Basic malaria microscopy training - sucls.sd · Plasmodium falciparum -- dangerous, multi-drug resistant, causes cerebral malaria, potentially life threatening. ... Falciparum Vivax

Basic malaria microscopy

training An approach to planning and implementing training in

malaria microscopy

Omar Abdelrazig Abdalla Microbiologist – CPDC Certified Trainer

11/24/2015 Omar Abdelrazig

Omar Abdelrazig

Opening

• Learning unit 1 Malaria, the disease

• Learning unit 2 Cleaning and storing microscope slides

• Learning unit 3 Keeping records

• Learning unit 4 Preparing blood films

• Learning unit 5 Staining with Giemsa stain

• Learning unit 6 The microscope

• Learning unit 7 Examining blood films

• Learning unit 8 Examining blood films for malaria parasites

• Learning unit 9 Routine slide examination

• Learning unit 10 Supervision in malaria microscopy

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Learning objectives

• Adequately prepare a set number of thick and thin blood films;

• Demonstrate the correct practice and precautions for preventing transmission of

blood borne pathogens when handling blood;

• Demonstrate the correct Giemsa staining procedures for staining thick and thin

blood films for malaria microscopy;

• Demonstrate and describe the methods used to maintain microscopes in

working order;

• Demonstrate and use the correct procedures for examining stained thick and

thin blood films for malaria parasites;

• Demonstrate their ability to identify correctly the components of normal

blood;

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Learning objectives

• Recognize and identify malaria parasites present in blood films; identify the stage(s) of plasmodia, the presence of individual species or mixed infections of Plasmodium falciparum, P. vivax, P. malariae and P.

ovale; and establish the density of malaria parasites in the film;

• Record the results of the microscopy examination on the correct form;

• inform those responsible of the findings in a timely manner;

• Demonstrate their understanding of the requirement to observe patient

confidentiality and ethical issues;

• Follow the correct procedures of the national programme, submit reports,

stores slides for audits and prepare requests for supplies, to ensure the smooth

functioning of the microscopy facility;

• Competently evaluate the gold standard test for malaria diagnosis.

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Basic rules for the laboratory

• Do not touch, open or smell bottles, jars and containers or chemicals unless you have been instructed to do so or unless you know what you are doing and know what is in the bottle.

• Clean up when you have finished your work: • do not leave dirty glassware or slides for others to wash.

• Do not eat or drink in the laboratory: eat and drink in designated areas.

• Do not smoke.

• Use the correct precautions when handling biological specimens, chemicals and ‘sharps’, such as needles and lancets.

• Take appropriate care when handling liquids that may be corrosive or acidic or have strong fumes. Wear protective gloves when handling materials contaminated with or holding blood.

• Discard contaminated materials into the designated receptacles; if you are not sure, ask someone who will know.

• As soon as a job is finished, wash your hands with soap and water.

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Silent body communication

• Body languish

– Smiling

– Good gesture

– Standing

– Facing the clients


Nonverbal communication.

• Positive body

language

– Smiling

– Good grooming

– Erect posture

– Face to face

– Zone of comfort

Negative body languish

• looking away

• Nervous behavior

• Deep sighing

• Crossing hands

• Griming

• Masticating gum

• Yawning


Negative body language/distracting

behaviors.

• Rolling eyes

• Nervous behaviors

• Deep sighs

• Crossed arms,

• Wrinkled forehead

• Throwing things around

• Chewing gum

• Yawning

Omar Abdelrazig

Learning objectives the disease

• After completing training, trainees will have successfully acquired the skills and competence to:

• describe the importance of malaria as a potentially life-threatening disease, in which early, accurate diagnosis and treatment are essential for patient recovery and survival;

• describe four common clinical signs and symptoms of malaria in patients;

• record on the correct laboratory or survey forms relevant patient details for subsequent information and patient follow-up; demonstrate their ability to prepare slides for blood filming correctly;

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Learning objectives

• describe why malaria is an important public health problem in many parts of the world;

• describe four common symptoms of malaria;

• describe why some people have malaria parasites in their blood but have no clinical symptoms;

• explain how malaria parasites produce disease in people; explain how some species of female Anopheles mosquito transmit malaria;

• explain why accurate diagnosis of malaria depends on correct microscopic identification.

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Malaria the disease

• Importance

• Current situation

• Participants role

• Life-cycle

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How is malaria transmitted?

• By the bite of an infected female Anopheles mosquito.

• Shared dirty needles between drug addicts.

• Blood transfusion from an infected donor.

• Congenital: (between infected mother and uninfected baby through the placenta).

• By contaminated blood to unprotected persons – e.g. through skin abrasions.

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Human Malaria’s

• Plasmodium falciparum -- dangerous, multi-drug resistant, causes cerebral malaria, potentially life threatening.

• P. vivax – relapses, drug resistant, causes severe anaemia, rarely causes death.

• P. malariae – untreated can last for more than 50 years, causes kidney problems.

• P. ovale – relapses, world-wide distribution but rare, often mis-identified as P. vivax.


One child dies every 30 seconds. 3 million people die per year from malaria.

How important is this disease?


The importance of malaria

• Please remember: In the 8 hours of these lectures:

will have died from this disease or its side effects, mostly but not only in Africa,

•


What is the distribution of malaria?

• Global and country figures are often considered weak – there is much disagreement on their accuracy between groups.

• Pf the most serious – a potential killer. • Pv becoming more serious – difficulty with

relapses and increasing drug resistance. • Combined they are the greatest of the

disease burdens; greater than HIV/Aids, diarrhoea or ‘flu.

• TB is possibly a greater burden but is grossly under-diagnosed and under-reported.


This is what it usually looks like:


The possible situation in 2050


The global malaria load

• Combined there is a calculated 40% of the world’s population in 107 countries at permanent risk (80% - 90% in Africa).

• Malaria, poor education and poverty go hand in hand.

• Malaria is an opportunistic disease.

• Countries spend 1.3% of their GNP on treatment and prevention – too high for most poor nations but still not nearly enough.

• Few countries will be able to meet the GDG (UN Global Development Goals) by 2015.


Beware! Country’s that have no transmission do have malaria:

Example 1:

Oman has had no recorded transmission since 2007 but has about 85,000 cases per year among immigrant labour from Pakistan,

Bangladesh, Iran, India and Afghanistan.

Example 2:

Although Thailand has malaria transmission it is calculated that more than 50% of cases per year come over the border from Myanmar and

Cambodia for treatment.


A thin blood smear stained with Giemsa from a fatal

human case of P. knowlesi malaria.

P. knowlesi Long-tailed macaque.


Simian Malaria’s Under certain conditions some monkey malarias can affect man (zoonoses). Normally not a problem, in Malaysia and parts of SE Asia, increased contact with forested areas results in humans diagnosed with P.

knowlesi. Pk’s 24 hour blood cycle leads to hyper-parasitaemia, rapid organ failure and death (2% mortality).

Further DISCUSS.


Sporogonous Cycle:

Mosquito Stages

Gametocytes

P. falciparum

P. vivax P. ovale P. malariae

Human Liver Stages

Exo-erythrocytic

(hepatic) Cycle:

Human Blood Stages

Erythrocytic Cycle:

Malaria Lifecycle


Signs and Symptoms:

• Fever • Chills • Headache • Body pains • Convulsions • Diarrhoea • Comatose • Cold and Flu-like

symptoms • Cough • Chest pains • More ….

Lookalikes: 1.Fevers of Unknown Origin (FUO):

2.Scrub typhus (SE Asia very common)

3.Meningitis

4.Typhoid fever

5.Flu

6.Heavy cold

7.Upset stomach

8.Cholera

9.Pneumonia

10.Dengue fever

11.More ……….


Chronic malaria can result in:

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Regular low level

parasitaemia

Absence from school or work

Poor education, no wages

Ignorance, poverty,

disease, ill health

Pregnancy, anaemia, weight

loss, miscarriage or underweight birth

Lowered immunity to other diseases;

fatigue,

Neurological problems, kidney involvement – nephrotic syndrome

(p.malariae), blackwater fever. Early death

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Severe malaria rapidly leads to:

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Severe malaria

Pf Hyperparasitaem

ia

Major organ involvement

Comatose

DEATH

Multi-drug resistance

Neurological involvement

Sequestration may mask the seriousness

of the infection

Rapid RBC damage and

loss

Severe anaemia

Kidney and liver failure

DEATH

Miscarriages

Stillbirths Underweight

babies

Retinopathy

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More malaria facts, the first attack:

Falciparum Vivax Malariae Ovale

Fever lasts 16-36 hours 8-12 hours 8-10 hours 8-12 hours

First attack Severe in non-immunes

Mild to severe Mild Mild

Parasite density per micro-litre

Average 20,000-500,000

20,000 6,000 9,000

Maximum 2,000,000 50,000 20,000 30,000

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Remember: with 5,000,000 RBC’s per microlitre the percentage of red cells parasitized may range from 0.4% to 40% with Pf. More than 20% parasitized red cells has little hope for recovery and ‘exchange transfusion’ must be considered.

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Malaria in Sudan

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Overview

High perineal transmission

Irrigated malaria

Seasonal malaria

Urban malaria

Desert-fringe malaria

P. falciparum …

A. arabiensis, A. funestus and A. gambiae…

Hypo-

Meso-

Hyper-

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Malaria Burden….

-

100,000

200,000

300,000

400,000

500,000

600,000

700,000 8

4

85

86

87

88

89

90

91

92

93

94

95

96

97

98

99

00

01

02

03

Ca

se

s

Year (GC)

Twenty Years Trend of Confirmed Malaria Cases by Year and Species

POSITIVE

P.f.

P.v.


Any Questions?


Constrains in malaria microscopy

Following SOPs

Quality of supplies

personnel

Environment


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Cleaning micro-slides

• Laboratory procedures

• Steps and standards

• SOP's

• Need for quality work

• 'How to' clean microslides

• Storing microslides

• QC of activities

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Cleaning the slides Learning objectives

• describe one standard operating procedure and explain its importance for malaria microscopy;

• select from previously used slides those suitable for making blood films and demonstrate why other slides are not suitable;

• demonstrate the two correct ways of washing, drying, wrapping and storing microscope slides for making blood films.

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Cleaning the slides

• The method:

• 1. Separate new slides one from the other and soak in detergent solution for 4–8 h, conveniently overnight.

• 2. After soaking, clean each slide on both sides by rubbing the two surfaces in the washing cloth or sponge between the forefinger and thumb.

• 3. Rinse the slides individually in clean water to wash off the detergent.

• 4. Drain excess water from the slides, before placing them in the jar of alcohol with the lid firmly screwed on. Keep out of direct sunlight.

• 5. When required, remove a slide and dry it thoroughly with a clean, lint-free cotton cloth. Always handle slides by the edges.

• 6. The slide is ready for use; it does not need wrapping.

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Keeping records

• Record forms

• Need for accuracy

• Confidentiality

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Record keeping :objectives

• By the end of the module participants should be able to:

• identify the correct record form(s) and register(s) for entering information on patients;

• demonstrate preparation of accurate, error-free records on the appropriate form; select the correct copy of each record form, or completed summary, for dispatch to the supervisor;

• describe examples of the possible consequences of mixing up patients’ records; and explain why a patient’s details are confidential and must not be shared with unauthorized persons.

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Record keeping Learning objectives

• By the end you should be able to:

• follow the correct procedures of the national programme, submit reports, stores slides for audits and prepare requests for supplies, to ensure the smooth functioning of the microscopy facility;

• use the handbook as a resource to teach health workers how to make thick and thin blood films, as part of the transfer of skills and team development; and organize, following the policies and requirements of the national malaria control programme, the collaboration necessary for regular supervision of the work of the laboratory.

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Record keeping

The record should cover:

• cover: the region, province, district or zone in which the work was

done;

• the town, village or locality in which the patient lives;

• the street and house number at which the patient can be contacted;

• the patient’s name, sex and age; the patient’s number, which may also be the blood film number;

• other details, such as symptoms, body temperature and weight;

• the results of the blood film examination, such as positive or negative for malaria parasites, species and stages seen and whether P. falciparum gametocytes were observed;

• any antimalarial treatment received before microscopy examination; and other comments, observations or instructions to the clinician

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What is the expected result?

1- when not writing the full name?

2 -when touching finger while sampling?

3- when using scratched un clean slides? 4 -when drying films with oven, incubator or direct

sunlight? 5- when films exposed to dust or flies . 6- when films are left to dry horizontally? ؟

7- when methanol reach the thick film? 8- when increasing Giemsa conc and lessening staining

time? 9-when fixing or staining simples while they are wet?

10- when leaving methanol open for long time? 11/24/2015 Omar Abdelrazig

What is the expected result

when using unsuitable buffer concentration? 11- 12-when filtering the stain?

13-when discarding the stain & cleaning the slide? 14-when cleaning oculars with lenses paper of oil

immersion? 15-when alternating lenses between different

microscopes? 16-when using non conforming immersion oil?

17-when using oil mixed with xylin ؟ 18-when cleaning lenses with xylin or toluene ? 19- when cleaning microscope stage with alcohol ?


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Film preparation Learning objectives

By the end you should be able to:

• adequately prepare a set number of thick and thin blood films;

• demonstrate the correct practice and precautions for preventing transmission of blood borne pathogens when handling blood;

• demonstrate the correct Giemsa staining procedures for staining thick and thin blood films for malaria microscopy;

• demonstrate and describe the methods used to maintain microscopes in working order;

• demonstrate and use the correct procedures for examining stained thick and thin blood films for malaria parasites;

• demonstrate their ability to identify correctly the components of normal blood;

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Preparing blood film learning objectives

By the end of this unit, you will be able to:

• explain why blood must always be regarded as potentially contaminated; name four diseases found in infected blood;

• demonstrate the normal precautions used when handling blood;

• demonstrate the action to take when blood contaminates something accidentally;

• list the materials required for making thick and thin blood films; • demonstrate the correct method for preparing a thick and a thin blood

films on the same slide, for malaria microscopy;* • demonstrate the correct way of labeling a blood films; separate thick

and thin blood films of acceptable quality from unacceptable ones, giving reasons for the selection;

• describe and identify common mistakes and faults in making thick and thin blood films and the causes.

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Tools for specimen collection

You will need:

• protective quality latex gloves without talcum powder (two to three pairs per person per exercise);

• cleaned, wrapped slides (more than are needed);

• sterile lancets (one per patient, plus 10%);

• 70% ethanol; absorbent cotton wool;

• a sharps container;

• a slide box or tray for drying slides horizontally and protecting them from flies and dust; four or five clean, lint-free cotton cloths; record forms or a register;

• ballpoint ink-pen for the record forms or register;

• and an HB lead pencil to write on the thin film and small sharpener.

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Precautions preparing blood films

• Wear protective gloves when taking blood samples or handling blood.

• Avoid getting blood, including dry blood from films, on your fingers or hands.

• Cover cuts or abrasions on your hands with a waterproof dressing.

• Avoid accidentally pricking yourself when handling sharp instruments that have been in contact with blood.

• Thoroughly wash your hands with soap and water as soon as you finish a job. If you get blood on your skin, quickly wipe it off with a cotton swab dampened with alcohol; then, wash the affected area with soap and water as soon as possible.

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Collection of Blood Smears

5. Touch the drop of blood to the slide from below.

4. Slide must always be grasped by its edges.

2. Puncture at the side of the ball of the finger.

3. Gently squeeze toward the puncture site.

1. The second or third finger is usually selected and cleaned.


Preparing thick and thin films

1. Touch one drop of blood to a clean slide.

2. Spread the first drop to make a 1 cm circle.

3. Touch a fresh drop of blood to the edge of another slide.

6. Wait for both to dry before fixing and staining.

5. Pull the drop of blood across the first slide in one motion.

4. Carry the drop of blood to the first slide and hold at 45degree angle.


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Blood films

• Making blood films

• Informed consent Common faults

• Storage of films

• Transporting films

• Precautions when

handling blood

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Arrange the slides on the top of the pepper with a draw of thick

& thin blood film on it


By Using micropipette place 2ul of the blood for the thin film.


and Place 6ul of blood on the thick film.


prepare the slides for mounting.


Keep the slides of the same batch in the same slide box label it and Wright

the results .


• Arrange mixed slides with different species, stages and density with negative slides for training and education.


Kinds of blood film

Used to search for

malaria parasites.

Consists of many layers

Of red blood cells.

So… can be examined

Quickly and easily.

Used to confirm the

malaria parasite species.

Consists of a single layer

of red blood cells.

The thin film The thick film

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Interpreting Thick and Thin Films

• THICK FILM – lysed RBCs

– larger volume

– 0.25 μl blood/100 fields

– blood elements more concentrated

– good screening test

– positive or negative

– parasite density

– more difficult to diagnose species

• THIN FILM

– fixed RBCs, single layer

– smaller volume


– good species differentiation

– requires more time to read

– low density infections can be missed


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Common faults in preparing blood films

• Poorly positioned blood films

• Too much blood

• Too little blood

• Greasy slides

• Edge of the spreader slide chipped

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Malaria Blood Smear

• Prepare smears as soon as possible after collecting venous blood to avoid

• Changes in parasite morphology

• Staining characteristics

• Take care to avoid fixing the thick smear • Risk of fixing thick when thin is fixed with methanol

if both smears on same slide

• Let alcohol on finger dry to avoid fixing thick

• Be careful if drying with heat


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Staining blood films

• Buffer water

• Giemsa stain

• Staining methods

• Care of equipment

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By the end of this unit, you will be able to:

• demonstrate correct operation of the analytical balance;*

• make up the buffered water used to dilute Giemsa stain;

• demonstrate correct use of the color comparator or pH meter;*

• make up the 2% correcting fluids used to adjust the pH of buffered water;

• explain why pH 7.2 buffered water is best for good Giemsa staining;

• demonstrate two correct methods of fixing thin blood films; explain when the ‘rapid’ and ‘slow’ Giemsa staining methods are used for malaria microscopy;

• demonstrate e mastery of the rapid and slow Giemsa staining methods;

• describe the correct ways of handling and storing Giemsa stain; and

• demonstrate the correct drying and storing of stained slides. 11/24/2015

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Buffered water

• The method:

• 1. Make sure that the pointer of the balance is set at zero by adjusting the balancing screw on the right arm.

• 2. Place a filter paper in each pan; set the balance to zero, this time by moving the gram weight along the gram scale arm.

• 3. Move the gram weight a further 0.7 g along the scale arm, ready for weighing the potassium dihydrogen phosphate.

• 4. Using a wooden spatula, place some of the KH2PO4 on the filter paper in the left-hand pan

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Buffered water

• 5. Transfer the weighed KH2PO4 to the glass beaker, add about 150 ml of water, and stir with a clean spatula until the salt dissolves.

• 6. Place a fresh filter paper in the left-hand pan.

• 7. Reset the balance as before, but this time adjust the gram weight to 1 g for the Na2HPO4.

• 8. Using a clean, dry spatula, add the Na2HPO4 to the right-hand pan, balancing the weight as described in step 4 above.

• 9. Add the Na2HPO4 to the solution in the beaker and stir as in step 5.

• 10. When the salts have dissolved, add the solution to the conical flask and top up to the 1 liter mark with water.

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Buffer preparation

1buffer component :

By weighing:

) Na2HPO4 59.24 g ( + ( NaH2PO4H2O 36.38g) dissolved in 1 liter of distilled water. For daily use 10 ml of buffer is added to 990 ml of D.W.


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2% correcting fluids

• To make up the 2% correcting fluids You will need: • an analytical balance accurate to 0.01 g or better (a two-pan trip

balance is ideal, or use an electrically operated one-pan balance);

• filter papers, 11 cm in diameter;

• two glass-stoppered bottles, each of 100 or 150 ml capacity;

• potassium dihydrogen phosphate (anhydrous) (KH2PO4)

• disodium hydrogen phosphate (anhydrous) (Na2HPO4);

• distilled or deionized water, about 200 ml; O wooden spatulas; two beakers of 250 ml capacity;

• one measuring cylinder of 100 ml capacity;

• and labels.

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Needed equipments

(Forceps) stop watch Measuring cylinder . Measuring flask.

latex gloves.

Cleaning gauge. Absorbent cotton. Registry log book . Lenses paper.


Reagents

concentrated Giemsa solution

7.2 ( Buffer .(

Ethanol 70 %

Absolute methanol. Recommended oil immersion100x

(refractive index1.52)

Distilled water. Detergent soap.


Giemsa Stain

Component 500 ml

Geimsa powder 3.8 gm

methanol 250 ml

glycerin 250 ml


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The stain

• The rapid (10%) method This is the commonest method for staining 1–15 slides at a time. It is used in laboratories where a quick result to determine a patient’s malaria status is required.

The method is efficient, but more stain is used. The need for speed justifies the additional cost.

• You will need:

• Giemsa stain, decanted from the stock solution into a 25-ml bottle;

• methanol;

• 1 absorbent cotton wool;

• test tubes of 5 ml capacity;

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• distilled or deionized water buffered to pH 7.2;

• a Pasteur pipette with a rubber teat;

• a curved plastic staining tray, plate or rack;

• a slide-drying rack;

• a timing clock;

• a small electric hair-drier.

Thick blood films must be completely dry before being stained. They can be dried quickly with warm air from a small hair-drier or by careful warming over a lamp or a light bulb. Avoid overheating slides as they can ‘heat fix’ and then stain poorly.

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The method:

1. Fix the thin film by dabbing it with a pad of cotton wool dampened with methanol or by briefly dipping the film into methanol. Avoid contact between the thick film and methanol, as methanol and its vapours quickly fix the thick film, and it does not stain well.

2. Using a test tube or a small container to hold the prepared stain, make up a 10% solution of Giemsa in the buffered water by mixing three drops of Giemsa from the stock solution, using the Pasteur pipette, with 1 ml of buffered water. Each slide needs approximately 3 ml of stain to cover it.

3. Depending on whether you are using a staining tray, plate or rack, place the slides to be stained face down on the curved staining tray or face upwards on the plate or rack.

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4. Pour the stain gently under the staining tray until each slide is covered with stain, or gently pour the stain onto the slides lying face upwards on the plate or rack.

5. Stain the films for 8–10 min. Experience with the stain you are using will help you to decide the exact time needed for good staining.

6. Gently wash the stain from the slide by adding drops of clean water. Do not pour the stain directly off the slides, or the metallic-green surface scum will stick to the film, spoiling it for microscopy.

7. When the stain has been washed away, place the slides in the drying rack, film side downwards, to drain and dry. Ensure that thick films do not scrape the edge of the rack.

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• Some important things to remember with regard to the stock solution of Giemsa stain are:

• Keep the bottle tightly stoppered to avoid evaporation and oxidation of the stain by high humidity.

• Store in a dark glass bottle in a cool, dry, shady place, away from direct sunlight. For daily requirements, measure small amounts of stain into a tightly stoppered bottle (about 25 ml), so that the stock solution is less likely to be contaminated.

• Do not add water to the stock solution; even the smallest amount will cause the stain to deteriorate, making staining progressively ineffective.

• Do not shake the bottle of stain before use. Shaking re-suspends precipitates, which settle on films during staining and obscure important details during microscopy.

• Do not return unused stain to the stock bottle or to the bottle used in your daily routine. Once stain is out of the bottle, it must be used

quickly or discarded.

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The microscope

• Binoc-light microscope

• What it can do

• Use power source

• Care of microscope

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Learning objectives

By the end you should be able to:

• demonstrate the correct set-up and use of a binocular microscope with artificial and with natural light;

• demonstrate the correct use of the x10 paired oculars and x100 oil immersion objective;*

• operate the mechanical stage correctly; name correctly 10 component parts of the microscope;

• describe the correct way in which to maintain a microscope in good working order;

• describe two ways of storing a microscope correctly; and

• demonstrate the correct way of packing a microscope for long- distance transport.

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The microscope

Specifications of the microscope

• -Binocular Electrical

• - with 2 Eyepieces wide filed x 10

• - 4 objectives lens X4 , X10 , X40 , X100,

• -Voltage 230 ± 10V , 50/60 HZ. 0.25 A Fuse.

• -With blue filter

• - plastic Cover. Mechanical stage

• -lamp 6V / 20w .


Maintenance of microscpe

Clean the microscope

1\Cleaning the lens After each use

2\ Cleaning the body Of the microscope

daily

Cover the microscope

when not in use.

(dust & other )

Checking function 1/Checking Lens

2/ Checking Mechanical

parts


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Examining blood films

• Normal blood

• How to examine blood films

CCMOVBD CCMOVBD

Trophozoites

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Routine film exam Learning objectives

• demonstrate consistency in Giemsa malaria microscopy; demonstrate competence and consistent accuracy in identifying malaria parasites;

• demonstrate competence and consistent accuracy in differentiating between P. falciparum and P. vivax infections;

• explain why thick films are routinely used for malaria diagnosis and any exceptions to this rule;

• explain why parasite counts are made and their use;

• demonstrate consistency in counting malaria parasites in thick blood films and expressing them as parasites per microlitre of blood.

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Examining blood film Learning objectives

• recognize and identify malaria parasites present in blood films;

• identify the stage(s) of plasmodia, the presence of individual species or mixed infections of Plasmodium falciparum, P. vivax, P. malariae and P. ovale; and establish the density of malaria parasites in the film;

• record the results of the microscopy examination on the correct form;

• inform those responsible of the findings in a timely manner;

• demonstrate their understanding of the requirement to observe patient confidentiality and ethical issues;

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Learning objectives

• Identify the parts of a malaria parasite correctly; distinguish malaria parasites in thin and in thick films, identifying trophozoite, schizont and gametocyte stages; identify, in thin and in thick films, the four human species of malaria parasite, P. falciparum, P. vivax, P. malariae and P. ovale;

• describe and demonstrate, in thick and thin blood films, the main morphological differences between the four species of human malaria parasite;

• demonstrate common contaminants seen in blood films that are often mistaken for blood components or malaria parasites;

• recognize and name other blood parasites common to humans in your area;

• and describe ways of preventing some artifacts from contaminating blood films

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What does a malaria parasite look like?

• Where do you find it?

• What shape, or shapes, is it?

• What colour, or colours, is it?

• What size is it?


Malaria parasites are found in the red blood cells )RBC’s( of human beings:

• The RBC cycle is called the Erythrocytic cycle; it is an asexually reproductive cycle

• MP’s occupy RBC’s and grow by consuming the contents of the RBC:

- In the process malaria pigment or haematin is produced; malaria pigment does not stain but is coloured from light brown to black and can help with diagnosis


What are the components and colours?

With Giemsa or another Romanowsky stain the colours are:

Chromatin: RED

Cytoplasm: BLUE

Vacuole:

Sometimes present

Malaria pigment: BROWN to BLACK

Stippling: May be, usually PINK

RBC enlargement: May be


What shape, or shapes are they?

• MP’s can have many shapes but they all start off as a small ‘ring stage’ and progress through to ‘mature trophozoite, schizont and/or gametocyte:

• Shapes, and size, in thin films can vary from those in thick films


Naming the parts:


RING TROPHOZOITE

SCHIZONT GAMETOCYTE

Blue Cytoplasm

Red Chromatin

Brown Pigment

Recognizing Erythrocytic Stages: Schematic Morphology


Parasite shapes


Parasite shapes:


Thin and thick film appearance:


Malaria Parasite Erythrocytic Stages

Ring form

Trophozoite Schizont

Gametocytes


Plasmodium falciparum

Rings: double chromatin dots; appliqué forms; multiple infections in same red cell

Gametocytes: mature (M)and immature (I) forms (I is rarely seen in peripheral blood)

Trophozoites: compact (rarely seen in

peripheral blood)

Schizonts: 8-24 merozoites (rarely seen in peripheral blood)

Infected erythrocytes: normal size

M I


Plasmodium vivax

Trophozoites: ameboid; deforms the erythrocyte

Gametocytes: round-oval Schizonts: 12-24 merozoites

Rings

Infected erythrocytes: enlarged up to 2X; deformed; (Schüffner’s dots)


Plasmodium ovale Infected erythrocytes: moderately enlarged (11/4 X); fimbriated; oval; (Schüffner’s dots)

“malariae - like parasite in vivax - like erythrocyte”

Rings

Trophozoites: compact

Schizonts: 6-14 merozoites; dark pigment; (“rosettes”)

Gametocytes: round-oval


Infected erythrocytes: size normal to decreased (3/4X)

Plasmodium malariae

Trophozoite: compact

Trophozoite: typical band form

Schizont: 6-12 merozoites; coarse, dark pigment

Gametocyte: round; coarse, dark pigment


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,

appliqu large ring,

amoeboid

large ring,

compact

small ring,

compact

Chromatin dot often double single large 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 Thin Films

P. falciparum P. vivax P. ovale P. malariae

Rings

Trophozoites

Schizonts

Gametocytes


Parasite or Artefact?









Omar Abdelrazig

Routine film examination

• Thick films

• Thin films

• Parasite counts and densities

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Omar Abdelrazig

Calculating Parasite Density

• Using 100X oil immersion lens, select area with 10-20 WBCs/field

• Count the number of asexual parasites and white blood cells in the same fields on thick smear

• Count ≥ 200 WBCs

• Assume WBC is 8000/l (or count it)

• parasites/l = parasites count X WBC count/l

WBC counted

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Omar Abdelrazig


• Count the number of parasitized and nonparasitized red blood cells (RBCs) in the same fields on thin smear

• Count asexual stages separately from gametocytes

• Count 500-2000 RBCs (fewer RBCs if

parasitemia is high)

• % parasitemia = # parasitized RBCs X 100

total # of RBCs

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Omar Abdelrazig


• Can interconvert results in % parasitized RBCs and parasites /l if you know the RBC or WBC counts

• If unknown, can assume

4,000,000 RBCs /l or 8000 WBCs /l

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Parasitemia and clinical correlates

Parasitemia Parasites /l

Remarks

0.0001-0.0004% 5-20 Sensitivity of thick blood film

0.002% 100 Patients may have symptoms below this level, where malaria is seasonal

0.2% 10,000 Level above which immunes show symptoms

2% 100,000 Maximum parasitemia of P.v. and P.o.


Parasitemia and clinical correlates

Parasitemia Parasites/l

Remarks

2-5% 100,000-250,00

Hyperparasitemia/severe malaria*, increased mortality

10% 500,000 Exchange transfusion may be considered/ high mortality

*WHO criteria for severe malaria are parasitemia > 10,000 /l and

severe anaemia (haemaglobin < 5 g/l).

Prognosis is poor if > 20% parasites are pigment containing

trophozoites and schizonts (more mature forms) and/or if > 5% of

neutrophils contain visible pigment. 11/24/2015 Omar Abdelrazig

Omar Abdelrazig

Estimating Parasite Density Alternate Method

• Count the number of asexual parasites per high-power field (HPF) on a thick blood film

+ 1-10 parasites per 100 HPF

++ 11-100 parasites per 100 HPF

+++ 1-10 parasites per each HPF

++++ > 10 parasites per each HPF

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4,000,000 RBCs /l or 8000 WBCs /l

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Omar Abdelrazig




4,000,000 RBCs /l or 8000 WBCs /l

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Omar Abdelrazig

The gold standard diagnostic test learning objective

By the of this unit participant should be able to :

• Identify different tests used in diagnosis of malaria.

• Evaluate which test could be the gold standard one.

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Malaria Diagnosis

• Clinical Diagnosis

• Malaria Blood Smear

• Fluorescent microscopy

• Antigen Detection

• Serology

• Polymerase Chain Reaction


Clinical Diagnosis

• Hyperendemic and holoendemic areas

• Laboratory resources not needed

• Fever or history of fever

• Sensitivity ranges from poor to high

• Often has poor specificity and predictive values

• Overlap with other syndromes


Malaria Blood Smear

• Remains the gold standard for diagnosis • Giemsa stain

• distinguishes between species and life cycle stages

• parasitemia is quantifiable

• Threshold of detection • thin film: 100 parasites/l

• thick film: 5 -20 parasites/l

• Requirements: equipment, training, reagents, supervision

• Simple, inexpensive yet labor-intensive

• Accuracy depends on laboratorian skill


Interpreting Thick and Thin Films

• THICK FILM – lysed RBCs

– larger volume


– blood elements more concentrated

– good screening test

– positive or negative

– parasite density

– more difficult to diagnose species

• THIN FILM

– fixed RBCs, single layer

– smaller volume


– good species differentiation

– requires more time to read

– low density infections can be missed


Malaria Blood Smear

• Prepare smears as soon as possible after collecting venous blood to avoid

• Changes in parasite morphology

• Staining characteristics

• Take care to avoid fixing the thick smear • Risk of fixing thick when thin is fixed with methanol if

both smears on same slide

• Let alcohol on finger dry to avoid fixing thick

• Be careful if drying with heat


Fluorescent Microscopy

• Modification of light microscopy

• Fluorescent dyes detect RNA and DNA that is contained in parasites

• Nucleic material not normally in mature RBCs

• Kawamoto technique – Stain thin film with acridine orange (AO)

– Requires special equipment – fluorescent microscope

– Staining itself is cheap

– Sensitivities around 90%


Quantitative Buffy Coat (QBC ®)

• Fluorescent microscopy after centrifugation

• AO-coated capillary is filled with 50-100 µl blood

• Parasites concentrate below the granulocyte layer in tube

• May be slightly more sensitive than light microscopy but some reports of 55-84%


Quantitative Buffy Coat (QBC ®)

• Useful for screening large numbers of samples

• Quick, saves time

• Requires centrifuge, special stains

• 3 main disadvantages – Species identification and quantification

difficult

– High cost of capillaries and equipment

– Can’t store capillaries for later reference


Malaria Serology – antibody detection

• Immunologic assays to detect host response

• Antibodies to asexual parasites appear some days after invasion of RBCs and may persist for months

• Positive test indicates past infection

• Not useful for treatment decisions


Malaria Serology – antibody detection

• Valuable epidemiologic tool in some settings

• Useful for – Identifying infective donor in transfusion-transmitted

malaria

– Investigating congenital malaria, esp. if mom’s smear is negative

– Diagnosing, or ruling out, tropical splenomegaly syndrome

– Retrospective confirmation of empirically-treated non-immunes


Malaria Antigen Detection • Immunologic assays to detect specific

antigens

• Commercial kits now available as immunochromatographic rapid diagnostic tests (RDTs), used with blood

• P. falciparum histidine-rich protein 2 (PfHRP-2)

• parasite LDH (pLDH)

• Monoclonal and polyclonal antibodies used in antigen (Ag) capture test

• Species- and pan-specific Ab

• Cannot detect mixed infections

• Cross reactivity with rheumatoid factor reportedly corrected


Omar Abdelrazig

Detection of Plasmodium antigens: pLDH (parasite lactate dehydrogenase

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Omar Abdelrazig

Detection of Plasmodium antigens

• A: HRP-2 (histidine-rich protein 2) (ICT)

• B: pLDH (parasite lactate dehydrogenase)(Flow)

• C: HRP-2 (histidine-rich protein 2) (PATH)

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Polymerase Chain Reaction (PCR)

• Threshold of detection at CDC – 0.1 parasite/µl if whole blood in tube

– 2 parasites/µl if using filter paper

• Definitive species-specific diagnosis now possible

• Can identify mutations – try to correlate to drug resistance

• Parasitemia not quantifiable

• May have use in epidemiologic studies

• Requires specialized equipment, reagents, and training


Preventing Transfusion-Transmitted Malaria (TTM) Detection of Parasites/Parasite Products

Parasite densities (parasites/l)

103 10 10-1 10-3 10-5

PCR (0.05 to 0.1 parasites/l)

Microscopy (5 parasites/l)

Antigen detection (10 to 100 parasites/ l)


Omar Abdelrazig

References

• Basic malaria microscopy, learners guide. WHO.

• Centre for disease control & prevention. U.S.A.

• John Storey, WHO consultant.

• Laboratories Administration, MOH. Sudan.

• Omar Abdelrazig, MOH. Sudan.

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Omar Abdelrazig

Questions?

Questions and disscussion .

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Documents

Basic malaria microscopy training - sucls.sd · Plasmodium falciparum -- dangerous, multi-drug resistant, causes cerebral malaria, potentially life threatening. ... Falciparum Vivax