Cover Image
close this bookBasic Malaria Microscopy (part I and II) (WHO - OMS, 1991, 72 p.)
View the document(introduction...)
View the documentPreface
View the documentIntroduction
View the documentLearning Unit 1. Malaria, the disease
View the documentLearning Unit 2. Cleaning and storing microscope slides
View the documentLearning Unit 3. Keeping accurate records
View the documentLearning Unit 4. Blood films
View the documentLearning Unit 5. Staining blood films with Giemsa stain
View the documentLearning Unit 6. The microscope
View the documentLearning Unit 7. Examining blood films
View the documentLearning Unit 8. Examining blood films for malaria parasites
View the documentLearning Unit 9. Artefacts in blood films
View the documentLearning Unit 10. Routine examination of blood films for malaria parasites
View the documentLearning Unit 11. Life cycle of the malaria parasite
View the documentLearning Unit 12. Supervisory aspects of malaria microscopy
View the documentBack Cover

Learning Unit 5. Staining blood films with Giemsa stain

Learning objectives

By the end of this Unit you should:

· be able to operate the simple chemical balance used in your laboratory

· be able to make up buffered water suitable for mixing with Giemsa stain, by correctly weighing the buffer salts and by the proper use of the Lovibond Comparator

· be able to make up 2% correcting fluids to adjust the pH of water for Giemsa stain

· know why it is essential for buffered water to be at pH 7.2 for good quality staining of blood films with Giemsa stain

· know when the regular method of staining or the rapid method should be used

· be able to demonstrate the use of Giemsa stain and the correctly buffered water to stain blood films suspected of containing malaria parasites, using either the regular or the rapid method

· be able to demonstrate the preparation of thick and thin blood films for staining

· know the correct and incorrect ways of handling and using Giemsa stain

· be able to demonstrate the correct drying and storage of stained slides.

Buffered water

Before you can stain blood films correctly, you first need to prepare the buffered water that is to be mixed with the stain. This buffered water needs to be at the correct pH to ensure that the quality of staining is good. Any malaria parasites that are present in blood films will then be properly stained and can be clearly seen under a microscope.

The term “pH” is used to express the acidity or alkalinity of a liquid. It is based on a scale of near 0 (very acid) to 14 (very alkaline). Liquids that are neither acid nor alkaline are called neutral and are indicated by a pH of 7.0.

The pH can be measured with a pH meter or with a colour indicator such as the Lovibond Comparator, which you will later learn how to use.

Water can be made more acid or more alkaline by the addition of certain buffer salts. These salts may be bought separately and then combined in the correct proportions, in a fixed volume of water, to give the required pH, or they may be bought as specially formulated tablets that produce a particular pH when added to a stipulated volume of water, e.g. 100 ml or 1 litre. If you plan to prepare your own buffer salts, you will need to weigh them using a balance. You must also ensure that the buffer salts have been properly stored and have not been affected by absorption of moisture from the air.

Making up buffered water


One chemical balance, readable to 0.01 g or better (a two-pan trip balance is ideal)
Two filter papers, 11 cm in diameter
One conical flask, capacity 1000 ml
One beaker, capacity 250 ml
Two wooden spatulas (wooden tongue depressors may be used)
Distilled or deionized water, 1000 ml
Potassium dihydrogen phosphate (anhydrous) (KH2PO4)
Disodium hydrogen phosphate (anhydrous) (Na2HPO4)


Step 1 Make sure that the pointer of the balance is set to zero. If it is not, adjust the balancing screw on the right-hand arm until the needle points to zero; your tutor or facilitator will help you to do this.

Step 2 Place one filter paper in each pan of the balance. Set the balance to zero again, by moving the gram weight along the gram scale arm.

Step 3 Now move the gram weight a further 0.7 g along the scale arm, ready for weighing the potassium dihydrogen phosphate

Step 4 Using a wooden spatula, take some of the potassium dihydrogen phosphate from the container and place it on the filter paper in the left-hand pan. Watch the balance pointer. Continue to add or subtract the salt until the balance pointer is again at zero.

Step 5 Transfer the 0.7 g of potassium dihydrogen phosphate to the beaker and add about 150 ml of water. Stir with the second spatula until the salt is dissolved.

Step 6 Replace the filter paper in the left-hand pan with a new one.

Step 7 After resetting the balance to zero to allow for the weight of the filter paper, adjust the gram weight to 1 g for the disodium hydrogen phosphate.

Step 8 Using the dry wooden spatula, add the disodium hydrogen phosphate to the filter paper, balancing the weight as described in step 4.

Step 9 Dissolve the disodium hydrogen phosphate in the water already in the beaker from step 5.

Step 10 When the salt is dissolved, add the fluid from the beaker to the conical flask; add water up to the neck of the flask (this makes approximately 1 litre).

The buffered water is now ready for adjusting to pH 7.2. However, before checking and adjusting the pH, you must have the correcting fluids ready for use: they will need to be made up next.

Making up the 2% correcting fluids


One chemical balance, readable to 0.01 g or better (a two-pan trip balance is ideal)
Two filter papers, 11 cm in diameter
Two glass-stoppered bottles, capacity 100 or 150 ml
Potassium dihydrogen phosphate (anhydrous) (KH2PO4)
Disodium hydrogen phosphate (anhydrous) (Na2HPO4)
Distilled or deionized water, 200 ml
Two wooden spatulas
Two beakers, capacity 250 ml
One measuring cylinder, capacity 100 ml


Step 1 Follow steps 1 and 2 of the method for making buffered water, then move the gram weight along the scale arm a further 2 g.

Step 2 Weigh 2 g of disodium hydrogen phosphate and add it to 100 ml of water in the beaker; stir with the wooden spatula until the salt has dissolved.

Step 3 Pour the solution into one of the glass bottles and label the bottle “2% disodium hydrogen phosphate”.

Step 4 Repeat steps 1 and 2 above, this time weighing out 2 g of potassium dihydrogen phosphate. Pour the solution into the second glass bottle and label it correctly.

Note: When not being used, the bottles should be stored in a cool place, away from sunlight.

Checking and adjusting the pH of the buffered water

It is very important that you check the pH of the buffered water before you use it. To alter the pH you will need to add small quantities of one of the correcting fluids: 2% Na2HPO4 if the pH is below 7.2 (too acid) or 2% KH2PO4 if the pH is above 7.2 (too alkaline). If the pH is not 7.2, it may be adjusted following the method outlined below.


One conical flask containing the buffered water
Correcting fluids (2% Na2HPO4 and 2% KH2PO4)
One Lovibond Comparator fitted with 2/1H bromothymol blue disc
Two Lovibond glass cells
One bottle of bromothymol blue indicator
One pipette, capacity 1 ml


Step 1 Pour the buffered water from the conical flask into each of the Lovibond glass cells until the 10-ml mark is reached. Place one cell in the left-hand compartment; this is the control cell.

Step 2 Pipette 0.5 ml of bromothymol blue indicator into the other cell. Mix the colour indicator and place the cell in the right-hand compartment.

Step 3 Holding the Lovibond Comparator up towards a clearly lit background, turn the colour disc until its colour matches that of the fluid in the right-hand cell.

Step 4 Adjust the pH of the water remaining in the conical flask by adding small quantities of the 2% correcting fluids. To make the water more alkaline you need to add the solution of disodium hydrogen phosphate; to make it more acid add the solution of potassium dihydrogen phosphate. Recheck pH using the Lovibond Comparator.

Staining the blood films

Regular method, for 20 or more slides


Stock of Giemsa stain
Absorbent cotton wool
Staining troughs (to hold 20 slides, placed back to back)
Distilled/deionized water, buffered to pH 7.2
Measuring cylinder, capacity 100-500 ml (depending on the number of slides to be stained)
Measuring cylinder, capacity 10-25 ml (depending on the amount of stock stain to be measured)
Flask or beaker (capacity will depend on the amount of stain to be made up)
Timing clock
Slide-drying rack

1 Methanol (methyl alcohol) is highly toxic and can cause blindness or death if swallowed. It should be stored in a lockable cupboard.


Note: For this method, it is better if slides have dried overnight.

Step 1 Fix each thin blood film by dabbing it gently with a pledget (small piece) of cotton wool dampened with methanol or by dipping it in a container of methanol for a few seconds. Avoid methanol, or its fumes, coming into contact with the thick film, otherwise fixation may take place and will prevent proper staining.

Step 2 Place the slides, back to back, in a staining trough, making sure that all thick films are at one end of the trough.

Step 3 Prepare a 3% solution of Giemsa stain by adding 3 ml of Giemsa stock solution to 97 ml of buffered water.

Step 4 Pour the stain gently into the trough until the slides are totally covered. Avoid pouring the stain directly on to the thick films.

Step 5 Leave the slides in the stain for 30-45 minutes. Experience will indicate the correct time for each batch of slides.

Step 6 Pour clean water gently into the trough to float off the iridescent “scum” on the surface of the stain. The water should be poured into the end of the trough where the thin films are, to avoid undue disturbance of the thick films.

Alternatively, gently immerse the whole trough in a bowl or basin filled with clean water.

Step 7 Gently pour off the remaining stain and rinse again in clear water for a few seconds. Then pour off the water.

Step 8 Remove the slides one by one and place them, film side downwards, in a drying rack to drain and dry, making sure that the thick film does not touch the edge of the rack.

Rapid method

The rapid staining method is generally used for between 1 and 5 slides at a time. You would use this method when you need to check urgently whether or not a patient has malaria. Much larger quantities of stain are required than for the regular method.


Giemsa stain in a 25-ml bottle
Absorbent cotton wool
Test tubes, capacity 5 ml
Distilled/deionized water, buffered to pH.7.2
Pasteur pipette, with rubber teat
Curved plastic staining tray or plate
Slide-drying rack
Timing clock
Small electric hair-drier or spirit lamp

1 Methanol (methyl alcohol) is highly toxic and can cause blindness or death if swallowed. It should be stored in a lockable cupboard.


Thick blood films must be thoroughly dry before they are stained. They can be dried more quickly with warm air blown from a small hair-drier or by exposing them to the heat from a spirit lamp. However, great care must be taken to avoid making slides hot to the touch, otherwise films will be heat-fixed and will not stain properly.

Step 1 Fix the thin film by dabbing it with a pledget of cotton wool dampened with methanol or by dipping it in a container of methanol for a few seconds. Avoid methanol, or its fumes, coming into contact with the thick film, otherwise fixation may take place and will prevent proper staining.

Step 2 Use a test-tube or small container to hold the prepared stain. Make up a 10% Giemsa solution with distilled/deionized water buffered to pH 7.2. If only one slide is to be stained, you will require about 3 ml of prepared stain. Allow 3 drops of stock Giemsa solution (from the Pasteur pipette) to each millilitre of buffered water to give a 10% solution.

Step 3 Gently pour the stain on to the slides (or use a pipette to drop the stain on to the slide).

Step 4 Stain the film for 5 to 8 minutes. Experience will indicate the correct time for each slide (or batch of slides).

Step 5 Gently flush the stain off the slide by adding drops of clean water. Never pour the stain off the slides, otherwise the surface scum will stick to the film and spoil it for microscopic examination.

Step 6 Place the slide in the drying rack, film side downwards, to drain and dry. Make sure that the thick film does not touch the edge of the rack.

Use of Giemsa stain

Giemsa stain, which is a mixture of eosin (pink-staining) and methylene blue, will be provided to you as a made-up stock solution in bottles of 100 ml, 250 ml or larger capacity.

There are a number of things that you should do, and other things you should not do, with the stock solution of Giemsa stain.

What you should do

· When the bottle of stock Giemsa is not being used, keep the stopper screwed tightly to prevent evaporation of the solvent and oxidation of the stain: the stock solution will then last longer.

· Keep the stain in a dark glass bottle and store it away from direct sunlight.

· Measure a small quantity of stain into a smaller bottle for one or two days’ use: again, the stock solution of stain will last longer.

· Store the stock solution in a cool dry place at all times.

What you should not do

· Never add water to the stock solution of stain: the smallest amount of water will cause deterioration of the solution so that it will no longer stain properly.

· Do not shake the bottle of stain before use: you will resuspend very small, undissolved crystals of stain, which can settle on the blood films during staining and obscure parts of the microscope field during examination.

· Never return unused stain to the stock bottle: it is better to measure out a small quantity for one or two days’ use.

Care of glassware

Glassware such as measuring cylinders, pipettes and staining troughs must always be clean and dry before use.

Any glassware that has been used for Giemsa stain should be rinsed in clean water immediately after use to remove as much of the stain as possible. It should then be soaked for some time, preferably overnight, in a detergent solution.

Washing glassware in detergent gives satisfactory results provided that you rinse it thoroughly in clean water. Deposits of detergent left on glassware can upset the pH of buffered water and spoil the staining, so always make sure that glassware is properly rinsed before being dried for future use.

Any stain deposits that are allowed to dry on glassware will become difficult to remove and may spoil the staining of subsequent blood films. They can be removed by soaking the glassware in methanol and then washing it with detergent in the normal way.