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Practical tips for field personnel

Collecting blood (Figure 17)
One essential precaution is to change pins between animals, so as not to transmit infections with the blood from one animal to another. Batches of used pins, as required, have to be sterilized (by boiling or treatment with a disinfectant). Avoid hollow needles, which are particularly dangerous for mechanical transmission of trypanosomes and other haemoparasites.

Many parasites are more concentrated in small vessels, so that low parasitaemias are easier to detect in so-called “capillary blood”. This is true not only for certain intracellular haemoparasites, such as Babesia bovis, but to some extent even for Trypanosoma congolense. The skin of the outer ear is cleaned of dirt and grease; if a liquid disinfectant is used, the skin surface should be completely dry before stabbing it with a sharp office pin, taking care to avoid visible veins; repeated stabbing may be necessary for sufficient blood to appear. One can also stab the tip of the tail, which is sometimes easier, particularly in a crush. The drop of blood is transferred to a slide (or taken up directly with the slide). If the skin is not dry, the blood will spread out on it and not make a suitable drop.

In many cases blood from a vein is satisfactory and often easier to collect. After cleaning and letting the skin dry, stab a visible vein on the outer ear with a sharp pin, or take a drop of blood from a tube in which jugular vein blood has been collected for other purposes, such as the preparation of serum. Some people prefer to take blood from the main vein under the tail, but this also depends on the way in which the animals are restrained.

Figure 17
Taking a blood sample

Figure 17

Collecting lymph
Choose a swollen lymph node, preferably the prescapular, which is the easiest to manipulate. Fix it firmly against the skin with one hand and insert with the other hand a large-bore needle, such as is used for IV injections, to a depth corresponding to the centre of the node. Some of the secrets of obtaining good lymph samples are:

  1. Avoid unnecessary movements, such as poking the needle around in the node, as this will result in bleeding and a useless sample (blood or lymph which is too diluted with blood).
  2. When inserting the needle, do not close its posterior end, so as to allow the lymph to enter the needle.
  3. Before and during the withdrawal of the needle, its rear end is closed with the thumb or by attaching a syringe, so that the lymph cannot flow back into the node. A syringe allows negative pressure to be applied, increasing the amount of lymph in the needle, but a clean syringe has to be used for each sample, as lymph is likely to enter the nozzle and contaminate the next sample.
  4. The contents of the needle are expelled on to a slide, by attaching a syringe and pushing on the plunger. A smear is made, fixed and stained, as is indicated below for blood smears.

Making blood smears
Blood smears should be even and thin, so that the red cells form one single layer. The thickness of the smear is influenced by the angle formed between the spreading slide used for drawing the blood over the second slide: the more acute the angle, the thinner the smear.

It is a good habit to make smears that are not only suitable for the diagnosis of extracellular parasites such as trypanosomes, but for intracellular organisms as well. The specific weight of white cells, as well as of red cells infected with protozoa, is slightly less than that of normal red cells and they tend to concentrate along the sides and in the tail of the smear. For the diagnosis of intracellular parasites and for establishing the white cell formula, it is therefore important that the smear does not reach the end and the edges of the slide, so that the tail and the sides can be properly examined with the microscope (Figure 18). The quantity of blood should be small, so that the tail of the smear tapers off before reaching the end of the slide, and one should cut (with sturdy scissors) one or two corners of one end of the spreading slide, so that the smear is less large than the slide on which it is made. It is also important that the edge of the spreading slide is smooth, and that the spreading movement is steady, in order to get an even smear. Another essential requirement for making good thin and even smears is that the slides are clean and free of grease. Do not use a slide twice for stained blood smears, as it is extremely difficult to clean them properly, and it is less expensive to buy new ones; one even risks finding stained parasites contained in rests of the first smear. (Such slides used for stained smears can be used after more normal cleaning for helminthological work or for examining fresh blood for trypanosomes.)

Figure 18
Preparing blood films

Figure 18

After the smear is made, it is essential to protect it from flies, which are attracted to the fresh blood, and will suck holes in the film, and may even cause pseudo-parasites to appear in the smear. Smears should also be protected from direct sun and other sources of excessive heat. The faster they are fixed and stained, the better, as old films stain badly and may even be useless, particularly in hot climates.

Staining of trypanosomes
Giemsa staining, after fixation by methanol, gives the best result, but it takes more time than more recent fast stains. This is a disadvantage for individual cases in the field, but much less when staining large numbers of smears in the laboratory.

Giemsa staining. Thin blood smears (or lymph smears) are fixed with pure43 methanol for at least two minutes (there is no maximum time limit), and then stained in Giemsa's solution, diluted 1:20 with buffered water, during some 30–60 minutes. The time needed to obtain the best results varies according to the make and sometimes even to the batch of Giemsa. (The make of Giemsa is important, some are much better than others.)

The diluted stain is made up just before use, while the smears are being fixed, by letting the concentrated Giemsa flow from a measuring pipette44 into a wide measuring cylinder holding the buffered water, and immediately swirling the cylinder slowly around to obtain an equal mixture. When adding the concentrated solution to the water, the tip of the pipette should be near the surface of the water without touching it. Avoid violent shaking and sharp shocks.

It is best to use special staining jars, in which the slides are maintained in a vertical position by grooves. There are jars for small or large numbers of slides. It is often convenient to use two such jars, one for methanol (which can be used several times, provided the methanol is protected against evaporation and against contamination) and one for the diluted Giemsa. (The interior of some staining jars can be taken out and transferred with the slides from the fixing to the staining jar.)

After fixation with methanol, the slides are transferred to the staining jar into which the diluted stain is then poured carefully until its covers the edges of the slides entirely. When preparing the stain and during the staining process, avoid shocks and contact with metal (e.g. forceps) of the diluted Giemsa, as this may induce precipitation of the stain.

After the required time for staining a flow of water is directed into the staining jar to flush out the staining solution with its covering film consisting of fine precipitate. Never allow smears to dry with staining solution adhering, as stain precipitate will cover the smear and make examination difficult or even impossible.

After washing, the slides can be put into a vertical position for drying (a wooden board with appropriate grooves is convenient). Drying may be accelerated by using an electrical hair drier, and this is particularly useful in a humid atmosphere, but avoid the use of excessive heat. Also avoid drying smears on filter paper, as this can damage the smear and moreover result in transferring cells and parasites from the smear to the paper and later to another smear. (This is not a hypothetical occurrence; because of this the author has found nucleus-containing red cells of birds in blood smears of cattle.)

The water used for the dilution of the concentrated Giemsa must be buffered, to a pH of 7.0–7.2 (7.2 giving the best result). Unbuffered tap water will not do. There are various formulas of salt mixtures which in solution will give the required pH. Salt tablets that give the pH wanted when diluted in a litre of water are commercially available and are most convenient. One can also make up one's own salt mixture, for example, a mixture of di-sodium mono-hydrogen phosphate and potassium di-hydrogen phosphate:

Na2 H PO43.0 g (anhydrous) or 7.5 g (hydrated: Na2 H PO4 - 12 H2.O)
K H2PO40.6 g
Waterone litre

The water used should be clean, if necessary filtered. Buffered water should not be kept for long periods before use and if at all it should be stored in a dark bottle, to avoid the growth of algae (which may even be the cause of pseudo-parasites in stained smears).

One can buy a concentrated solution of Giemsa ready for dilution, or make up one own's concentrated solution from powder. The latter procedure is cheaper and preferable, because the shelf-life of the solution is limited, especially in a hot climate, while the powder, if kept dry, can be kept almost indefinitely, and the concentrated solution can be made up in limited quantities, as required.

Mix 3.8 g of Giemsa powder with 250 ml of pure glycerol and 250 ml of pure methanol. Leave the mixture in a well-closed bottle containing glass beads for at least 48 hours and agitate frequently during that time. The mixture can be kept for many months. Filter through filter paper before use. Dilution with buffered water is carried out just before staining.

In some countries May-Grünwald-Giemsa staining (which includes fixation) is commonly used, but it is more expensive, does not give a better result for the staining of parasites and has the great disadvantage that there are often stain deposits on the smear, particularly in hot climates. (It does give a slightly better result for the differentiation of white blood cells, but not for blood parasites, including trypanosomes.)

43 By pure we mean laboratory quality and not contaminated with water.

44 The use of a measuring pipette is much more accurate than counting drops, the size of which depends on the orifice from which they fall.

Other staining methods
Smears of individual animals may be stained more quickly for rapid results with fast stains such as Diff-Quik, RAL 555, Field's stain or CAM's Quick-stain, after methanol fixation.

Making brain smears
Although brain smears are not commonly required for the diagnosis of trypanosomosis, they are often essential for the differential diagnosis of diseases causing central nervous symptoms, which may be due to infections by Trypanozoon, Babesia, Theileria, but also of course rabies, plant poisoning, listeriosis, etc.

Grey matter is used for smears, as it contains many capillaries. Opening the skull in order to have access to the brain is often a major undertaking in the field, and may be dangerous as brain material, potentially infected with rabies, is likely to spatter around. It is safer to take cerebellar material with a sharp spoon (curette) through the foramen magnum,45 or by making a hole in the skull with a large nail and a hammer, through which cerebral material can be taken with a large-bore needle and a syringe.

A small quantity of grey matter (corresponding to the size of one or at most two match heads) is crushed between two slides, which are pulled one across the other while pressure is maintained, resulting in two brain smears.

The smears are fixed and stained in the same way as blood smears.

45 The hole in the skull through which the spinal cord joins the medulla oblongata.

Artefacts and other pseudo-parasites

Thrombocytes in smears often mislead novices, as in the process of drying and disintegrating they may assume various shapes, and even ressemble small trypanosomes. When a thrombocyte lies across a red cell, it is sometimes taken for an intracellular haemoparasite, such as a Babesia.

Stain deposits may prevent a proper examination of the smear and are sometimes mistaken for Anaplasma or for Eperythrozoon.

Flies may deposit micro-organisms on a fresh smear, which may be mistaken for haemoparasites, and pseudo-parasites may also result from yeast or algae in the water used in the staining procedure.

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