciparum (pLDH specific for P.

falciparum) and non-falciparum antigens (pan specific pLDH or aldolase),

respectively.

• With the PfpLDH/ the pan-pLDH or aldolase tests, a color change on the

control line and the pan-specific line indicates non-falciparum infection.

• A color change on all 3 lines indicates the presence of P. falciparum infection,

either as mono infection or as a mixed infection with non-falciparum species.

• If the PfpLDH line is visible when the pan-specific line is not, the test is

interpreted as positive for P. falciparum infection.

Table 6. 3 Limitations of RDT Results

Repetition of the test after 1 to 2 days may therefore be indicated if illness persists

or if the patient’s condition deteriorates.

6.8. Test Procedure

Malaria RDT test procedure varies from manufacturer to manufacturer. Therefore,

adherence to manufacturers’ instruction is crucial. An example of the procedure of

one RDT product (CareStart multispecies) is illustrated below:

1. Seat the patient in comfortable position and explain to him or her what you

are about to do

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2. Wear a pair of gloves

3. Open the test kit pack

4. Take out the test kit and remove the desiccant

5. Clean the finger tip of patient’s left hand with alcohol swab

6. Make a prick with a sterile lancet and squeeze the finger tip

7. Wipe away the first drop of blood

8. Gently squeeze the pipette provided, immerse the end in the blood drop and

gently release the pressure to draw blood up to the black line/ring (5μl)

9. Place all the collected blood in the sample well of the test kit

10. Add two drops (60μl) of assay buffer into the buffer well

11. Read the test result in 20 min.

Result reading and interpretation: Multi species

6.9. Strengths and Challenges of RDT

6.9.1 Strengths

▪ Easy to use with minimal training

▪ Give rapid results & permitting immediate treatment in the site

▪ Do not rely on electricity and special equipment

▪ Do not require refrigeration

▪ Uses whole blood

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6.9.2 Challenges

▪ Costs per test exceed microscopy

▪ Short shelf-life,

▪ Requires effective transportation, storage and distribution systems

▪ Can’t estimate parasite density (qualitative test)

▪ Reinforce patient confidence in the diagnosis & health service

6.10. RDT Kit Selection and Handling

The specific performance requirements of a test will vary depending on the intended

usage. When considering whether or not to use malaria rapid diagnostic test in a

particular setting, it is important to consider their strengths and plan how to manage

the challenges. Important considerations in choosing an RDT include:

• The plasmodium species to be detected.

• Accuracy (sensitivity and specificity).

• Shelf-life and temperature stability during transport, storage and use.

• Ease of use

• Cost per test.

6.10.1. The Plasmodium Species to Be Detected

The appropriateness of P. falciparum-specific, pan-specific and non-falciparum

RDTs varies with the relative prevalence of the different human malaria species in

the intended area of use. These areas are categorized by the WHO as:

• Zone 1. P. falciparum only, or with non-falciparum species occurring almost

always as coinfections with P. falciparum. This includes most areas of sub-

Saharan Africa and lowland Papua New Guinea.

• Zone 2. Falciparum and non-falciparum infections occurring commonly as

single-species infections. This includes most endemic areas in Asia and the

America as well as isolated areas in Africa, particularly the Ethiopian

highlands.

• Zone 3. Areas with non-falciparum malaria only; this includes mainly vivax-

only areas of East Asia and central Asia and some highland areas elsewhere.

6.10.2. Accuracy (Sensitivity and Specificity)

A test method is said to be accurate when it measures what it is supposed to

measure. Accuracy of RDTs are expressed through several measures, the most

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widely used being sensitivity and specificity. These measures complement each

other.

Sensitivity: is the ability of a test to correctly identify individuals who have a given

disease or disorder. A high sensitivity of a malaria RDT means that it will produce a

true positive result when used in a population infected with malaria as compared to

the reference malaria gold standard test, microscopy. The sensitivity of an RDT for

detecting malaria parasitemia (or recent parasitemia) depends on the concentration

of circulating antigens in the patient’s blood, and the ability of the labeled antibody

on the RDT to bind the antigen and accumulate to form a visible line.

Specificity: Specificity is the ability of a test to correctly exclude individuals who do

not have a given disease or disorder. A high specificity of a malaria RDT means that

it will produce a negative result when used in a population not infected with malaria

as determined by the reference malaria gold standard test of microscopy. In other

words, it measures how often the test is negative when malaria is absent.

Both sensitivity and specificity are influenced by product storage and conditions of

use. In general, it is recommended that at least 95% of P. falciparum infections

should be detected at 100 parasites/µl and higher parasite densities, which is likely

similar to good field microscopy.

6.10.3. Shelf Life and Stability

To ensure that a product will retain its quality, it should be stored and transported

within the transporting requirements and necessary precautions. A longer shelf-life

reduces the pressure on the supply chain and the likelihood of wasting expired tests.

A minimum of 18 months of shelf life (e.g., at least 15 months after purchase) is

recommended since the RDT can be used at least for one-year period which means

for one major and minor transmission season in Ethiopian context.

6.10.4. Ease of Use

The intended conditions of use must be considered when choosing an RDT. If the

RDTs are to be used in a remote area without temperature-controlled storage,

stability (temperature requirements and shelf life) will be of great importance,

compared to storage and use in temperature-controlled laboratories. In Ethiopia for

example, RDTs are recommended to be used at health posts by health extension

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workers, since an easy-to-use format is more important in settings where

laboratories are absent.

6.10.5. Cost

RDTs are often costlier than microscopy and this should be taking into consideration

when deciding purchase in quantities and the level of use in the health care system.

Points to remember when performing RDTs

▪ Instructions should be strictly followed

▪ Blood-safety precautions

▪ Should be discarded if the envelope is punctured or damaged

▪ The test envelope should be opened only when it has reached ambient

temperature

▪ The RDT should be used immediately after opening

▪ RDT cannot be re-used

Negative Result Positive Result

Sometimes a negative test result does not

exclude malaria with certainty, because:

- There may be insufficient parasites to

give a positive result

- The RDT may have been damaged,

reducing its sensitivity

- Illness may be caused by another

species of malaria parasite which the RDT

is not designed to detect.

A positive result does not always signify malaria

illness, because:

- Antigen may sometimes be detected after the

infecting parasites have died (i.e., after treatment)

or due to the persistence of malaria gametocytes

which do not cause illness

- Presence of other substances in the blood may

occasionally produce a false-positive result

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CHAPTER SEVEN: Document and Record

Keeping

Chapter Description

This chapter will provide information for the training participants on the essential

elements of record keeping and the content of Laboratory request and report forms;

to provide information on how microscopy results correctly record in the laboratory

register.

7.1 Essential Elements of Recording and Reporting

Accurate document and record keeping in the malaria laboratory is essential.

Proper recording and reporting of patient data used to:

• Determine work load

• Planning of resources

• Compile important epidemiological information for malaria prevention and

control program.

7.2 Laboratory Request and Report Forms

In many countries, the laboratory request form and the microscopy report form are

combined into a single sheet of paper. This enables better tracking of reports and

reduces the time to transcribe the patient- and sample-related information on

separate report forms and hence reduces transcription errors.

A Laboratory request form must be submitted with the patient information and it

must match the information on the slides exactly. If the form is incomplete but the

patient is available, ask the patient for the required information.

A completed Laboratory Request Form should contain the following information:

• Name of the health facility

• Date (day, month, year)

• Patient’s name, address, age and sex

• Type of specimen

• Patient ID/ Specimen ID number (laboratory serial number)

• Clinical impression

• Signature of person requesting the exam

Microscopy Report

After the blood film has been read, immediately write the result in the result form.

Check that the specimen ID (laboratory serial numbers) on the slide matches that on

96

the laboratory request form. Subsequently, write the results in the Laboratory

Register, again checking to make sure that the laboratory serial number matches

exactly.

The microscopy report should include the following information:

• Specimen ID number

• Date and time of specimen Collection

• Date of specimen analysis (day, month, year)

• Blood film result (presence of parasite, species, stage and parasite

count/density against WBC)

• If necessary (high parasitemia) percentage of infected RBC in thin films

• Name and signature of laboratory technician performed the test

• Name and signature of laboratory technician reviewed the result

Once completed, the microscopy report should be made available as soon as

possible. Send the report to the referring health facility or clinician. Ensure that the

health facility or clinician receives the result. (See the detail in Annex 8)

7.3 Entry of Data into the Laboratory Register

Laboratory Register Book: The laboratory register book is a record book

maintained by the laboratory personnel responsible for blood film

examination of patients with suspected malaria. The health facility laboratory

register must include the following data for each patient with suspected

malaria:

• Laboratory serial number

• Date of specimen collection

• Date of specimen testing (day, month, year)

• Patient’s name, sex, age, and address

• Type of test

• Test result(s)

• Signature of person responsible for tests

Make sure all necessary columns are completed accurately. Reset the laboratory

register number to one on July 8 (Hamile 1st) (which corresponds to the start of the

Ethiopian fiscal year). DO NOT reset at the end of each day, week or month. Enter

patients successively, increasing the line number by one each time. The line

number is sufficient for identification of the request form.

97

Malaria Data Report

• Compiled from the laboratory log book on weekly, monthly, quarterly and

yearly basis.

• This will give us evidence on the workload of health facility laboratory for

further planning of resources.

• Have significant input to show the prevalence of laboratory confirmed

malaria cases in the facility.

7.4 Consequences of Incorrect Reporting False Negatives: A false negative is a result that was reported negative when it was

truly positive. Patients with malaria who receive a false negative result may not be

treated, resulting in ongoing disease, disease transmission and possibly death.

False Positives: A false positive is a result that was reported positive when it was

truly negative. Patients who receive a false positive result for malaria will be treated

unnecessarily, given the wrong medications (malaria medications) and in turn

medications will be wasted.

7.5 Importance of Malaria Data

Malaria data are usually compiled from a laboratory register on a weekly basis and

further summarized to receive monthly, quarterly and yearly data. Errors in

reporting will affect:

• Accurate determination of malaria cases at specific localities

• Determination of work load in specific laboratories

• Epidemic detection

• Malaria transmission trends based on confirmed cases

7.6. Laboratory Confirmed Malaria Case Report Form

• Compilation of microscopically confirmed cases can be done by reviewing

the malaria register and counting the number of blood films performed with

their results reported as positive with species name and negative slides.

• The report is usually submitted to the health facility manager and to the

respective health offices. This will assist to: -

o Determine the work performed at the end of each week, month, quarter

and year.

o Determine consumption of supplies based on the workload.

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o Provide epidemiological data on microscopically confirmed cases of

malaria at the health facility.

7.7 Malaria Laboratory Performance Report Form

• Done by reviewing the malaria register and counting the number of blood

films reported as positive with their species name and negative slides, and

age classification (under 5 and ≥5). (See the detail in Annex 9)

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CHAPTER EIGHT: Supply and Logistics

Management in Malaria Laboratory Diagnosis

Chapter Description

This chapter is designed to describe types and list of reagents and supplies required

for malaria microscopy diagnosis and its inventory control system.

8.1. Logistics Management

The establishment of an effective supply chain is essential to foresee and provide

all the equipment and supplies that are needed to sustain an uninterrupted flow of

reliable malaria diagnoses.

The main target of supply management is to ensure that patients always get the

laboratory services they need with concept of “No product, No program”.

For successful supply Management, the system must fulfill the Six rights of supply

chain management.

• The right products

• In the right quantity

• with the right quality

• On the right Place

• At the right time

• By the right cost.

8.2. Supply List for Malaria Diagnosis

Equipment and Materials:

1. Slide box

2. Staining jar (to hold 20 slides, placed back to back)/rack.

3. Drying rack

4. Forceps

5. Measuring cylinder

6. Slide boxes

7. Binocular microscope- Microscope spare bulb: 1 per microscope for 1

year, Microscope

8. Spare mirror, fuses, eyepiece, and oil immersion objective: 1 per

microscope for 10 years

9. Tally counter(s)

10. Funnel

100

11. Dropper (with rubber bulb)

12. PH meter

13. Timer

14. Beam balance

15. brown bottle

16. spatula/spoon

Consumables

1. Blood lancets

2. Absorbent cotton wool/cotton

3. Alcohol (70% ethanol)

4. Disposable gloves

5. Clean glass slides

6. Pencil/pen/marker

7. Sharps container

8. Biohazard containers

9. Distilled water/ buffered water

10. Lens paper

11. Immersion oil (Type A)

12. Lens cleaning solution

13. Filter paper (e.g., What man #1)

14. Measuring cylinder, capacity 100-500 ml (depending on the number of

slides to be stained)

15. Measuring cylinder, capacity 10-25 ml (depending on the amount of

stock stain to be measured)

16. PH tab

17. Beads

18. QC slides

19. Ethanol

20. Plaster

21. Weighting paper

Reagents

1. Absolute methanol

2. Giemsa powder

3. Glycerol

4. 10% Giemsa working solutions

5. Giemsa stock solution

Documents and records

1. Laboratory Registration book

2. Laboratory test request & result reporting form

101

3. Material safety data sheet (MSDS)

4. Bin card

5. Stock card

6. SOP

7. Job aid

8. Different formats

Supply list of Malaria RDT

1. RDT Device

2. Sterile Blood Lancet

3. Alcohol Swab

4. Sample transfer Pipette

5. Buffer

6. Glove

7. Timer

8. Sharp container

9. Biohazard bag

10. Labeling pencil / Pen/ permanent marker

11. Waste Bin

8.3. Logistics Management Information System (LMIS)

Three essential data items are required to run a logistics system

1. Stock on Hand: Quantities of usable stock available at a particular point in

time.

2. Consumption/Usage Data: The quantity of laboratory commodities used

during the reporting period (every two months).

3. Losses and Adjustments: Losses are the quantities of products removed from

the stock for anything other than the provision of laboratory services to

patients or those issued to another facility (e.g., expiry, lost, theft or damage)

and are recorded as negative (-) numbers.

Adjustments are quantities of a product received from any source other than

Ethiopian Pharmaceutical Supply Agency (EPSA) or issued to anyone other than

your facility’s laboratory. An adjustment may also be a correction due to an error in

mathematics. An adjustment may be a negative (-) or a positive (+) number.

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There are only three activities that happen to laboratory commodities within a

logistics system:

• Stored in inventory

• Moved between facilities

• Used to provide laboratory services to patients.

8.4. Stock Management

Stock management is properly maintaining adequate supplies to ensure

uninterrupted service. It involves:

• Performing a stock count (physical inventory)

• Maintaining proper inventory records

• Determining how much and when to order

• Placing orders properly

• Inspecting and verifying supplies received

• Ensuring proper storage of items on stock.

Stock management ensures the availability of staining reagents, supplies and RDT

kits which avoids the use of old reagents or expired RDT kits, and minimizes waste.

It is important not to under-stock or overstock supplies at the testing site

• Under-stocking will result in insufficient supplies while testing clients, which

interrupts the testing process.

• Over-stocking can result in storing of laboratory commodities for much

longer periods leading to deterioration or expiring of reagents and RDT kits

before use.

8.4.1 Inventory Control

An inventory control system is to inform personnel when and how much and what

types of a commodity to order and how to maintain an appropriate stock level to

meet the needs. A well-designed and well-operated inventory control system helps

to prevent shortage, oversupply, and expiry of commodities.

The inventory control system designed for the laboratory logistic system is a forced

ordering Maximum and Minimum inventory control system. Therefore, every

service delivery point (SDP) in the system is required to report at the end of every

other month and order all laboratory commodities back up to the maximum level. If

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stock levels ever fall below 2 weeks (0.5 months) of stock before the end of the

reporting period, an emergency order should be placed.

To maintain adequate stock levels, the maximum months of stock, minimum

months of stock and an emergency order point have been established for each

service delivery point in the system.

• The maximum months of stock signifies the largest amount of each

laboratory commodity a facility should hold at any one time.

• The minimum months of stock is the approximate level of stock on hand at

the end of the reporting period when an order is placed.

• The emergency order point is the level where the risk of stocking out is likely,

and an emergency order should be placed immediately.

8.4.2 Assessing Malaria Stock Status

A maximum/minimum inventory control system is a system designed to ensure that

quantities of stock fall within an established range – a maximum level and a

minimum level. In order to know if the stocks are within that range, you must assess

your stock status.

When you review the stock status, you determine how much of each laboratory

commodity you have available at your facility and how long these stocks will last.

You can review your stock status by counting the stock available, as you do during

a physical count. When you finish, you will have an absolute quantity of stock

available. But it is much more important to know how long the stocks will last and

if you have enough stock available until you receive your next order.

Months of stock is the number of months’ laboratory commodities will last, based

on the present consumption rate. When you review your stock status, you need to

determine how many months of stock you have in your facility.

Three months of stock means that your stock will last three months, as long as

consumption remains at the current rate.

Determining Months of Stock by calculating the months of stock, a facility can

determine if the right quantities of laboratory commodities are stocked. To

determine how long stock will last, the following simple formula can be used:

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8.4.3 Conducting a Physical Count

A physical count of the products is done to verify that the stock balance found on

the stock card shows the correct number of usable commodities that are available

in the storeroom. If the quantity on the stock card does not match the quantity on

the shelf, the stock card should be updated and an adjustment entered.

A physical count of laboratory supplies should be conducted ONLY at the end of

every month and the stock cards should be updated.

8.4.4 Conducting a Visual Inspection

A visual inspection should be completed each time products are handled: when

receiving, issuing or dispensing supplies, or when conducting a physical count.

When conducting a visual inspection, be sure to check the following:

• Package and product integrity: check for supply, missing or illegible

identification information.

• Labeling: make sure that products are labeled with the date of manufacture

or expiration, lot number and manufacturer’s name.

• Storage condition how the reagent is placed, temperature, humidity and store

area.

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8.4.5 Record Keeping

Efficient stock management depends upon accurate record keeping. Keeping

accurate records ultimately saves time.

Stock Card

The stock Card contains a list of all items in the store. It must be updated routinely

when orders are placed and received. It also serves as a reference to track orders

that have been placed and not received. The information recorded in the stock book

regarding when orders are placed and when they arrive may help a site to adjust

the reserve quantities of supplies that are kept on-site to ensure uninterrupted

testing.

Table 8. 1 Example of A Stock Book

Item

Physical

count

performed

Date

physical

count

performed

Quantity

Requested

(Units)

Date

Requested

Quantity

Reserved

Date

Received

Total

stalk in

hand

Expiry

date

Table 8. 2 Example of Stock Card

Item

required

Quantity

(units)

requested

Date

Requested

Quantity

Received

Date

Received

Lot number

expiry date

Quantity

used Balance

8.4.6 Calculation of Required Supplies

Calculations of supplies required for a malaria microscopy and RDT can be based

on the actual number of patients performed during a quarter and a stock count of

supplies on hand. This will give you more accurate information about the actual

condition. It is performed with a spreadsheet (see example worksheet below).

8.5. How to Calculate Required Supply Levels:

1. Determine the number of blood films or RDT performed in a quarter (A).

2. Determine the amount of each item required for a single blood film examination

or RDT (B).

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3. Multiply the two values (A x B = C).

4. Add a reserve quantity (D) of each item (C+D). Note: The reserve quantity can

be a fixed amount equal to the quantity of each item required for one quarter of

operation.

5. From that estimate, subtract the supplies you already have on hand, E ((C+D) –

E).

6. The result will be the amount of items you must order to ensure uninterrupted

testing during the next quarter of operation.

Key: A = the number of blood films examined in the quarter

• B = the amount of material required per test

• C = A x B

• D = Reserve quantity

• E = Stock at hand Stock needed = (C+ D) – E

Example; what is the number of slide required for the coming quarter?

• The # of blood films done in the quarter = 500 = (A)

• The # of slides required per test = 1 = (B)

• The # of reserve slide required = 500 = (D)

• The # of slide at hand = 100 = (E)

• Therefore: A x B = C →500 x 1 = 500

• C + D – E → 500 +500 - 100 = 900

• The required # of slides for the coming quarter is 900 pieces/18 boxes.

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Table 8. 3 Example of A Quarterly Supplies Request and Report, Requirement

Form

Table 8. 4 Example of A Quarterly RDT Supplies Requirement Form

You should assess your stock status any time you suspect that the stock levels do

not fall within the recommended maximum and minimum stock levels for your

facility. This may occur if there is a loss of supplies due to damage, expiry or theft

or if there is an unexpected increase or decrease in consumption.

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8.6. Storage of Malaria Laboratory Commodities

• Storage conditions will affect the quality of the laboratory products being stored.

Rooms that are too hot, stacks of cartons that are too high, and other poor

storage conditions can cause damage or cause a reduction in shelf life.

• A well-organized storeroom will simplify a facility’s work; time will not be wasted

trying to find needed supplies.

• Each commodity has a shelf life that is specified by the manufacturer. When the

commodity reaches the end of its shelf life, it has expired and should not be

distributed to patients or used in the laboratory. Some laboratory products have

short shelf lives. Because of these short shelf lives, it is important that proper

storage procedures are followed, so that the shelf life is protected. Always check

the expiry dates before issuing or using, and do not use products that have

expired.

• In some cases, a product will not have an expiration date on it, but it will have

the manufacturing date. By knowing the date, it was manufactured and the shelf

life of the product, one is able to determine the expiration date of the product.

8.6.1. Guideline for Malaria Laboratory Diagnosis Supply Storage

In order to manage storage of reagents and equipment:

• Keep staining reagents in well-closed bottles and out of direct sunlight.

• Kept tightly stoppered and free of moisture; stock Giemsa stain is stable at

room temperature indefinitely (stock stain improves with age).

• Make working Giemsa stain fresh daily. If a large number of smears are made,

the working stain may need to be changed throughout the day.

• Label all stock bottles containing staining reagents by name, date of

preparation and person who prepared it.

• When storing new microscope slides, make sure that they are as dry as

possible to prevent fungus growth.

• Store microscopes and their spare parts in a well-ventilated, dry, dark and

safe place. Spare bulbs should always be available at the laboratory, while

objectives, eyepieces, and other less frequently required parts can be stored

at regional level. Optical parts must be kept in a dry place to prevent damage

from fungus.

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• In general, supplies should be protected from sun, heat and water. Follow

manufacturer recommendations for storing supplies. This information is

usually printed on the product carton and boxes.

8.6.2. Handling Damaged or Expired Stocks

If expired or damaged stocks are found at any time during a visual inspection or a

physical count (or upon receipt of a consignment), these items should be removed

from the laboratory. These items also need to be moved to a separate place so they

cannot be dispensed or used. Damaged items should be safely disposed off.

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CHAPTER NINE: Quality Assurance of Malaria

Laboratory Diagnosis

Chapter Description:

This chapter generally describes the basic principles and components of malaria

quality assurance and different EQA methods that will be used to increase quality of

malaria testing laboratories using either microscopy or RDT methods.

9.1. What is Quality Assurance?

Quality Assurance (QA) is a system designed to improve the reliability and efficiency

of laboratory services, which are critical to the success of malaria control programs.

All parts of the testing system must be monitored to ensure the quality of the overall

process, to detect and reduce errors, and to improve consistency between testing

sites. To ensure reliability and to reduce errors, a quality system must address all

parts of laboratory testing.

The components of a quality assurance program for laboratory diagnosis of malaria

are the following:

Quality Control (QC): QC is systematic internal monitoring of work practices,

technical procedures, equipment and materials, including the quality of stains.

External Quality Assessment (EQA): EQA is a schematic assessment by an external

entity of a laboratory’s performance in testing of known and standardized but

undisclosed content and comparing the results with those of other participating

laboratories to assess laboratory practices and identify problems and weaknesses.

EQA includes onsite evaluation of laboratories, proficiency panel tests and blinded

smear rechecking.

Quality Improvement (QI): QI is a process by which the components of blood film

microscopy diagnostic services are analyzed with the aim of identifying and

permanently correcting any deficiencies. Data collection, data analysis, and creative

problem solving are skills used in this process

Generally quality assurance has three phases: pre-analytical, analytical and post

analytical phases as summarized in the figure below.

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Figure 9. 1 The quality assurance cycle source:

http://wwwn.cdc.gov/mpep/labquality.aspx accessed on 04/09/2012

9.2. The Need for Accurate Malaria Laboratory Diagnosis

Parasitological confirmation of the diagnosis of malaria provided by high-quality

microscopy or, where this is not available, by RDTs is recommended for all

suspected cases of malaria. In settings where malaria incidence is very low,

parasitological diagnosis for all fever cases may lead to considerable expenditure to

detect only a few patients who are actually suffering from malaria. In such settings,

health workers should be trained to identify patients that may have been exposed

to malaria (e.g. recent travel to a malaria endemic area, or lack of effective

preventive measures) and have symptoms that may be attributable to malaria

before they conduct a parasitological test. A parasitological confirmation of malaria

in stable high-transmission settings improves the differential diagnosis of fever,

improves fever case management, and reduces unnecessary use of antimalarial

medicines.

A high-quality microscopy service is one that is cost-effective, provides results that

are consistently accurate and timely such that they have a direct impact on

treatment. To achieve this, a comprehensive and active quality assurance (QA)

program is required. The primary aim of malaria microscopy QA programs is to

ensure that microscopy services are strengthened by competent and motivated

staffs, supported by effective training and supervision that maintain a high level of

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staff competency and performance, and by a logistics system that provides and

maintains an adequate supply of reagents and equipment.

9.3. Errors Compromising Quality Laboratory Diagnosis

Laboratory errors can be seen at three different vital steps of the procedure. The

forecast and recognition of each source of error may create a way to correct

inaccurate results.

The following are common sources of errors by phase:

A. Pre-analytical phase

• Incorrect test request or test selection (vague or ambiguous requests)

• Incomplete request forms

• Poor or inadequate patient preparation

• Poor method of specimen collection, labeling and transportation

• Use of dirty slides

• Defective equipment (microscope, weighing balances) and/or improper use

of equipment

• Substandard or expired reagents, and poor reagent preparation and storage

B. Analytical phase

• Poor procedure in blood film preparation and staining

• Inaccurate reading by unqualified or incompetent laboratory staff

• Lack of adherence to SOPs

C. Post- analytical phase:

• Poor reporting and recording

• Inaccurate calculations, computation or transcription

• Delay in reporting results to the clinician

• Incorrect results or misinterpretation of results

9.4. Objectives of Quality Assurance Programs

• To improve the overall performance of laboratory personnel at each level of

the laboratory system.

• To sustain the highest level of accuracy (in sensitivity and specificity) in

confirming the presence of malaria parasites.

• To systematically monitor Malaria laboratory diagnostic procedures,

reagents and equipment.

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The quality of Malaria Diagnosis Depends Upon

• Agreement with quality standards

• Availability of supplies, equipment’s and infrastructure

• Good working condition of the microscope

• The training of laboratory personnel

• Regular supervision

• Quality of reagents and stains

• Cleanliness

• Optimal work load, and

• Technical ability of laboratory technician and type of techniques used.

9.5. Challenges in Malaria Laboratory Diagnosis

It has been difficult to maintain good quality microscopy, especially at the periphery

of the health system where most patients are treated presumptively in spite of the

fact that the importance of light microscopy is well recognized. The current

challenges of malaria microscopy include the:

• Poor quality of microscope and RDT, particularly at the peripheral level.

• Difficulties in maintaining microscopy facilities in good order, logistic

problems and high costs of maintaining adequate supplies and equipment.

• Delays in providing microscopy results to requesting clinician or physician

• Lack of adequate training and retraining of laboratory staff.

• Poor QA system and supportive supervision of laboratory services.

9.6. Setting up a QA System

To set up a national QA system, the following should be considered

1. Description of the tiered laboratory network and responsibilities of each level

2. An inventory of available resources: The aim of this step is to establish the

minimum acceptable level of microscopy resources, including properly trained

technicians, functional microscopes, supplies, means of communication and

program supervision. Future and historic resources from national and

international stakeholders need to be included in this assessment.

3. Analysis of the annual number of slides collected at each laboratory, and an

estimate of the slide positive rate. Availability of the data.

4. Evaluation of the status and effectiveness of current QA activities.

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5. Comparative perspective: consider the adaptation of QA program of other,

comparable countries.

9.7. Principles of QA in Malaria Laboratory Diagnosis An effective QA scheme must be:

• Realistic, feasible and sustainable

• Compatible with the different situations and needs of each country

• A catalyst for change to a culture of quality

• Able to promote the best quality in the prevailing circumstances

• Able to appropriately recognize and accredit good performance

• Identify diagnostic settings (laboratories) and laboratory personnel with

serious problems that lead to poor performance

Generally, QA in malaria laboratory diagnosis includes: Correct specimen collection,

Preparation of good quality blood films, Staining using quality reagents from a

reliable source, Examination using a quality electric binocular microscope, Correct

interpretation and reporting

Therefore, incorrect, delayed, or misinterpreted tests: Have serious consequences

for patients and community, Undermine confidence in the diagnostic service, Waste

scarce resources

To implement QA for Malaria Lab Diagnosis; The health facilities or laboratory

personnel have to take training on malaria laboratory diagnosis and QA, should

avail all required equipment, laboratory commodities, and document and records,

then initiation of EQA. This process should be supervised and mentored by

expertise.

9.8. Components of Quality Assurance in Malaria Microscopy

Quality Assurance includes:

• Internal quality control (IQC)

• External Quality Assessment (EQA)

• Quality Improvement (QI)

(For more details refer to “Malaria Laboratory Diagnosis External Quality

Assessment Scheme Guidelines, EPHI, January2018.)

9.8.1. Quality Control (QC)

QC comprises those measures implemented by the laboratory during each test run

to verify that the test or procedure is correct and working properly, for example,

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checking the quality of the stain by use of a control blood film and Checking the

internal control line in RDTs

QC helps to ensure that the results produced by a laboratory are accurate, reliable

and reproducible. The QC program should be performed regularly and, to be

effective, the process must be practical and readily included in standard laboratory

reporting practices. All laboratory professionals are responsible for performing,

recording and reporting results of QC. Many components of QC are either performed

in conjunction with routine testing or as part of the regular quality management of

the laboratory.

IQC in malaria includes correct specimen collection by qualified laboratory staff;

preparation of good quality blood films; staining using quality reagents from a

reliable source; examination using a quality Microscope; and correct result

interpretation, recording and reporting.

The quality of Giemsa working solution should be checked before its use. Therefore,

IQC is performed for every new batch of Giemsa stock solution, when there is a

doubt in the quality of the reagent, and on a regular basis using control slides.

Preparation of Control Slides

• Collect a blood sample in an EDTA tube from a patient’s blood known to have

malaria infection (an ideal blood sample has at least one parasite in every 2-

3 fields on thin smears).

• If we can’t get known patient having malaria infection, we can prepare panel

slides from negative patient.

• Make as much blood films as possible from collected blood samples,

preferably within one hour of drawing the blood from the patient. Allow the

blood film to air dry at room temperature.

• Fix the thin blood film in absolute (100%) methanol and allow them to dry.

• Label the slide with date and “positive or Negative control

• Place them back to back in a slide box with separating grooves.

• Label the box with the species, date of preparation and “Giemsa Control

Slides”.

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• Cover the slide box containing control slides with plastic envelop and store

in -70 refrigerators for longer storage.

• The slides can be stored at room temperature for a maximum of one week.

Staining of Control Slides

If frozen, remove the slide from the box before use and allow the condensation to

evaporate at room temperature. The blood film can be then stained and examined

to check that the working solution of Giemsa stain is working properly or not using

the criteria listed below.

Evaluation The background of a well-stained thin film should be clean and free from

debris; the color of erythrocytes is a pale green pink and do not appear pink to red;

Neutrophil leukocytes will have deep purple nuclei and well-defined granules; the

chromatin of malaria parasites is a deep purplish red and cytoplasm a clear purplish

blue; Stippling should show up as Schuffner’s dots in erythrocytes containing P.

vivax or P. ovale, and Maurer’s spots in erythrocytes containing the larger ring forms

of P. falciparum.

In a well stained thick blood film, above 90% of the red cells should be completely

lysed. The background should be clean and free from debris, with a pale mottled-

grey color derived from the lysed erythrocytes; Leukocytes’ nuclei are observed as

a deep, rich purple; Malaria parasites are well defined with deep-red chromatin and

pale purplish blue cytoplasm. In P. vivax and P. ovale infections the presence of

Schuffner’s stippling in the “ghost” of the host erythrocyte can be seen especially

at the edge of the film. If the explained criteria are not met, the reagent preparation

techniques and the pH of the distilled water used for working solution preparation

should be checked. Stained blood film that is too pinkish suggests low pH or over-

staining and too bluish or purplish suggests high pH or under staining. All QC results

should be documented.

9.8.2. External Quality Assessment (EQA)

EQA refers to a system of objective checks of laboratory results by means of an

external agency. EQA schemes are effective means of assessing a laboratory’s

performance. The objective of EQA is to help laboratories identify errors and

improve practices for better performance.

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Effective EQA is a collaboration of laboratories at every level. The performance of

each laboratory is determined, areas of weakness are identified and corrective

measures undertaken. Intra-laboratory performance can be compared. Data or

information collected can be used for accreditation purposes and to evaluate the

use of certain laboratory equipment or techniques in the field.

Involvement in an EQA activity should not be seen as a threat, but rather as an

opportunity to strengthen skills. Most laboratory technicians want to provide

accurate testing. Good performance in EQA activities reassures them that their

results are contributing to accurate malaria diagnosis and control. EQA activities are

performed as per the National EQA structure. (For more details refer to “Malaria

Laboratory Diagnosis External Quality Assessment Scheme Guidelines, EPHI,

January 2018.)

National EQA Structure

1. National Level:

• Responsible for planning, budgeting, implementing and monitoring the

QA network.

2. Regional Level:

• Responsible for supervising and monitoring activities to maintain the

quality of the district and peripheral laboratories.

• Provide feedback of EQA scheme

• Planning and implementation of training and retraining activities

• Ensuring equipment is maintained in good working order.

• Ensure supply chain does not break down

3. Peripheral Level: Comprises

• Primary malaria diagnostic facilities (health posts) and

• Secondary malaria diagnostic facilities, such as laboratories within a

hospital or health centers that deal with inpatients and outpatients

There are three EQA methods for evaluating performance of malaria laboratory

diagnosis: proficiency panel testing blinded rechecking and on-site supervision.

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9.8.2.1. Panel Testing

Panel testing refers to the process by which the laboratory (known as the ‘test

laboratory’) performs malaria microscopy on a set of prepared slides received from

the reference laboratory. This exercise can check both the laboratory’s staining

procedure as well as the ability of the laboratory professional to recognize and

identify malaria parasites present.

Major advantage

• It provides a rapid picture of the proficiency of many laboratories in a country

(or region).

• The same set of panel slides will be distributed to all sites; Distribution of the

same panel to different laboratories will Identify sites most in need of

improvement and Allow comparison between sites.

Activities in Panel Testing

• Panel slides are arranged in set of ten slides which comprises five stained

and five unstained blood films.

• Source of panel slides are national and regional reference laboratories.

• Health facility laboratories at all levels of the public health laboratory system

in the public & private sectors are eligible.

• The laboratories are assigned a unique code number which is common to all

NEQAS.

• Panels are packed and shipped using standard procedures for handling

hazardous material.

• Feedback for participant laboratories will be sent within 30 days up on scoring

the results.

• A final summary report and improvement plan will be developed for

appropriate corrective actions.

Frequency

• Panel slides are distributed three times a year.

9.8.2.2. Blinded Rechecking

Blinded rechecking refers to the process by which a random selection of slides

collected from the ‘testing laboratory’ is reexamined at a higher level laboratory.

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Slides are checked for quality of blood film preparation (appropriate size and

thickness), quality of staining and accuracy of the result. Rechecking reflects the true

performance of laboratories offering routine diagnostic services at health facility

level. The purpose of the exercise is to allow a statistically valid assessment of the

proficiency of the peripheral laboratory.

Rechecking may detect malaria misdiagnosis in routine work and assess the overall

quality of testing. This should not be considered a criticism of the person who

performed the routine examination. Misdiagnosis in routine examination is more

frequently caused by different reasons such as high workload, poor equipment and

not necessarily lack of skill by the reader.

Each round of rechecking must be followed by feedback in the form of a written

report, showing details of incorrect scoring, if applicable, and offering suggestions

for quality improvement (corrective action). For the purpose of blinded rechecking,

slides are selected from those stored in the health facility.

Slide Storage in the Health Facility

• All positive and negative slides need to be stored in a slide box or dust free

carton box, and away from excessive heat and humidity until the slides have

been selected.

• Store slides consecutively according to laboratory number so there is a direct

link between the results in the laboratory register and the slide location.

• Selection must be done from the laboratory register and not directly from the

slide storage boxes.

Systematic Slide Selection Techniques

• Thirty slides per health facility should be re-examined every three months for

accuracy. The following selection technique should be applied during

sampling (See also example 1):

• Ten stained malaria slides are selected each month to determine accuracy: 5

positive slides and 5 negative slides.

• If less than 10 slides are examined in the facility, select all slides for

rechecking.

• If the number of positive slides examined is less, make up the difference with

negative slides.

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• Ideally malaria slides should be stored for 1 month and the selection made

before discarding the slides. The slide selection procedure will be conducted

on weekly basis by the laboratory head/quality officer using the procedure

described above (select slide from registration book and note the serial

number - put a mark on the register book to identify the selected slides).

• During collection of selected slides, the supervisors should counter check the

conformity of the selected slides with the laboratory registration book.

• Slides should always be stored for at least 1 week, to allow for patient follow

up. If slides are

• selected weekly, select as follows:

• Week 1 - randomly select 2 positive slides & 1 negative slide

• Week 2 - randomly select 1 positive slide & 1 negative slide

• Week 3 - randomly select 1 positive slide & 1 negative slide

• Week 4 - randomly select 1 positive slide & 2 negative slides

These numbers are the minimum sample size required for statistical analysis (see

below). More slides can be selected provided there is sufficient capacity for accurate

rechecking of all slides. Either the site supervisor or the facility laboratory personnel

should transfer the data of the collected 30 selected slides of each participating

health facility laboratory from the laboratory registration book into appropriate

form. (All forms for Blind Rechecking are available under Annex 10 and 11).

Frequency

Blind Rechecking is conducted quarterly bases; four times a year.

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9.8.2.3. On-Site Supervision

On-site supervision of malaria microscopy and RDT requires regular supervisory

visits to obtain a realistic picture of laboratory conditions and practices for malaria

microscopy. On-site supervision includes a comprehensive assessment of

laboratory organization, equipment, adequacy and storage of supplies, reagent

quality, availability and usage of SOPs, reading and reporting of results, and

infection

control measures – guided by the use of a supervisory checklist. On-site supervision

is the ideal way to obtain a realistic assessment of the skills practiced in the testing

laboratory or facility, to provide problem solving strategies and corrective action,

and assess the need for training. Supportive supervision includes assessment of test

performance, provision of on-site training and strengthening of services.

Malaria microscopy on-site supervision is conducted in accordance with NEQAS two

times a year by quality officers and malaria experts and others working on malaria

quality improvement. Onsite supervision provides an opportunity for basic

supervision, including assessment of laboratory supplies storage and inventory,

basic procedures, availability of functional equipment, quality of reagents, training

status of the laboratory staff, review of laboratory practical skills, work load, safety

and waste disposal system, performance of internal QC and result recordkeeping

practice. A major advantage of on-site supervision is the ability to identify sources

of errors detected by panel testing or rechecking and to implement appropriate

measures to resolve problems.

Sufficient time must be allotted for the visit to include observation of all the work

associated with malaria microscopy, including preparing films, staining, reading of

films by the laboratory personnel and examining a few stained positive and

negative films by supervisors to observe the quality of film preparation and staining

as well as condition of microscope.

On-site supervision checklist should be completed during the visit and discussed

with the test performer before the supervisor leaves the health facility, Filled

checklists should be submitted to the onsite supervision organizer after completion

of each visit, Feedback would be reported to each respective health facility in a

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month period and summary reports would be submitted to the regional &national

reference laboratories, Implementation of the feedback/corrective actions should be

followed up.

9.8.3. Quality Improvement

Quality Indicators for Malaria Microscopy

The following are quality indicators for malaria laboratory diagnosis:

• Laboratories should have SOPs, Job Aids and Bench Aids for malaria

microscopy diagnosis, and adhere to the procedures.

• Qualified staff

- Laboratories should have laboratory personnel trained on malaria

microscopy and RDTs

• Functional Equipment

Microscope is in good working order (electrical binocular microscope)

o Availability of maintenance and cleaning records

o Functional timer and tally counter

• Reagent Preparation and Storage

o Daily preparation of fresh working reagent from stock solution

o Storage according to the manufacturer instructions (Giemsa stain

should be stored in brown bottle)

o Reagents are labeled clearly with day of preparation/opening and

expiry dates

• Quality Control

o Check regularly the quality of every new batch of reagents using

known positive and negative blood films.

o RDT pre and post procurement lot testing

o RDT selection based on WHO recommended criteria

• EQA (External Quality Assessment)

o Laboratory participation in EQA

o Mechanisms for implementing corrective action, including retraining

• Correct Blood Film Specimen

o Completed request

o Labeled with unique ID and matching the request

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o The thin film should consist of a single layer of RBCs with a feathered

end, uniformly spread on the slide

- The thick film should be:

✓ round in shape

✓ 10mm away from the edge of the slide

✓ 10mm in diameter

✓ have 10-12 WBC per single 100x objective field

✓ newsprint or hands of a watch can be seen through the film

• Proper storage of slides for rechecking

✓ in a slide box

✓ away from excessive heat and humidity

✓ consecutively according to laboratory register number

✓ Free from immersion oil.

✓ Clearly labeled laboratory numbers

✓ Results should not be written on slides

Safety and safe waste disposal procedures (see Laboratory safety section for

details)

9.9. Quality Assurance (QA) of Malaria RDTs

A QA process for malaria RDTs aims to ensure high accuracy of test results in the

hand of end-users. This will include both monitoring of the technical standard of the

RDTs, processes to minimize environmental biosafety, and training and monitoring

of preparation and interpretation by end-users.

9.9.1. Planning for RDT Introduction

This requires a strategic plan with clear timelines to ensure that the various

components of the RDT program are in place at the right time. A Quality Assurance

coordinator (or coordinators) should be designated to oversee the overall

implementation plan and ensure that all agencies involved understand the process

and their particular roles, and that none are neglected. WHO has produced a

document “Good Practices for Selecting and Procuring rapid diagnostic tests for

Malaria” to guide country programs through the process of choosing, procuring and

planning for RDT rollout.

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9.9.2. Procurement

Procurement from manufacturers with ISO 13485:2003 compliance is

recommended.

Sensitivity and specificity are difficult to assess, as they are dependent on quality of

the test, the parasite density and other characteristics of the testing population, on

RDT preparation and interpretation, and on the quality of the reference standard.

Data on test accuracy should be obtained from the manufacturer and interpreted

with caution. In order to evaluate test sensitivity and specificity, lot-testing and field

monitoring are essential.

Thermal stability data should be obtained from the manufacturer and compared

with conditions of intended transport, storage and use. The parasite density (antigen

concentration) of the standard used to assess stability should be noted, as a heat-

damaged RDT may still detect samples with high parasite density.

Staggered delivery is useful policy (splitting delivery from the manufacturer into 2

or 3 batches several months apart), as it reduces the burden on central storage

facilities and allows new products to be received closer to the expected time of use,

shortening storage duration and effectively lengthening the shelf-life of the overall

procurement.

9.9.3. Lot Testing: Pre- and Post-Market

It is recommended that all lots (batches) of RDTs be tested before deployment to the

field. A ‘lot’ to be tested is normally defined as a production run using a particular

batch of monoclonal antibodies and nitrocellulose. Lots are usually defined by a lot

number provided by the manufacturer, and usually consist of 40,000 to 80,000 tests.

Lot testing can be done either

• Before purchase, directly arranged with the manufacturer and a lot-testing

center, or

• After purchase, before distribution to the field.

Lot testing is done

• To identify inter and intra-lot variation in specific lots of products

• To ensure no reduction in performance has occurred as a result of

inadequate storage conditions during transport to the country,

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• To convince stakeholders (clinicians, users, regulatory authorities) that tests

are in working order.

9.9.4. Monitoring Performance in the Field

Field monitoring is difficult, partly due to the inherent problems of accuracy of field

microscopy, with which RDTs must be compared.

At present, the following procedure is recommended:

• Compare RDT results with expert light microscopy. RDTs and blood films should

be taken from the same patients in selected health facilities where RDTs have

undergone typical storage, distribution and usage. E.g., every month, 40 RDTs

(20 positives and 20 negative) should be cross-checked against the

corresponding 40 BF obtained from the same patients and examined by an

expert laboratory technician. Where >10% discordant results occur, a more

detailed field evaluation should be rapidly performed or the remaining RDTs

should be returned for laboratory testing (see ‘lot testing’ above).

• It is important that the microscopists selected for the evaluation of RDT

performance are highly competent.

• In addition, it is important to supervise the health workers performing RDTs on

a regular basis at least every 3 months in order to evaluate the laboratory

personnel’s capacity of interpreting a set of prepared RDTs.

• Regular review of diagnosis and treatment records.

• Ensure that good blood safety practices are maintained.

• Ensure that sufficient supplies are in place for management of malarial and non-

malarial fever.

9.9.5. Training and Instructions for Users

Appropriate training of health workers prior to the introduction of RDTs is necessary.

Teaching instructions and instructions provided as job aids need to be clear, in a

locally-appropriate language, if required, and tested.

9.9.6. Use of Results and Community Education

• There is extensive evidence that RDT (and microscopy) results are frequently

ignored when treatment decisions are made. To address this problem, it is

essential to:

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• Ensure and demonstrate the accuracy of the RDTs (through the quality

assurance processes described above),

• Provide management algorithms for appropriate management of parasite

negative cases (non malarial febrile illness) and train health workers in their

use,

• Provide health workers with the means to manage parasite positive and

negative cases appropriately,

• Educate (sensitize) the community on the importance of parasite-based

diagnosis.

9.9.7. Storage and Transport

Standard supply management procedures should be applied to minimize storage

times and exposure to extremes in temperature, similar to those for the handling of

drugs. These include staggered delivery of large purchases, ‘First Expiry, First Out’

stock management principles, a temperature-controlled centralized storage, and

minimizing of storage in peripheral facilities lacking temperature control. Direct

exposure to sunlight should be avoided and transport coordinated to minimize

exposure to temperatures exceeding the manufacturer’s recommended storage

temperature.

9.10. Quality Assurance of Malaria RDTs in Remote Areas

The QA focus at this level should concentrate on initial training, supervision and

continuous education so that personnel working in remote areas achieve and retain

competence and motivation. Training should not only include test procedure

methodology but also trouble shooting guidelines, especially on how to suspect

RDT failure, and operating procedures for reporting suspected failed tests and recall

procedures of all proven failed batches of tests to the first referral level or

distribution point.

9.10.1. Ensuring Quality of RDTs

a. Pre-analytical phase

Important points to consider are:

- Ensure the quality of batches or lots for RDTs as they come into the country.

- Store and transport RDTs within temperature ranges recommended by the

manufacturer.

- Check expiration date of RDTs prior to use.

- Check integrity of packages prior to use.

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b. Analytical phase

- Ensure packages are opened only immediately prior to use.

- Ensure product instructions are accessible and tests are performed as

instructed by the manufacturer. - Read results within the time frame

stipulated by the manufacturer.

c. Post-analytical

- Ensure that RDTs are not re-used.

- Ensure that all used RDTs and accessories are discarded in a safe place for

incineration.

9.10.2. External Quality Assessment of Malaria RDTs

External quality assessment is one of the methods used to ensure that the RDTs are

handled and used in a correct way to provide valid and reliable results. Of all the

EQA methods, on-site evaluation is commonly used in our situation to assess the

storage conditions, observe how the test procedure is performed and on bio-safety

methods, using standardized checklist. Corrective measures are taken based on the

gaps identified and trainings organized when found necessary.

On-site supervision for malaria RDT should be performed three times a year by

supervisors of health extension workers and other partners working on malaria RDT

quality improvement. On-site supervision provides an opportunity for assessment

of RDT supply storage area and temperature, inventory, basic procedures including

sample collection, the RDT performance skills of the health extension worker,

internal quality control, result interpretation, reporting and recording, safety and

waste disposal, and need of retraining, by using a supervisory checklist.

A major advantage of on-site supervision is the ability to identify sources of errors

and provide on-site corrective action to improve the quality of result output and

implement appropriate measures to resolve problems.

Standardized checklists should be developed to assist supervisors during on-site

visits and to allow for the collection and analysis of standard data for subsequent

remedial action. Checklists should be reviewed and revised as needed to capture all

aspects of the testing process including laboratory related matters in order to

improve the entire process.

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9.10.3. Quality Indicators of Malaria RDT

• RDTs in use have been checked for the quality of batches or lots as they come

into the country.

• Storage and transport of RDTs is within the manufacturer’s recommended

temperature ranges.

• SOP and Job Aid are in place and the adherence to the procedure.

• Expiration date is checked and recorded prior to use of RDT.

• Integrity of the packages is checked prior to use (ensure that packages are opened

only prior to testing). An RDT should be discarded if its envelope is punctured or

badly damaged. If the procedure is delayed beyond the recommendation of the

manufacturer after opening the envelope/package, humidity can damage the

RDT.

• Tests are performed by personnel trained on malaria RDT and manufacturer’s

instructions are strictly followed.

• Test results are read only within the time limit specified by the manufacturer. Test

lines may become ‘positive’ several hours after preparation.

• RDTs are not re-used. - Participation in the EQA scheme (onsite supervision).

• Mechanisms for implementing corrective actions, including retraining, are in

place.

• All used RDTs are discarded in a safe place for incineration.

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CHAPTER TEN: Professional Ethics and Good

Laboratory Practices

Chapter Description:

This chapter is designed to equip laboratory professionals to increase core

competencies of laboratory ethics scientifically, professionally and culturally

acceptable laboratory services for patients /their customers.

Secnario

A SMALL TRUTH MAKES LIFE 100%

IF

A B C D E F G H I J K L M N O P Q R S T U V W X Y Z

IS EQUAL TO

12 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26

Luck: L+U+C+K = (12+21+3+11 = 47%

Love: L+O+V+E = 12+15+22+5 = 54%

Money: M+O+N+E+Y= 13+15+14+5+25 = 72%

Behavior: B+E+H+A+V+I+O+R = 2+5+8+1+22+9+15+18 = 80%

Leadership: L+E+A+D+E+R+S+H+I+P= 12+5+1+4+5+18+19+9+16 = 89%

• Then what makes 100%?

Is it Knowledge? ... NO!!

K+N+O+W+L+E+D+G+E =11+14+15+23+12+5+4+7+5 = 96%

Hard Work? ... NO!!! H+A+R+D+W+O+R+K= 8+1+18+4+23+15+18+11 = 98%

• Every problem has a solution, only if we perhaps change our attitude.

• To go to the top, to that 100%, what we really need to go further... a bit

more...

ATTITUDE A+T+T+I+T+U+D+E; 1+20+20+9+20+21+4+5 = 100%

It is OUR ATTITUDE towards Life and Work that makes OUR Life 100%!!!

Individual reflection

What is ethics?

What is health care ethics?

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10.1 What Is Ethics

The word ethics is derived from the Greek word ethos, meaning custom or

character. It is a major branch of philosophy which study values and customs of a

person or group. It is concerned with distinguishing between good and evil in the

world, between right and wrong human actions, and between virtuous and non-

virtuous characteristics of people.

It is concerned with what is right or wrong, good or bad, fair or unfair.

Ethics refers to standards of behavior that tell us how human beings ought to act

in the many situations in which they find themselves:

• As friends,

• Parents,

• Children,

• Citizens,

• Businesspeople,

• Teachers,

• Professionals, and so on.

• Ethics is not the same as feelings

• Ethics is not religion

• Ethics is not following the law

• Ethics is not following culturally

accepted norms

• Ethics is not science

▪ Moral Principles.

▪ What is good and bad.

▪ What is right and wrong.

▪ Based on value system.

▪ Ethical norms are not universal –

depends on the sub culture of the

society.

Ethics is not---

Ethics is ----

Ethics is a system governing human behavior

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Health Care Ethics:

It is a set of moral principles, beliefs and values that guide us to make choices about

healthcare. The field of health and healthcare raises numerous ethical concerns,

including issues of health care delivery, professional integrity, data handling, use of

human subjects in research and the application of new techniques.

10.2. Why is Ethics Important

The role of ethics in our society is very important because it is the basic beliefs and

standards that make everything run smoothly. Ethics are involved in all

organizations and institutions around us whether it be political, medical, lawful,

religious, or social.

Ethics influence and contribute to:

• Employee commitment.

• Customer loyalty and confidence.

• Customer satisfaction.

• The ability to build relationships with stakeholders.

• Cost control.

• Performance, revenue, and profits.

• Reputation and image: - “One of an organization’s most prized assets

is its reputation.”

10.3. Types of Ethics

Philosophers nowadays tend to divide ethical theories into different areas: but for

this purpose we will discuss about four types of ethics

1. Applied Ethics

2. Professional Ethics

3. Organizational Ethics:

4. Work Ethics:

Applied Ethics; - Is a discipline of philosophy that attempts to apply ethical theory

to real-life situations. It refers to the practical application of moral considerations. It

is ethics with respect to real-world actions and their moral considerations in the

areas of private and public life. For example, the bioethics community is concerned

with identifying the correct approach to moral issues in the life sciences.

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Examples of applied ethics: -

• Medical ethics (e.g. Bioethics)

• Business ethics

• Medical Laboratory ethics

• Journalism ethics

• Engineering ethics

• Legal ethics

Professional Ethics; - Is the moral principle, which should guide members of the

profession in their dealings with each other and with their patients, the patrons

(clients), the state etc. It encompass the personal, and corporate standards of

behavior expected by professionals Adherence to professional standards is

expressed through taking a professional oath and accepting professional code of

ethics Professionals and those working in acknowledged professions exercise

specialist knowledge and skill. How the use of this knowledge should be governed

when providing a service to the public can be considered a moral issue and is

termed professional ethics. It is capable of making judgments, applying their skills,

and reaching informed decisions in situations that the general public cannot

because they have not attained the necessary knowledge and skills. One of the

earliest examples of professional ethics is the Hippocratic oath to which

medical doctors still adhere to this day.

Organizational Ethics; is the application of morality related choices as influenced

and guided by values, standards, rules, principles, and strategies associated with

organizational activities.it is the ethics of an organization, and it is how an

organization responds to an internal or external stimulus. Example: -Uniform

Treatment of All Employees

Work Ethics: A standard of conduct and values for job performance. The importance

of developing a strong work ethic and how the work ethic you develop will impact

your future as an employee .it is also a set of moral principles that an employee uses

in the performance of his job. It also refers to how you feel about your job or career,

so it covers your attitude and behavior. It also pertains to how you do your job, or

the responsibilities that come attached with it. The level of respect you show your

co-workers and people you come into contact with at work, and how you

communicate and interact with them, also defines your work ethic. It refers to key

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characteristics that you should have, and they include honesty,

integrity, humility and accountability, among others.

Top ten work ethics are;

• Attendance

• Character

• Team work

• Appearance

• Attitude

• Productivity

• Organizational skill

• Communication

• Cooperation

• Respect

10.4. Elements of A Strong Work Ethics

A. Integrity: It means doing the right things, at all times, even if no one is watching,

much less your boss. Its greatest impact is seen in your relationships with the

people around you, which is why integrity is seen as one of the most important

ingredients of Trust.

B. Emphasis on Quality of Work; - If you show dedication and commitment to

coming up with very good results in your work, then your work ethic will

definitely shine. While some employees do only the barest minimum, or what

is expected of them, there are those who go beyond that.

They do more, they perform better, and they definitely go the extra mile to come

up with results that surpass expectations. Clearly, these employees are those

who belong to the group with a solid work ethic.

C. Professionalism: - The word “professionalism” is often seen as something that

is too broad or wide in scope, covering everything from your appearance to how

you conduct yourself in the presence of other people. It is so broad and

seemingly all-encompassing that many even go so far as to say that

professionalism equates having a solid work ethic.

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D. Discipline; Work ethic is something that emanates from within. You can tell an

employee to do this and that, be like this and like that, over and over, but if they

do not have enough discipline to adhere to the rules and follow through with

their performance, then there is no way that they can become the productive

employees that the company wants.Discipline involves focus, dedication and

determination on your part to do what you should.

E. Sense of Responsibility; - The moment you became part of the organization and

assigned tasks and duties, you have a responsibility that you must fulfill. If you

have a strong work ethic, you will be concerned with ensuring that you are able

to fulfill your duties and responsibilities. You will also feel inclined to do your

best if you want to get the best results.

F. Sense of Teamwork; - As an employee, you are part of an organization. You are

simply one part of a whole, which means you have to work with other people.

If you are unable to do so, this will put your work ethic into question. Work ethic

is also continuously shaped by relationships, specifically on how you are able

to handle them in achieving goals, whether shared or individual.

10.5. Principle of Ethics

The major principles of medical laboratory ethics are:

A. Autonomy: means independence and ability to be self-directed in health care.

Autonomy is the basis for the client’s right to self-determination. We have an

obligation to respect the autonomy of other persons, which is to respect the

decisions made by other people concerning their own lives. This is also called

the principle of human dignity. It gives us a negative duty not to interfere with

Ethical principles are the fundamentals of ethical analysis because they are the

viewpoints that guide a decision. There are four fundamental principles of

health care ethics.

1. Autonomy

2. Beneficence

3. Non-maleficence

4. Justice

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the decisions of competent adults, and a positive duty to empower others for

whom we’re responsible.

B. Beneficence: is the ethical principle which morally obliges health workers to

do positive and rightful things. It is “doing what is best to the patient”. Acting

in the best interests of patients (doing or promoting good). This principle is

the basis for all health care providers. We have an obligation to bring about

good in all our actions.

C. Non-maleficence: means to avoid doing harm. The principle refers to “avoid

doing harm”. Patient can be harmed through omitting or committing. Health

professionals should not inflict harm on patients. We have an obligation not

to harm others: "First, do no harm."

D. Justice: fair, equitable and appropriate treatment. Treat all patients equally –

no unfair discrimination. Justice refers to fair handling and similar standard

of care for similar cases; and fair and equitable resource distribution among

citizens. It is the basis for treating all clients in an equal and fair way. We have

an obligation to provide others with whatever they are owed or deserve. In

public life, we have an obligation to treat all people equally, fairly, and

impartially.

10.6. Core Values of Ethics

• Trustworthiness; - The trait of deserving confidence

• Respect: - Showing due deference to the innate dignity and value of others

• Responsibility: - That for which someone is responsible or answerable

• Fairness; - Consistent with rules, logic or ethics

• Caring: - Feeling and exhibiting concern and empathy for other

• Citizenship; - Exercising the duties rights, and privileges of being a citizen

10.7. Confidentiality and Informed Consent.

Confidentiality

Confidentiality in healthcare ethics underlines the importance of respecting the

privacy of information revealed by a patient to his or her health care provider, as

well the limitation of healthcare providers to disclose information to a third party.

The healthcare provider must obtain permission from the patient to make such a

disclosure.

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The information given confidentially, if disclosed to the third party without the

consent of the patient, may harm the patient, violating the principle of non-

maleficence. Keeping confidentiality promotes autonomy and benefit of the patient.

The high value that is placed on confidentiality has three sources:

• Autonomy: personal information should be confidential, and be revealed

after getting a consent from the person

• Respect for others: human beings deserve respect; one important way of

showing respect is by preserving their privacy.

• Trust: confidentiality promotes trust between patients and health workers.

Informed Consent

Informed consent is legal document whereby a patient signs written information

with a complete information about the purpose, benefits, risks and other alternatives

before he/she receives the care intended. It is a body of shared decision making

process, not just an agreement. Patient must obtain and being empowered with

adequate information and ensure that he/she participated in their care process.

For consent to be valid, it must be voluntary and informed, and the person

consenting must have the capacity to make the decision. These terms are explained

below:

A. Voluntary: the decision to either consent or not to consent to treatment must be

made by the person him or herself, and must not be influenced by pressure from

medical staff, friends or family. This is to promote the autonomy of the patient.

B. Informed: the person must be given all of the information in terms of what the

treatment involves, including the benefits and risks, whether there are

reasonable alternative treatments and the consequences of not doing the

treatment. This will help to avoid harm—patients may harm themselves if they

decide based on unwarranted and incorrect information.

C. Capacity: the person must be capable of giving consent, which means they

understand the information given to them, and they can use it to make an

informed decision.

10.8. Right and Obligations of Medical Laboratory Professionals

The Code of Ethics describes the expected ethical obligations and principles that

patients, the profession and the public believe will guide the professional and

personal conduct of all medical laboratory technologists (MLTs). These principles

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can be thought of more as exhibited behaviors than the knowledge and skills listed

in a Standards of Practice document.

Honesty

• In performing lab testing

• In reporting lab results

Dependability

• Taking your position seriously

• Being at work when you assigned to be

Kindness and Firmness

• Use compassion with patients and co-workers

• Remain firm in your duties and in doing what is right and best for your

patient

Humanity and Justice

• Be fair in running all patients’ laboratory tests

• Put yourself in the patient’s position before you argue or say something you

will regret

• Put yourself in your co-worker’s position before you argue or say something

you will regret

Maintaining Good Reports

• Use good hand writing when signing outpatient results

• Take your time to record all patient information in all log books

• Be honest in all your reporting

Adaptability

• Be willing to change work hours to help a co-worker

• Be willing to stay at work a little longer if patient work is not completed

• Be willing to make changes in testing procedures and other areas as needed

Co-operation

• If you expect others to co-operate with you, you need to co-operate with

them

• Team work will finish lab work quicker than an individual

Ethical Behaviors

• In all that you do, base your actions and decisions on ethical behavior

Rights of Medical Laboratory Professionals

• Safe working environment

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• Appropriate wage, allowance

• Legal protection

• Security

• Insurance for occupational hazard

• Access to medication

• Professional risk allowance

• Standard work load

• Proper leave (education, maternity, paternity, sick)

• Right to exercise the profession scientifically, ethically and legally

• The right to participate in health facility management and policy making

• Safety to new technology

• The right to receive Vaccination

• Be rewarded for innovative ideas

10.9. Professional Malpractice

Malpractice refers to Negligence or misconduct by a professional person. A

scientifically unsound or technically unjustified omission, manipulation, or

alteration of procedures or data.

An act or continuing conduct of a professional which does not meet the standard of

professional competence, and which results in provable damages to his/her client

or patient.

Such an error or omission may be through negligence, ignorance (when theprofes

sional should have known), or intentional wrongdoing.

Malpractice: Area of concern

Fraud:- the crime of obtaining money or property by deceiving people:

• The deliberate falsification of analytical or quality assurance results, where

failed method requirements are made to appear acceptable during

reporting.

• The intentional recording or reporting of incorrect information

• Fraud is purposeful and intentional

• Fraud is not a mistake.

• Fraud is an intentional misrepresentation of lab data to hide known or

potential problems.

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• Fraud makes data look better than it really is, with the intent to

deceive/mislead.

• Data manipulation

• Failure to follow Policy, SOPs/reference methods

• Falsifying existing data

• Incomplete record keeping

• Reporting data for samples not analysed

• Failure to respect customers

• Partiality/Injustice

• Unsafe practice

• Failure to adhere professional ethics

What are the Penalties for Malpractice?

Some Possible Legal Actions

• Suspension

• Civil Prosecution

• Criminal Prosecution

Suspension with pay: - is when an employee is sent home from work, usually while

receiving full pay. Employers are entitled to suspend an employee pending an

investigation of gross misconduct or other serious disciplinary matter.

Suspension without pay: -Suspension from work, without pay (unpaid suspension),

is the temporary removal of an employee from performing his/her work duties and

from receiving pay, as a disciplinary measure. Many employers who have

progressive discipline policies use unpaid suspension for employee misconduct:

such as theft, unsafe work behavior and company policy violations.

Civil Prosecution is a term used to describe a civil court action brought by one

person against another that may result in money damages being paid eg a libel

action or an action for wrongful death.

Criminal Prosecution: -The prosecution is the legal party responsible for

presenting the case in a criminal trial against an individual accused of breaking the

law

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Prevention of malpractice

• Create effective policies:

• Zero Tolerance – fraud is grounds for immediate dismissal

• Be Proactive:

✓ Develop a Code of Conduct and

✓ Provide Ethical declaration/ Agreement

✓ Training /orientation

✓ Provide job description

✓ Create effective communication system

✓ Write SOPs (manual integration

10.10. Ethics and Law

The Relation between Ethics and Law

• Law – the authority is external

• Ethics – the authority is internal

Ethics as discussed in the previous sessions, is considered as a standard of behavior

and a concept of right and wrong beyond what the legal consideration is in any

given situation.

Law is defined as a rule of conduct or action prescribed or formally recognized as

binding or enforced by a controlling authority. Law is composed of a system of rules

that govern a society with the intention of maintaining social order, upholding

justice and preventing harm to individuals and property. Law systems are often

based on ethical principles and are enforced by the police and Criminal justice

systems, such as the court system.

Ethics and law support one another to guide individual actions; how to interact with

clients and colleagues to work in harmony for optimum outcome; provision of

competent and dignified care or benefits of clients/ patients. Ethics serves as

fundamental source of law in any legal system; and Healthcare ethics is closely

related to law. Though ethics and law are similar, they are not identical.

Individual reflection

What is the relationship between ethics and law?

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Often, ethics prescribes higher standards of behavior than prescribed by law; and

sometimes what is legal may not be ethical and health professionals will be hard

pressed to choose between the two. Moreover, laws differ significantly from one

country to another while ethics is applicable across national boundaries. The

responsibilities of healthcare professionals and the rights and responsibilities of the

patient is stipulated in legal documents of EFMHACA like regulation 299/2013,

directives and health facility standards.

10.11. Good Laboratory Practices (GLPs)

Laboratory services are an integral part of disease diagnosis, treatment, monitoring

response to treatment, disease surveillance programs and clinical research.

Laboratory test results, therefore, should be reliable, accurate and reproducible.

Generation of such 'quality' results involves a step wise process of meticulous

planning, perfect execution and thorough checking of results by the whole team

involved.

Good Laboratory Practice (GLP) embodies a set of principles that provides a

framework within which laboratory studies are planned, performed, monitored,

recorded, reported and archived. In the clinical and research arena, the phrase good

laboratory practice or GLP generally refers to a system of management controls for

laboratories and research organizations to ensure the consistency and reliability of

results

GLP includes

• Data management (recording, reporting and archiving)

• Using Standard operating procedure (SOP)

• Safety in laboratories (to protect both staff and the environment

• Ethical considerations

• Quality assurance: the total process whereby the quality of laboratory

reports can be guaranteed.

• IQC

• External quality assessment

• Internal audit (identify problems and weak points in the system and suggest

remedial measures

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Key massage

Laboratory Staff; -you are the most critical part of the

✓ quality system

✓ The laboratory’s greatest asset

✓ An important partner in patient care

SO: Bring your integrity and professionalism to the healthcare community

References of the chapter

• Ethiopian Medical Laboratory Association (Ethiopia Medical Laboratory

Association) Code of Ethics for Medical Laboratory Technologists Practicing

in Ethiopia, 2008

• Medical Ethics Manual, World Medical Association, 2005

• James M. Gripando Nursing Perspectives and issue; Delmar publishers INC

3rd edition

• International Federation of Biomedical Laboratory Science (IFBLS) code of

ethics IFBLS general assembly of delegates, 1992

• Immigration Advisors Authority: Ethics tool kit, Dec,2013

• Introduction to Ethics, Medical ethics, Bioethics Main ethical approaches,

Prof. Marija Definis-Gojanović, October 201

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References

1. CDC. DPDx: Laboratory Identification of Parasites of Public Health Concern/Parasites and

Parasitic Diseases; Blood-borne Parasites: Malaria http://www.dpd.cdc.gov/dpdx, accessed

on March 15, 2009.

2. Chansuda W, Mazie JB, et.al. (2007). Review of Malaria Diagnostic Tools: Microscopy

and Rapid Diagnostic Test (RDT) American journal of tropical medicine and hygiene, 77,

pp. 119– 127.

3. Cheesbrough M (1998). District laboratory Practice in Tropical Countries. Part 1,

Cambridge University Press, UK.239-258.

4. Chotivanich K, Silamut K, Day N (2007). A Short review of methods; Laboratory

Diagnosis of malaria infection, New Zealand Journal of medical laboratory science, 61

(1):4-7.

5. EHNRI (2008). Acid fast bacilli sputum smear microscopy training module.

6. . EHNRI (2008). National AFB participant manual (Draft), April 2008.

7. EHNRI (2009). Malaria Laboratory Diagnosis External Quality Assessment Scheme

Guidelines. pp62.

8. EHNRI/TLCP (2008). Guidelines for Quality Assurance of Smear Microscopy for

Tuberculosis Diagnosis EHNRI/ TLCP - Federal Ministry of Health Ethiopia(DRAFT)

9. FMOH (1999). Health and Health Related Indicators.

10. FMOH (2008). Ethiopia National Malaria indicator survey 2007 technical summary-

Federal Democratic Republic of Ethiopia – Ministry of Health 2008

11. FMOH/ANVER/WHO (2007). Entomological profile of malaria in Ethiopia, September

2007.

12. http://chsr.aua.am/malaria/eng/diagnostics.php. Accessed on April 25, 2012

13. http://en.wikipedia.org/wiki/Sch%C3%BCffner's_dots. Accessed on April 25, 2012

14. http: //www.rph.wa.gov.au/malaria/diagnosis.html. Accessed on April 10, 2009.

15. IMaD (2008). Draft of National Guideline for Laboratory diagnosis of malaria, Ghana

Ministry of health.

16. IMaD (2008). Draft of SOPs for Laboratory diagnosis of malaria, Ghana Ministry of health.

17. Kakkilaya BS. (2003). Rapid Diagnosis of Malaria. Malaria site Lab Medicine. 8(34):602-

608.

18. Lawrence M. Tierney Jr., Stephen J. McPhee., Maxine A., Papadakis (2006). Current

Medical Diagnosis and Treatment, 15th edition (Kindle Edition).

19. Mandy FF (2004) General Laboratory Safety Issues, National HIV Immunology

Laboratory, Health Canada.

20. MOH (2006). National Five-Year Strategic Plan for Malaria Prevention and Control in

Ethiopia: 2006–2010 Federal Democratic Republic of Ethiopia Ministry of Health, Addis

Ababa.

21. MOH/EHNRI (2009). Master plan for the public health Laboratory System, Second

Edition, 2009-2013 Ethiopian Health and Nutrition Research Institute Federal Democratic

Republic of Ethiopia – Ministry of Health

22. National Strategic Plan for Malaria Prevention, Control and Elimination in Ethiopia, 2010

– 2015

23. PMI (2009). President malaria initiative, malaria operational plan, Ethiopia.

144

24. SCMS (2007). Standard operating procedures manual for the management of the national

laboratory logistics system to support HIV/AIDS prevention, treatment and support

programs, December.

25. SNNPR (2008). Implementation Guideline for malaria microscopic Diagnosis Quality

Assurance, SNNPRS Regional Health Bureau in collaboration with Malaria Consortium,

September 2008- Hawasa.

26. Transfusion, 2008; The American Red Cross

27. WHO (1991) Basic malaria microscopy. Part I. Learner’s guide. WHO, Geneva

(Switzerland).

28. WHO (2000) Bench aids for the diagnosis of malaria infection.

29. WHO (2006). The use of malaria rapid diagnostic tests, second edition.

30. WHO (2008) Methods Manual for Laboratory Quality Control Testing of Malaria Rapid

Diagnostic Tests, Version 5(a), Geneva, Switzerland.

31. WHO (2008). Malaria QA updates: Quality Assurance of Malaria Rapid Diagnostic Tests

buying well and maintaining accuracy.

32. WHO (2008). How to use Rapid Diagnostic Test (RDT) - A Guide for training at a village

level modified for training in the use of generic Pf- pan test for falciparum and Non

falciparum malaria.

33. WHO (2009). Malaria microscopy quality assurance manual. Version 1

34. WHO SEARO/WPRO (2005). Malaria Light Microscopy Creating A Culture of Quality

Report of Who SEARO/WPRO Workshop on Quality Assurance For Malaria Microscopy

Kuala Lumpur, Malaysia, 18–21 April 2005

35. WHO (1999). New perspectives Malaria Diagnosis report of a joint WHO/USAID informal

consultation - 25-27 October 1999 – World Health Organization – Geneva

36. WHO. Malaria slide bank project protocol (WHO regional office for the western pacific)

37. WHO/TDR/FIND (2008). Methods Manual for Laboratory Quality Control Testing of

Malaria Rapid Diagnostic Tests. Version Five.

38. WHO/USAID/FIND/TDR/. Malaria Rapid Diagnostic Tests. Available at

http://www.wpro.who.int/sites/rdt/home.htm

39. World Malaria Report 2010. Available at

http://whqlibdoc.who.int/publications/2010/9789241564106_eng.pdf

40. WHO (2010). Guidelines for the treatment of Malaria. 2nd edition.

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Annexes

Annex 1: Microscope: Types, Parts, Care and Handling A. Types of Microscope for Malaria Diagnosis

There are two types of microscopes:

a) Simple microscope the simple microscope is an ordinary magnifying glass which may

have a magnification of 5x, 10x, 20x or more.

b) Compound microscope A compound microscope has a much higher magnification than

the simple microscope. The typical compound light microscope is capable of increasing

our ability to see details 1000 times enlarged, so that objects as small as 0.1 micrometer

(µm) or 100 nanometers (nm) can be seen. This microscope uses at least two lenses

positioned at different places. A magnified image of the object is first produced by one

lens and this image is further enlarged by a second lens to give a more highly magnified

object. These two lenses are placed one at the end of each tube. The first lens which is

near to the object is known as the objective lens. While the second lens which is near

the eye is known as the eyepiece lens

Types of Compound Microscope

Based on the type of illumination system, different types of compound microscopes are

available:

• Light microscope

• Fluorescent microscope

• Dark field microscope

• Phase contrast microscope

Based on the available number of eyepieces, we can have at least two types of compound

microscopes:

a. Monocular microscopes

o Have a single eyepiece

o Are convenient for use by beginners, for field work where there is no electricity and for

photographing clinical specimens.

b. Binocular microscopes

o Have two eyepieces

o Are recommended where much microscopic work has to be done, i.e. in routine

examinations.

The total magnification power of a microscope is the magnification of its objective multiplied

by that of its eyepiece. For example, using a 10x objective and 10x eyepiece, the total

magnification of microscope is 100x.

The resolving power of a microscope is the ability of an objective to distinguish the dots

separately and distinctly. It is the limit of usable magnification.

For example

• The human eye can separate /Resolve/ dots that are 0.25 mm in diameter

• A light microscope can separate dots that are 0.25µm apart

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• The electron microscope can separate dots that are 0.5 nm apart.

Parts of a Compound Microscope

1. Microscope Stand

The stand of a basic microscope includes

Tube - Holds the eyepiece and objectives in line and at the correct distance

Stage - Is a flat surface where the specimen to be examined is placed. - In the center of the

stage there is circular hole that allows the light from the mirror or lamp to pass through

Mechanical stage - This enables the slide on which the specimen is mounted to be moved in

a controlled way, vertically or horizontally.

Sub stage - Immediately below the stage is the sub stage which holds a condenser lens with an

iris diagraph and a holder for light filters and stops.

Foot/Base - This ensures microscope stability on the laboratory bench.

Compound Microscope.

1. The Mechanical Adjustment System

Coarse adjustment

o Usually used to focus using low-power objectives

o Controlled by a pair of large knobs positioned one on each end of the body

o Rotation of these knobs moves the tube with its lenses or, in some microscopes, the

stage up or down fairly rapidly.

Fine adjustment

o Use to focus objectives for high-power objectives because they require a fine

adjustment

o Moves the objectives or stage up or down very slowly.

o Controlled/moved by two smaller knobs on each side of the microscope.

Condenser adjustment

o The condenser has an adjustment system for its focusing light onto the specimen on the

stage. This is done by opening and closing of its aperture.

o It can also be swung aside to remove it or to exchange it with another.

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o The condenser is usually focused by rotating a knob to one side of it.

3. Optics of a Light Microscope

Objectives

o Objectives are the most important parts of a microscope because the quality and most

of the magnification of the image depend on them.

o Modern objectives are described according to their magnification and older objectives

are often described according to their equivalent focal length (EFL)

For most routine medical laboratory work, 10x, 40x and 100x objectives are required.

The low power objective: 10x

o Used for initial scanning and observation in most microscopic works.

o Used for initial focusing and light adjustment of the microscope.

The high power objective: 40x

o Used for more detailed study as the total magnification with 10x eyepiece is 400.

o Used for the diagnosis of intestinal protozoal parasites, urine sediments/cells, casts

crystals, and histological sections.

The oil immersion objectives: 100x

o This lens has a very short focal length and working distance.

o The objective lens rests almost on a microscopic slide when in use.

o Known as oil immersion objective since a special grade oil must be placed between the

objective and the slide.

o Oil is used to increase the numerical aperture and the resolving power of the objective.

Ocular (Eyepiece)

o A lens that magnifies the image formed by the objectives.

o The usual magnification of the ocular is 10x, others are 4x, 6x, 7x, 15x and sometimes

as high as 20x.

o The higher the power, the greater the total magnification of the microscope. The lower

the power of the eyepiece, however, the brighter and sharper is the image.

Condenser

o A large lens with an iris diaphragm placed below the stage.

o It directs and focuses the beam of light from the light source, lamp or mirror, to the

specimen under examination.

o Usually consists of two or sometimes three lenses

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o The lenses are curved so that the light can pass to the objectives at a sufficiently wide

angle.

o The condenser position is adjustable; it can be raised and lowered beneath the stage and

the light must be correctly focused on the material to be examined.

Iris diaphragm

o It controls the amount of light passing through the specimen under examination.

o Located at the bottom of the condenser, under the lenses, but within the condenser body.

o It can be opened or closed as necessary to adjust the light intensity.

Mirror

o Used in the microscope without built in illumination

o It reflects the beam of light from the light source upwards through the iris in to the

condenser

The illumination system

o The modern compound microscope most often has a built-in illumination system with

a controller to adjust the amount of light comfortable for the microscopists.

B. Routine Use of a Basic Microscope Procedure:

1. Place the microscope on a firm bench and make sure it is not exposed to direct sunlight.

2. Select the source of light. If it is a built-in source, switch it on.

3. Place the specimen slide to be examined on the stage. Make sure the underside of the

slide is completely dry.

4. Select the objective to be used.

• It is usually better to begin examination with low power (10X) objectives. Once in

focus, all the other objectives also will be in focus provided that they are par focal.

5. Focus the objectives

• Move the objectives carefully downwards using the coarse adjustment knob and

looking at it from the side until the lens is near the specimen but not touching it.

• Then while looking through the eyepiece, move the objectives slowly upwards, still

with the coarse adjustment, until the image comes into view and is sharply focused.

6. Focus the condenser.

Open the iris of the condenser fully and, using the condenser adjustment knob, focus the

condenser on the detail of the light source until the image of the diaphragm appears sharp.

7. Adjust the opening of the condenser iris according to the specimen examined.

• Specimen like stained smears give off a little glare and for these the condenser iris

should be opened more widely giving a well-illuminated image with fine details.

• Unstained specimen like urine and saline preparations of stool give off a lot of glare

and require a reduced condenser iris to increase the contrast.

8. Examine the specimen using the mechanical stage to move it.

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9. For a higher magnification, swing the 40x into place. Focus the 40x objectives using the

fine adjustment.

10. For the highest magnification, add a drop of immersion oil to the specimen and swing the

100x oil immersion objectives in to place. Open the iris fully to fill the objectives with light.

Note: If examining a stained smear directly with the oil immersion lens and it is

not possible to focus it, remove the slide and check that the oil has been placed on

the smear side of the slide.

C. Setting of the Köhler Illumination for Light Microscope

1. Plug in the microscope and turn on the illuminator. Rotate the nosepiece so that the 10X

objective is locked into place.

2. Put the specimen slide on the stage and center it under the10X objective.

3. If there is a swing out (flip) condenser, be sure it is in the light path. Adjust the intensity of

the light to a comfortable level with the transformer.

4. Open the field diaphragm all the way and close the condenser diaphragm all the way.

5. Move up (rack up) the stage to its highest position.

6. Adjust the oculars for interpapillary distance so that when looking with both eyes only one

circle of light is seen.

7. Rack up the condenser as high as possible with the height adjustment knob.

8. Close the field diaphragm half way and focus on the specimen at 10X using the coarse

adjustment knob.

9. Close the field diaphragm until the diameter of the illuminated image is smaller than the

field of view. Note: If there is a flip condenser, it may need to be swung out at this time

to achieve this view of the illuminated image.

10. Lower the condenser with the positioning knob until a sharp, focused image of the edges

of the field diaphragm is achieved.

11. Using the centering screws on the side of the condenser, adjust the condenser so that the

circle of light is centered in the field.

12. Open the field diaphragm until the illuminated image is just larger than the field of view.

If more light is needed, use the transformer. Köhler illumination is now set.

D. Microscope Specifications

• Microscope must be completely UL*, CSA* and CE* tested, listed, and approved to

ensure fire and/or shock safety. Only UL listed components or line cords are not

acceptable. Must have10x/18mm eye pieces.

• Must have auto compensating Siedentop style binocular with diopter scale for

interpupillary distance (must have visible diopter scales).

• Must have 4-position reversed nosepiece of metal construction with internal ball

bearing stops. External clip system not acceptable.

• Must have 4x HI-Plan, 10x HI-plan, 40x HI-plan, and 100x oil HI-plan par focal and

par centered infinity corrected objectives.

• Mechanical stage must be of built-in design with metal rack and pinion X-Y drives. No

polymer belts, metal cables, timing belt systems or nonmetallic components are

acceptable in the drive mechanism. Coaxial controls must be low mounted for ease of

use.

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• Pre-aligned Abbe condenser with graduated iris diaphragm wheel with markings to

show where iris aperture should be set for each objective magnification.

• Focus drive must be a self-tensioning, three ball design of all metal construction. Fine

focus must have graduations of 100 divisions and 3 microns per division. Focusing

knobs on both sides must have these markings.

• All gears throughout the microscope: mechanical stage, focus, condenser rack and

pinion must be made of metal, brass, stainless steel or aluminum – no plastic

components.

• Illumination system must be designed for 12v/35w tungsten halogen 2,000-hour

average life bulb.

• Microscope must have hinged lamp door that is angled to help prevent breakage.

Sliding “drawer” type bulb covers not acceptable for safety reasons.

• Must have blue filter fixed into its mount, not loose. In Koehler kits, lollipop filters

have “locking slots” to prevent them from falling out when tilted.

• Microscope base temperature must not exceed 37 degrees centigrade using a 12v/20w

halogen lamp at full voltage for 6 hours.

• Power supply must be voltage sensing 85-265 volts with surge suppression and soft

start lamp control.

• Lamp intensity must be conveniently located in stand armrest and controlled via an

illuminated rotating wheel. Stage finger assembly is to be slide friendly that does not

damage or break slides.

• Microscope must have ergonomic design.

*UL: Underwriters Laboratories Inc.

*CSA: Canadian Standards Association

*CE: Conformance European

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Annex 2: SOP for Care and Preventive Maintenance of Microscopes Purpose This SOP provides guidelines for the proper use and preventive maintenance of

microscopes.

Principle A microscope magnifies minute objects making them visible to the eye. The

microscope consists of mechanical components, a system of lenses that magnify the specimen

placed on the microscope stage, and a light source that illuminates the specimen.

Materials, Reagents and Equipment

1. Lens cleaning solution (80/20 Ethyl Ether Alcohol)

2. Lens paper

3. Microscope

4. Plastic cover

5. Wooden storage Box

Procedure for Installation of Microscope

1. Place the microscope on a firm bench, free from vibration, near an electric power outlet

and away from direct sunlight.

2. During installation of new microscopes, follow manufacturer’s instructions.

Procedure for using microscope

1. Always follow the manufacturer’s instructions.

2. Connect to the power supply, and switch on the light source

3. Adjust the eyepieces by sliding them horizontally until both eyes fit comfortably and

the two fields merge.

4. Centre the condenser as follows:

• Swing the x10 objective into position.

• Raise the condenser to the uppermost position.

• Open the iris diaphragm fully.

• Open the light diaphragm to illuminate the whole field.

5. Clean and dry the underneath of a glass slide by wiping with cotton gauze.

6. Rotate the nosepiece so that the lowest power objective is in position. Slight resistance

is felt as the objective moves into the correct position.

7. Place the slide carefully on the stage.

8. Never place the slide on the stage when the x 40 or x 100 objectives are in position, to

prevent scratching of the lenses.

9. Adjust the illumination:

• Open the lamp rheostat fully to obtain a bright light.

• Reduce the iris diaphragm to control brightness.

10. Focus the specimen by racking the stage carefully upwards with the x10 objective in

position.

11. Using the coarse adjustment knob, rack downwards slowly using the coarse adjustment

knob until the image comes into view. Use the fine adjustment knob to focus the image

sharply.

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12. Swing the x40 and x100 objectives into position to examine in more detail using the

fine adjustment knob to focus.

13. After examination, lower the stage or swing the lowest power objective into position

before removing the slide.

• Never remove the slide when the x 40 and x 100 objectives are in position as this

may scratch the lenses.

14. Wipe off any oil from the lenses and microscope stage using a piece of lint free cotton

gauze soaked in lens cleaning solution (80/20 Ethyl Ether solution). Clean with lens

tissue.

15. Switch off the microscope, disconnect from the power source and cover to protect from

dust.

Procedure for Care and maintenance of Microscope

1. Always follow the manufacturer’s instructions carefully.

2. Clean the lenses with lens tissue and not a cloth or ordinary paper. Use lens cleaning

solution (80/20 Ethyl Ether solution) and not use xylene, methylated spirit or acetone;

these may dissolve the cement holding the lenses.

3. For removal of heavy contamination from the instrument surface, use a mild soap

solution – never use acetone.

4. At the end of every day, disconnect the power source by switching off at the wall socket

and removing the plug, or disconnecting the battery terminals.

5. Cover the instrument after use.

6. To protect against fungus in humid climates, place the microscope in a

7. Small cabinet or cupboard that is heated continuously from below by a low watt bulb.

Do not store the microscope in its carrying case or under a plastic hood in humid

climates.

8. Protect the microscope from power surges using a voltage stabilizer.

9. Replace blown bulbs, following the manufacturer’s instructions. 10. If the equipment

is faulty, consult a qualified biomedical engineer.

10. All microscopes in the laboratory must be scheduled for routine cleaning and check-

up daily using daily microscope maintenance chart. (Appendix 1)

Troubleshooting

1. Always refer to the operations manual.

2. If the microscope fails to switch on, check the electric socket outlet, plug and fuse or

the battery terminals.

3. Do not dismantle any part of the microscope. If the microscope is not

4. Functioning properly, consult a qualified biomedical engineer

Related procedures and documents

Microscope user manual

References

1. KEMRI, Kisumu Malaria SOPs, February 2006.

2. WHO Documents (CD). EQAS, September 2007

3. RITM, Parasitology Manual of SOPs, August 2007.

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Annex 3: SOP for Capillary Blood Collection and Preparation of Malaria

Blood Films

Purpose: This SOP provides instructions for capillary blood collection from the finger (or

earlobe or heel in infants) and preparing good quality thick and thin malaria blood films

(MBFs).

Principle: Capillary blood obtained by direct pricking of the finger (in adults), or the earlobe

or heel of the foot (in infants). The blood is used immediately to make thin and thick blood

film therefore it does not need anticoagulant.

Materials and supplies

1) Alcohol (70% ethanol)

2) Disposable sterile lancets

3) Absorbent cotton

4) Disposable gloves

5) Clean frosted end glass slides

6) Lead Pencil/Glass writing pencil

7) Slide drying Tray

8) Biohazard containers (for infectious waste)

9) Sharp Container

10) Patient Register

Safety Precaution

1. Wear protective gloves when handling or taking blood samples.

2. Cover any cuts or abrasions on your hands with adhesive dressing.

3. Always wash your hands with soap and water after handling blood sample.

4. If blood gets on to your skin, wipe it off quickly with cotton wool soaked with

alcohol and wash the affected area with soap and water as soon as possible.

5. Take care not to accidentally prick yourself.

6. Never use disposable lancets more than once.

Procedure of blood collection using capillary

1 Label the frosted end of the slide with the patient ID number and date.

2 Disinfect the finger (in adults) or the earlobe or side of heel (in infants) thoroughly

with an alcohol swab.

3 Let the alcohol air dry.

4 Prick the finger/earlobe/heel with a disposable sterile lancet, deep enough for the

blood to flow freely.

5 Wipe the first drop of the blood with dry cotton.

6 Apply gentle pressure to the finger/earlobe/heel for the blood to flow

7 Discard used lancets directly into the sharps disposal container.

8 From the pricked finger/earlobe/heel, collect blood directly in to the pre-labeled

glass slides

9 Make both thick and thin blood films on the same slide as follows:

10 By touching the slide on the blood, place a small drop(2µl) of blood in the middle

portion of the slide and 1 bigger drop(6µl) on the portion next to the frosted end.

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Allow some space between the thick and thin films to be made on the same slide

(See Appendix Figure 3a).

Procedure for preparation of the thin film (See Figure 3c, Illustration 1 and 2 below).

1. Working quickly, obtain a second clean and polished slide (spreader) and place in

front of the small blood drop at a 30º - 45º angle. Pull back the slide and hold until

the blood is evenly spread along the edge of the slide. Do not delay between

applying and spreading the drop.

2. Rapidly push the slide forward in a single, smooth, continuous motion. Avoid

hesitation or jerky motions when spreading the blood. (A feathered end of the film

should have red blood cells that are lying individually without overlapping and

relatively evenly distributed).

Procedure for Preparation of thick blood film (See Figure 3c, Illustration 3 below).

1. With one corner of the spreader slide, in a circular motion, spread the blood out to

make a circle with approximately 1cm (1/3 inch) in diameter, finishing off at the

center.

2. 2. The ideal thickness of the smear should allow for printed text to be readable when

it is placed on it. 3. Discard the spreader into an appropriate slide container and

DON’T re-use it for another patient’s blood sample.

3. Allow both blood films to air dry in a horizontal position on a slide tray. Slow drying

prevents cracking. Avoid using a fan or blow dryer to dry these slides.

Procedural Notes

A number of errors are common in making blood films. These can affect the labeling, the

staining or the examination.

a) Badly positioned blood films

Care should be taken that the blood films are correctly sited on the slide. If they are not, it may

be difficult to examine the thick film. Also, portions of the films may even be rubbed off during

the staining or drying process.

b) Too much blood

After staining films made with too much blood:

• The background to the thick film will be too blue.

• There will be too many white blood cells per thick film field, and these could obscure

or cover up any malaria parasites that are present.

• If the thin film is too thick, red blood cells will be on top of one another and it will be

impossible to examine them properly after fixation.

c) Too little blood

If too little blood is used to make the films:

• There will not be enough white cells in the thick film field and you will not examine

enough blood in the standard examination.

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d) Edge of spreader slide chipped

When the edge of the spreader slide is chipped:

• The thin film spreads unevenly, is streaky and has many “tails”.

• The spreading of the thick film may also be affected.

e) Thin film too big & thick film in the wrong place

• The thick film will be out of place and may be so near the edge of the slide that it

cannot be seen through the microscope.

• During staining or drying, portions of the thick film will probably be scraped off by

the edges of the staining trough or drying rack.

• It may be very difficult, or impossible, to position the thick film on the microscope

stage so that it cannot be fully examined.

Quality Control

Monitor the quality of the preparation of thick and thin smears

1. Follow proper collection procedures.

2. Glass slides must be clean and free from grease.

3. Thick films and thin films must be prepared properly while drying protects blood

films from dust, flies and insects.

4. Do not dry expressed to direct sun light.

5. Too thin a film may not have adequate quantity of blood for detection of parasite.

6. Blood film spread unevenly on a greasy slide makes examination difficult.

7. Thin film too long, leaves less space for thick film.

8. When fixing the thin film, be careful not to let methanol touch the thick film.

9. Wet slides are wrapped together and the slides stick to one another.

10. Never add a pinch of EDTA powder directly to the sample tubes. High concentration

of EDTA leads to shrinking of RBC and destroys the structure of WBC and platelets

11. Never add the blood before the EDTA solution is completely dried. It will dilute the

blood

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Illustrations

Illustration of Blood film preparation

a. Template for Thick and Thin Malaria Blood Films

• The edge of a clean slide is placed at about 45 angle in front of the smaller blood drop

for thin film (see Illustration below).

• Slowly pull this second slide back into the drop while securing the sample slide with

the forefingers of the other hand.

• Barely touch the drop of blood and, as the blood spreads laterally along at least two

thirds of the edge of the “spreader” slide,

• rapidly push the spreader slide forward in a smooth, continuous and rapid motion, not

stopping until the clean slide leaves the bloody slide.

• A properly prepared thin film is thick at the beginning end and thin or "feathered" at

the other end. The feathered end of the smear should not reach to the end of the glass

slide. The feathered end should have areas optimal for microscopy that are only one

cell layer thick.

• The thin smear is best prepared immediately after applying the drop of blood, before

any drying occurs.

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d. Illustration of thin Blood film making

The clean slide was placed just before the blood drop (to the right) then pulled back (to the left)

and pushed forward to the right leaving a feather edged thin film. The blood for the thick film

remains untouched at this stage.

Use the corner of the same clean slide to make the thick film by gently swirling the drop of

blood to form an even circle of approximately 10mm diameter using the paper template over

which the slide is placed during slide preparation. Once the drop(s) are evenly spread, lift the

corner of the clean slide out of the center of the smear, trying not to leave any bubbles. If

bubbles are present, stir again with the corner of the slide until no bubbles remain, and/or break

the bubbles with the sharp corner of the spreading slide.

e. Illustration of making Thick blood film

Once the thin film area has been produced, use the corner of the clean slide to make the thick

blood film.

Allow the blood smears to dry in a horizontal position before staining in order to obtain the

best staining quality.

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Annex 4: SOP Preparation of Giemsa Stock Solution Purpose: This SOP provides instructions for preparation of Giemsa stock stain

Principle: Light microscopy, usually applying the Giemsa staining technique, is the

established method for the laboratory diagnosis of malaria.

Giemsa is a Romanowsky stain used for staining blood films. Romanowsky stains contain

Eosin Y, an anionic acidic dye, and Azure B, a cationic basic thiazine dye obtained by oxidation

of methylene blue. When the dyes are diluted in a buffer, the anionic dye stains the acidic

components (nucleus) of cells red, and the cationic dye stains the basic components

(cytoplasm) of cells blue.

Materials and Reagents

1. Giemsa Powder

2. absolute methanol

3. Glycerol

4. Measuring cylinder

5. Glass beads

6. Funnel

7. Brown bottle

Special Safety Precaution

• Highly flammable with flash point 12 0c and Keep away from sources of ignition

• Avoid inhaling fumes and contact with skin

Procedure of Preparing Giemsa stock solution

1. Weigh the Giemsa and transfer to a dry brown bottle of 500 ml capacity which contains a

few glass beads.

2. Using a cylinder measure the methanol and add to the stain. Mix well.

3. Using the same cylinder measure, the glycerol and add to the stain. Mix well.

4. Tightly stopper the bottle.

5. Shake the bottle for 2-3 minutes.

6. Add measured glycerol and repeat the shaking.

7. Continue shaking for 2-3 minutes at 30 minutes’ intervals at least 6 times.

8. Keep the bottle for 2-3 days; shaking it 3-4 times each day.

9. Keep small amount of the stock in a small bottle

10. Label with the name of the reagent and date of preparation and mark “inflammable”.

Store at room temperature in the dark.

What you should do after preparation of Giemsa stock solution

• Keep the stopper screwed tightly

• Must be diluted with distilled water (Buffered water) with pH of 7.2.

• Should be tested for proper staining reaction

• Must be Protected from Moisture and direct sunlight

• Stored in a cool dry place in a dark bottle

• Measure a small quantity of stain into a smaller bottle for one or two days’ use

159

What you should not do after the reagent preparation

• Never add water to the stock Giemsa solution.

• Do not shake the bottle of stain before use: you will re-suspend very small, un

dissolved crystals of stain.

• Never return unused stain to the stock bottle

Quality controls of Giemsa stain

• Done to ensure the staining quality and performance of Giemsa stain

• Use known positive and negative films with each new batch of working Giemsa stain

• Blood films can be prepared using EDTA anticoagulated blood from a patient’s

• Allow the blood films to dry quickly

• Fix the films using absolute methanol

• Place them, touching back to back, in a box with separating grooves

• Label the outside of the box with the species, date and “Giemsa control slides”

• The slides can be stored at room temperature for a minimum of 1 week but will last

longer if stored at -200C or below –70 °C

• Just before use, remove the slide from the box and allow the condensation to

evaporate

• Label the slide with the date and “Positive control”

• The blood film can then be stained and examined

When do we perform IQC for Giemsa Stain?

• For newly opened/prepared Giemsa stock solution

• Regularly at least once per week

• When encountered unexpected staining result

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IQC RESULT RECORDING FORM

References

1. Cheesbrough M. District laboratory Practice in Tropical Countries. Part 1,

Cambridge University Press, UK. 1998:239-258.

2. Methods Manual for Laboratory Quality Control Testing of Malaria Rapid

Diagnostic Tests. Version Five A.

3. WHO Bench Aids for the Diagnosis of Malaria Infections. 4. WHO Basic Malaria

Microscopy, Learners Guide, 1991.

161

Annex 5: SOP Preparation of Giemsa Working Solution

Purpose: This SOP provides instructions for preparation of Giemsa working solutions from

Giemsa stock.

Principle: Light microscopy, usually applying the Giemsa staining technique, is the

established method for the laboratory diagnosis of malaria.

Giemsa is a Romanowsky stain used for staining blood films. Romanowsky stains contain

Eosin Y, an anionic acidic dye, and Azure B, a cationic basic thiazine dye obtained by oxidation

of methylene blue. When the dyes are diluted in a buffer, the anionic dye stains the acidic

components (nucleus) of cells red, and the cationic dye stains the basic components

(cytoplasm) of cells blue.

Materials and Reagents

1. Giemsa stock solution

2. Buffered Distilled water

3. Measuring cylinder 10 and 100ml capacity

4. Filter paper 5. Funnel

Special Safety Precaution

• Highly flammable with flash point 12 0c and Keep away from sources of ignition

• Avoid inhaling fumes and contact with skin

Procedure of Preparing 10% Giemsa working solution 1. Pour 90 ml of buffered water (pH 7.0

– 7.2) into the measuring cylinder. 2. Add 10 ml of filtered Giemsa stock into the measuring

cylinder 3. Mix well before using.

Procedure of Preparing 3% Giemsa working solution

1. Pour 97 ml of buffered water (pH 7.2) into the measuring cylinder.

2. Add 3 ml of filtered Giemsa stock into the measuring cylinder.

3. Mix the stain well before using.

Quality Control Check the staining quality using known QC slides for every batch of Giemsa

stain solution.

162

Annex 6: SOP for Preparation of Buffered Water

Purpose: This SOP provides instructions for preparation of buffered water (pH 7.2)

Principle: The importance of buffering the Giemsa stain solution resides in creating the

optimal PH environment for staining.

Materials and Reagents

1. Beaker, 250ml capacity

2. Graduated cylinder, 1000ml capacity

3. Buffer tablet

4. Distilled water

5. Procedure

6. Add 150ml of distilled water to beaker

7. Add one tablet

8. Shake the water until the tablets dissolve.

9. When dissolved add the fluid from the beaker to the measuring cylinder.

10. Fill the fluid in the measuring cylinder with distilled water until it is made up to 1L

mark.

Quality control Check expiry date of buffer tablet

References

1. WHO Publication: “Bench Aids for the Diagnosis of Malaria Infection”

2. WHO Basic Malaria Microscopy, Learners Guide, 2007 (revised edition)

3. Manufacturer instruction.

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Annex 7: SOP for Examination of Malaria Blood Films and Estimation of

Parasitemia

Purpose: This SOP provides instructions for the proper detection, identification and

quantification of malaria parasites in Giemsa-stained MBFs.

Principle: Examination of both thick and thin blood film is used to detect & identify malaria

parasite respectively and estimation of parasitemia.

In the thick blood film, the red blood cells (RBCs) are lyses and dehemoglobinized while the

malaria parasites are left intact and concentrated and used as a screening test to detect the

presence of malaria parasite.

In the thin blood film, when fixed with absolute methanol, enables the RBCs to retain their

original morphology with malaria parasites, if present, visible inside the RBCs, is used to

identify the species and stages of malaria parasites.

Materials, Reagents and Equipment.

Materials

1. Patient Register

2. Pen

3. Lens paper

Reagents

1. Immersion oil

2. Lens cleaning solution (80/20 Ethyl Ether solution)

Equipment’s

1. Binocular microscope

2. Tally counter(s) / Differential counter

3. Slide boxes

Procedure for Focusing and scanning blood films

1. Place the MBF on the microscope stage, switch on the light and adjust the light source

optimally by

2. looking through the ocular and the 10X/40X objectives.

3. Place a drop of immersion oil on the dry stained slide. To avoid cross contamination,

ensure that the tip of immersion oil dropper never touches the slide.

4. Slowly change to the oil immersion objective, and a thin film of oil will form between

the slide and the lens.

5. Adjust the light source optimally by looking through the 10x ocular (eyepiece) and

the100X objective and use the fine adjustment knob to focus the field; the lens should

not be allowed to touch the slide.

6. Examine the slide in a systematic fashion. Start at the left end of the thick film and

begin reading at the periphery of the field and finish at the other end. When the field is

read, move the slide right to examine adjacent fields.

164

Procedure for examining the thick blood film

1. Scan the thick film under oil immersion objective (x100) and ascertain whether a smear

is positive or negative.

2. Use the “WHO Bench Aids in the Diagnosis of Plasmodium Infections” for the

characteristics and illustrations of Plasmodium species.

3. If positive, determine all species and stages present. 4. Read a minimum of 200 oil

immersion fields before declaring the slide as negative. If time permits, scan the whole

thick film.

Procedure for examining the thin blood film

1. If the blood film is positive for malaria parasite on the thick blood film a careful

examination of the parasite morphology should continue on the thin blood film for

verification of species.

2. If different species are observed, all types should be recorded.

Procedure for Estimating Parasite density

A. Parasites/µl of blood by counting parasites against 200 WBCs in the thick film

1. Select a part of the thick film, under oil immersion objective, where the white cells are

evenly distributed and the parasites are well stained.

2. Using a piano-type tally counter (or 2 single tally counters), count parasites while

simultaneously counting WBCs in each field covered.

3. Count asexual parasites on the thick film against 200 or 500 WBCs.

4. Stop counting after counting 200 WBCs if the asexual parasites counted are greater than

150.

5. Continue counting up to 500 WBCs if parasites are less than 150 after 200 WBCS have

been counted.

6. All parasites in the final field must be counted even if a count of 200 or 500 WBCs has

been exceeded. Record actual number parasites and WBCs counted.

7. Compute for the number of parasites/µl of blood using the formula:

B. Proportion of parasitized erythrocyte / 5000 RBCs count in thin film

1. This method will indicate the percentage of erythrocytes that are infected by malaria

parasites

2. The number of parasitized erythrocyte (asexual forms) present in 25 microscopic fields

is counted divided by the total number of erythrocyte present in these fields (about

5000), and multiplied by 100.

165

Quality control

Before reading the slide, examine the thick and thin films grossly under 40 x objectives to

check the quality of the slide as follows and ensure the following:

a) Thick film is >90% intact and red cells should be completely lysed, except around the

edges. b. WBCs in the thick and thin films are properly stained (i.e., purple granules

visible within the cytoplasm of the neutrophils).

b) RBCs in the thin film do not appear pink to red.

c) Thin film has RBCs that are in one single, distinctive layer.

d) Thick or thin films have no significant debris.

If these criteria are not met, aim to collect another specimen from the patient.

Related Procedures and Documents

1. Patient Register

2. Laboratory request form

166

Annex 8: SOP for Recording and Reporting of Malaria Blood Film Results

Purpose: This SOP provides instructions for interpretation, recording and reporting of results

of MBFs.

Materials

1. Pen

2. Laboratory Request Form

3. Patient Register

Procedure of Recording of MBF Results

1. All MBFs examined, whether for routine diagnosis, referrals, confirmation or

validation, should be recorded accurately in the Patient Register.

2. MBFs for research, projects and trials should be recorded separately in study-specific

logbooks.

Procedure of Reporting of MBF Results Report all species and stages seen and if necessary

provide parasite count, according to the table below.

species stage % parasites

P.falciparum

Trophozoites, schizonts (asexual stage),

Gametocyte (sexual) stage

P.vivax

P.malarea

P.ovale

No malaria parasites seen

Example

a) P. falciparum, Trophozoites stage ;2% of RBCs are infected

b) P. vivax, Trophozoites Schizonts and gametocytes are found

c) No malaria parasites seen

References

1. WHO Bench Aids for the Diagnosis of Malaria Infections.

2. WHO Basic Malaria Microscopy, Learners Guide, 2007 (revised edition).

3. RITM, Parasitology Manual of SOPs, August 2007.

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Annex 9: Monthly Malaria Case Report Format

Region ____________________ Zone ____________________

Wereda / District _____________

Health facility ____________________________

Month ____________________ Year _________

Need separate column for persons suspected of malaria (total tested) and duplicates, and

RDT results (P. vivax or P. falciparum), number tested with both RDT and microscopy.

Total suspected malaria (total tested) = (Blood smear Pos + Blood smear negative) + (RDT

Negative +RDT positives) – (Repeat Microscopy + repeat/duplicate RDT).

168

Annex 10: SOP for Malaria Blood Film Slide Storage and Selection for

Blinded Rechecking

Purpose: This SOP provides instructions to ensure that malaria blood films (MBFs) are

properly stored and readily accessible. MBFs and their associated data records must be stored

for blinded rechecking.

Materials: For storing Malaria blood films:

1. Slide boxes

2. Tissue paper

3. cabinet

Precautions

1. Store slides by protecting from dust, direct sun light

2. Wear protective gloves when handling slides.

Procedure for Labeling and Storage of Malaria blood for External Quality Assessment

(EQA)

1. All MBFs collected for blinded rechecking must be placed in slide boxes labelled on

the outside with the short title, collection site, month and date.

2. Example of box label: Malaria EQA Program, -----HC/Hospital, 30/09/ 2003 EC

3. Store slides consecutively according to laboratory number so there is a direct link

between the results in the laboratory register and the slide location.

4. Stored slides should be free from immersion oil. Remove the oil by either gently wiping

the film with lens tissue or leaving the slides overnight with the smear side facing down

on lint free tissue paper.

5. Slides must have laboratory numbers clearly visible. Slides without laboratory numbers

cannot be used for validation purposes.

6. Results should not be written on slides; these slides cannot be used for validation

purposes

Procedure for Selection of MBFs for blinded rechecking

1. Ten stained malaria slides are selected each month to determine accuracy: 5 positive

slides and 5 negative slides.

2. If less than 10 slides are examined in the facility, select all slides for rechecking.

3. If the number of positive slides examined is less, make up the difference with negative

slides.

4. Ideally malaria slides should be stored for 1 month and the selection made before

discarding the slides. The slide selection procedure will be conducted on monthly basis

by the laboratory head/quality officer using the procedure described above (if number

of examined blood films >1000/month selection will be conducted in weekly basis)

5. Select slide from registration book and note the serial number - put a mark on the

register book to identify the selected slides.

6. During collection of selected slides, the supervisors should counter check the

conformity of the selected slides with the laboratory registration book.

7. The laboratory number and results of the selected slides from the registration book

should be recorded on the format of Annex c-1.

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Annex 11. Blinded Rechecking Result Recording and Feedback Forms

11.1. Selected Slide Result Recording Form for Rechecking

Region ___________Zone__________Woreda________Health Facility

Date sent to Rechecking Laboratory Total No. of slides

Date received at Rechecking Laboratory Total No. of slides received

Name and Initial of Receiver at Rechecking Laboratory

Slide

ID.

Diagnostic Result at the Health Facility from Laboratory

Registration Book (1st Reader)

Parasite

Density

Remark

Neg. Positive Stage of Malaria

Parasite

(for positive Slide) PV PF Mixed Others

Total

Name and signature of laboratory personnel Date

11.2. Slide Reader Result Record Form for Rechecking (2nd Reader)

Rechecking Laboratory

Region _________Zone ______________Woreda__________ Health Facility

Total slides Received Source

Name of laboratory personnel, who examine the slides

Slide

ID

2nd Reader result (At the Rechecking Lab.) Parasite

Density

Slide quality grading Remark

Neg. Positive Stage

(for

positive

Slide)

Excellent Good Poor

PV PF Mixed Others

Total

NB: -Quality of blood film includes size and thickness of the film and quality of the

staining.

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Name of 2nd reader Signature Date

General comment __________________________________________________________

11.3. Slide Reader Result Record Form for Rechecking (3rd Reader for Discordant

Result)

Rechecking Laboratory

Region __________Zone ______________Woreda __________Health Facility__________

Total slides Received Source

Name of laboratory personnel, who examine the slides

Slide ID 3rd Reader result (At the Rechecking Lab.) Parasite

Density

Slide quality grading Remark

Neg. Positive Stage

(for

positive

Slide)

Excellent Good Poor

P

V

PF Mixed Others

Total

NB: -Quality of blood film includes size and thickness of the film and quality of the

staining.

Key for Slide quality grading Excellent

Gross appearance: Both thin and thick film prepared on the same slide, thick film 10 mm diameter, newsprint read under thick film before staining, 10 mm from frosted end and thick film, 10 mm between thick and a thin film with distinct head, body and tail. Microscopic appearance: Demonstrates RBCs lysed in thick film and a monolayer of RBCs, with normal and abnormal morphology in thin film. Staining allows the trophozoites, gametocytes and/or schizonts and the white blood cells to be clearly distinguished against the background.

Good Gross appearance: Thick film with irregular and uneven thickness, thin film with uneven tail, too thick, too wide or too long. Microscopic appearance: Demonstrates RBCs lysed in thick film and a monolayer of RBCs, with normal and abnormal morphology in thin film. Staining allows the trophozoites, gametocytes and/or schizonts and the white blood cells to be clearly distinguished against the background.

Poor Gross appearance: Film with ragged tail, too thick, too wide or too long with uneven thickness. Microscopic appearance: Distorted appearance of the RBCs, malaria parasite and the white cells. Difficult to spot fields with monolayer of cells on thin film, lack of white blood cells to be clearly distinguished against the background and no properly lysed RBCs in thick film.

171

Name of 3rd reader Signature Date

General comment

11.4. Performance Notification Form

To: _____________________________________ Notification No: ___________

From: ___________________________________ Code No. _________________

I. Total No. of slides with correct reading IV. Grading of performance by % of Agreement

• Excellent (>90%)

• Very good (80-90%

• Good (70-80%)

• Poor (≤70%)

• % of false positive

• % of false negative

II. Total number of slide with discordant results

III. Type of discordance:

• # Positive diagnosed as negative

• # Negative diagnosed as positive

• # Species misdiagnosis

III) Recommendation

General

___________________________________________________________________________

Specific____________________________________________________________________

___________________________________________________________________________

Key for Slide quality grading Excellent

Gross appearance: Both thin and thick film prepared on the same slide, thick film 10 mm diameter, newsprint read under thick film before staining, 10 mm from frosted end and thick film, 10 mm between thick and a thin film with distinct head, body and tail. Microscopic appearance: Demonstrates RBCs lysed in thick film and a monolayer of RBCs, with normal and abnormal morphology in thin film. Staining allows the trophozoites, gametocytes and/or schizonts and the white blood cells to be clearly distinguished against the background.

Good Gross appearance: Thick film with irregular and uneven thickness, thin film with uneven tail, too thick, too wide or too long. Microscopic appearance: Demonstrates RBCs lysed in thick film and a monolayer of RBCs, with normal and abnormal morphology in thin film. Staining allows the trophozoites, gametocytes and/or schizonts and the white blood cells to be clearly distinguished against the background.

Poor Gross appearance: Film with ragged tail, too thick, too wide or too long with uneven thickness. Microscopic appearance: Distorted appearance of the RBCs, malaria parasite and the white cells. Difficult to spot fields with monolayer of cells on thin film, lack of white blood cells to be clearly distinguished against the background and no properly lysed RBCs in thick film.

172

Feedback Summary Table

Slide

ID.

Result Slide Quality Remark

Correctly

read

Discordant Good Poor

Pos. Report

as Neg.

Neg. Report

as Pos.

Species

Misdiagnosed

Total

References

1. Malaria Laboratory Diagnosis External Quality Assessment Scheme Guidelines, EHNRI,

2009 2. KEMRI Kisumu Malaria SOPs, 2006. 3. RITM, Parasitology Manual of SOPs, August

2007.

173

Annex 12: Exposure Reporting Form Dear colleague here is just part of our care for your wellbeing. The hospital is committed to

create a healthy working environment. We always advise universal precautions in all your

efforts in caring for others. But in case you happen to be exposed to any suspicious body fluid

that may put you at risk of HIV infection please fill this exposure reporting form and call or

get to the PEP focal person. In case you start PEP drugs please report to your prescriber if you

have any side effects in addition to your recommended (scheduled) visits.

Is the exposed person willing to be tested for HIV: □Yes □No If yes, test Result: □ HIV+□

HIV– IS the staff eligible for PEP: □ Yes □ No If eligible for PEP:

Other base line lab done and results: WBC………………………. Hgb………………….

ALT…………………………. Regimen provided: …………………………………....………

How long after exposure did the HCW start PEP medication: □ < 4 hours □ 4-24hours □

24- 72 hours

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Annex 13. List of Contributors

Ser. No Name Organization/Institute

1 Abnet Abebe EPHI

2 Teshome Degefa Jima University

3 Edosa Kifle Nekemite University

4 Ziad Amin Harari PHDRL

5 Alemayehu Belay EPHI

6 Ayalew Jejaw Gonder University

7 Siyum Obasa Nekemite RLPHDRC

8 Tsegaye Yohanes Arbamich University

9 Wondwossen Kassa EPHI

10 Seid Mohammed Afar PHDRLC

11 Gonfa Ayana EPHI

12 Tadese Menjetta Hawassa University

13 Tesfaye Kassa AA

14 Jemal Mohammed Haromaya University

15 Bedada Teshome Adama PHDRLC

16 Degaga Kenea Arsi University

17 Zerfie Tadesse APHI (Dssie)

18 Biruk Zerfu Addis Ababa University

19 Desalegn Nega EPHI

20 Getachew Tegaye Benishangul Gumuz PHDRLC

21 Gebeyaw Zeleke AA

22 Adugna Abera EPHI

23 Bisrat Nigussie EPHLA

24 Gohu Belay Hawassa PHDRLC

25 Asmare Mekonnen EPHI

26 Wondimenh Liknaw EPHI

27 Kelali Kaleaye Tigray Public Health Institute

This publication is made possible by the generous support of the American people through the United States for International Development (USAID) Transform: Health in Developing Regions.

The contents are the responsibility of the Ministry of Health-Ethiopia and do not necessary reflect the views of USAID or the United States Governmont.

USAID Transform: Health in Developing Regions is implemented by Amref Health Africa in partnership with Project HOPE, IntraHealth International and General Elctric.