The ideal thickness is such that on microscopy there is some overlap of red cells throughout much of the film’s length (see p. 30). The leucocytes should be easily recognizable throughout most of the film. With poorly made films the leucocytes will be unevenly distributed, with monocytes and other large leucocytes being pushed to the end and the sides of the spread. An irregular streaky film will occur if the slide is greasy, and dust on the surface will cause patchy spots (Fig. 4.1).

The films should be allowed to dry in the air. In humid conditions the films may be exposed to a current of warm air (e.g. from a hairdryer), but this should be in a microbiological safety hood.

Automated Methods

The manufacturer’s instructions should be followed unless local experience has demonstrated that variation of the recommended technique achieves better results.

Labelling Blood Films

The film should be labelled immediately after spreading. Write either a laboratory reference number or the name of the patient and the date in pencil on the frosted end of the slide or on the film itself (writing on the thickest part, which is least suitable for microscopic examination). A label written in pencil will not be removed by staining. A paper label should be affixed to the slide later. If blood films are to be stored for future reference, apply the paper label in such a manner that it is easily read when the slides are filed.

In a computerized laboratory, bar-coded specimen identification labels are convenient and preferable. These should have the patient’s name, the date and the laboratory number as well as the barcode.

Fixing Blood Films

To preserve the morphology of the cells, films must be fixed as described on p. 59. This must be done without delay, and the films should never be left unfixed for more than a few hours. If films are sent to the laboratory by post, it is essential that, when possible, they are thoroughly dried and fixed before dispatch.

Bone Marrow Films

The method for preparation of films of aspirated bone marrow is described on p. 126. They should be made without delay. Films must be thoroughly dry before they are fixed or artefactual changes will occur. At least one film should be fixed for a Perls’ stain on the initial bone marrow aspirate of each patient, and, if necessary, films should be fixed in the appropriate fixatives for special staining (Chapter 15); others should be fixed and stained with a Romanowsky stain as described later. Crushed bone marrow particles and touch preparations from trephine biopsy specimens can be stained in the same manner.

Staining blood and bone marrow films

Romanowsky stains are used universally for routine staining of blood films, and satisfactory results can be obtained. The remarkable property of the Romanowsky dyes of making subtle distinctions in shades of staining, and of staining granules differentially, depends on two components: azure B (trimethylthionin) and eosin Y (tetrabromo-fluorescein).^1,²

The original Romanowsky combination was polychrome methylene blue and eosin. Several of the stains now used routinely that are based on azure B also include methylene blue, but the need for this is debatable. Its presence in the stain is thought by some to enhance the staining of nucleoli and polychromatic red cells; in its absence, normal neutrophil granules tend to stain heavily and may resemble ‘toxic granules’ in conventionally stained films.³

There are a number of causes of variation in staining. One of the main factors is the presence of contaminants in the commercial dyes and a simple combination of pure azure B and eosin Y might be considered preferable to the more complex stains because this ensures consistent results from batch to batch.^1,^4,⁵ However, in practice, absolutely pure dyes are expensive, and it is sufficient to ensure that the stains contain at least 80% of the appropriate dye.⁶ Among the Romanowsky stains now in use, Jenner is the simplest and Giemsa is the most complex. Leishman’s stain, which occupies an intermediate position, is still widely used in the routine staining of blood films, although the results are inferior to those obtained by the combined May–Grünwald–Giemsa, Jenner–Giemsa, and azure B–eosin Y methods. Wright’s stain, which is widely used in North America, gives results that are similar to those obtained with Leishman’s stain, whereas Wright–Giemsa is similar to May–Grünwald–Giemsa.

A pH to the alkaline side of neutrality accentuates the azure component at the expense of the eosin and vice versa. A pH of 6.8 is usually recommended for general use, but to some extent this depends on personal preference. (When looking for malaria parasites, a pH of 7.2 is recommended to see Schüffner’s dots.) To achieve a uniform pH, 50 ml of 66 mmol/l Sörensen’s phosphate buffer (see p. 622) may be added to each litre of the water used in diluting the stains and washing the films.

The mechanism by which certain components of a cell’s structure stain with particular dyes and other components fail to do so depends on complex differences in binding of the dyes to chemical structures and interactions between the dye molecules.⁷ Azure B is bound to anionic molecules, and eosin Y is bound to cationic sites on proteins.

Thus, the acidic groupings of the nucleic acids and proteins of the cell nuclei and cytoplasm of primitive cells determine their uptake of the basic dye azure B, and, conversely, the presence of basic groupings on the haemoglobin molecule results in its affinity for acidic dyes and its staining by eosin. The granules in the cytoplasm of neutrophil leucocytes are weakly stained by the azure complexes. Eosinophilic granules contain a spermine derivative with an alkaline grouping that stains strongly with the acidic component of the dye, whereas basophilic granules contain heparin, which has an affinity for the basic component of the dye. These effects depend on molar equilibrium between the two dyes in time-dependent reactions.² DNA binds rapidly, RNA more slowly, and haemoglobin more slowly still; hence the need to have the correct azure B to eosin ratio to avoid contamination of the dyes and to stain for the right time. Standardized stains and staining method have been proposed (see p. 61).

The colour reactions of the Romanowsky effect are shown in Table 4.1; causes of variation in staining are given in Table 4.2.

Table 4.1 Colour responses of blood cells to Romanowsky staining

Cellular component	Colour
Nuclei
Chromatin	Purple
Nucleoli	Light blue
Cytoplasm
Erythroblast	Dark blue
Erythrocyte	Dark pink
Reticulocyte	Grey-blue
Lymphocyte	Blue
Metamyelocyte	Pink
Monocyte	Grey-blue
Myelocyte	Pink
Neutrophil	Pink/orange
Promyelocyte	Blue
Basophil	Blue
Granules
Promyelocyte (primary granules)	Red or purple
Basophil	Purple black
Eosinophil	Red-orange
Neutrophil	Purple
Toxic granules	Dark blue
Platelet	Purple
Other inclusions
Auer body	Purple
Cabot ring	Purple
Howell–Jolly body	Purple
Döhle body	Light blue

Table 4.2 Factors giving rise to faulty staining

Appearances	Causes
Too blue	Incorrect preparation of stock, eosin concentration too low
	Stock stain exposed to bright daylight
	Batch of stain solution overused
	Impure dyes
	Staining time too short
	Staining solution too acid
	Film too thick
	Inadequate time in buffer solution
Too pink	Incorrect proportion of azure B:eosin Y
	Impure dyes
	Buffer pH too low
	Excessive washing in buffer solution
Pale staining	Old staining solution
	Overused staining solution
	Incorrect preparation of stock
	Impure dyes, especially azure A and/or C
	High ambient temperature
Neutrophil granules not stained	Insufficient azure B
Neutrophil granules dark blue/black (pseudo-toxic)	Excess azure B
Other stain anomalies	Various contaminating dyes and metal salts
Stain deposit on film	Stain solution left in uncovered jar
Stain deposit on film	Stain solution not filtered
Blue background	Inadequate fixation or prolonged storage before fixation
Blue background	Blood collected into heparin as anticoagulant

Preparation of Solutions of Romanowsky Dyes

Buffered Water

Make up 50 ml of 66 mmol/l Sörensen’s phosphate buffer of the required pH to 1 litre with water at a pH of 6.8 (see p. 622). An alternative buffer may be prepared from buffer tablets, which are available commercially. Solutions of the required pH are obtained by dissolving the tablets in water.

Staining methods

May–Grünwald–Giemsa Stain

Dry the films in the air, then fix by immersing in a jar of methanol for 5–10 min. For bone marrow films, allow a longer time to ensure thorough drying and then leave it for 15–20 min in the methanol. Films should be fixed as soon as possible after they have dried. If they are left unfixed at room temperature it may be found that the background of dried plasma stains a pale blue that is impossible to remove without spoiling the staining of the blood cells. It is important to prevent any contact with water before fixation is complete. Methyl alcohol (methanol) is the fixative of choice, although ethyl alcohol (‘absolute alcohol’) can also be used. To prevent the alcohol from becoming contaminated with absorbed water, it must be stored in a bottle with a tightly fitting stopper and not left exposed to the atmosphere, especially in humid climates. Methylated spirits must not be used because it contains water.

Transfer the fixed films to a staining jar containing May–Grünwald stain freshly diluted with an equal volume of buffered water. After the films have been allowed to stain for about 15 min, transfer them without washing to a jar containing Giemsa’s stain freshly diluted with 9 volumes of buffered water, pH 6.8.

After staining for 10–15 min, transfer the slides to a jar containing buffered water, pH 6.8, rapidly wash in three or four changes of water, and finally allow to stand undisturbed in water for a short time (usually 2–5 min) for differentiation to take place. This may be controlled by inspection of the wet slide under the low power of the microscope; with experience, the naked-eye colour of the film is often a good guide. The slides should be transferred from one staining solution to the other without being allowed to dry. Because the intensity of the staining is affected by any variation in the thickness of a film, it is not easy to obtain uniform staining throughout a film’s length.

When differentiation is complete, stand the slides upright to dry. This method is designed for staining a number of films at the same time. Single slides may be stained by flooding the slide with a combined fixative and staining solution (e.g. Leishman’s stain, discussed later), but it is important to ensure that the methanol used as fixative is completely water-free. As little as 1% water may affect the appearance of the films, and a higher water content causes gross changes (Fig. 4.2). The red cells will also be affected by traces of detergent on inadequately washed slides (see Fig. 26.5, p. 612).

Figure 4.2 Blood film appearances following methanol fixation. Photomicrographs of Romanowsky-stained blood films that have been fixed in methanol containing: (A) 1% water; (B) 3% water; (C) 4% water; and (D) 10% water. The red cells and leucocytes are well fixed in (A), reasonably well fixed in (B) but badly fixed in (C) and (D).

The diluted stains usually retain their staining powers sufficiently well for several batches of slides to be stained in them. They must be made up freshly each day, and it is probably best to stain the day’s films in two batches, morning and afternoon. There is no need to filter the stains before use unless a deposit is present.

Standardized Romanowsky Stain

A standardized Romanowsky stain ^2,⁵ based on a method with pure dyes has been proposed by the International Committee for Standardization in Haematology. It is useful for checking the performance of routine stains. The method is described fully in previous editions.

Jenner–Giemsa Stain

Jenner’s stain may be substituted for May–Grünwald stain in the technique described on the previous page. The results are a little less satisfactory. The stain is used with 4 volumes of buffered water and the films, after being fixed in methanol, are immersed in it for approximately 4 min before being transferred to the Giemsa stain. They should be allowed to stain in the latter solution for 7–10 min. Differentiation is carried out as described earlier.

Leishman’s Stain

Air dry the film and flood the slide with the stain. After 2 min, add double the volume of water and stain the film for 5–7 min. Then wash it in a stream of buffered water until it has acquired a pinkish tinge (up to 2 min). After the back of the slide has been wiped clean, set it upright to dry.

Automated Staining

Automatic staining machines are available that enable large batches of slides to be handled. They may be either stand-alone staining machines or a part of a large automated blood counting instrument. In many instances, the instrument spreads, fixes and stains blood films. Some automated instruments incorporating staining can only be programmed to prepare and stain a single film per sample. Others can prepare and stain multiple films from a single blood sample; this is useful for preparing slides for teaching large numbers of students. Some systems apply staining solutions to slides lying horizontally (flat-bed staining), whereas others either immerse a slide or slides in a bath of staining solution (‘dip-and-dunk’ technique) or spray stain onto slides in a cytocentrifuge. Problems include increased background staining, inadequate staining of neutrophil granules, degranulation of basophils and blue or green rather than pink staining of erythrocytes. These problems are usually related to the specific stains and staining protocols used rather than to the type of instrument, although flat-bed stainers are more likely to cause problems with stain deposit. However, as a rule, staining is satisfactory provided that reliable stains are used and there is careful control of the cycle time and other variables.⁸ Flat-bed stainers may not stain an entire film (e.g. a bone marrow film) if the film exceeds the standard length.

Rapid Staining Method

Field’s method ^9,¹⁰ was introduced to provide a quick method for staining thick films for malaria parasites (see below). With some modifications, it can be used fairly satisfactorily for the rapid staining of thin films. The stains are available commercially ready for use, or they can be prepared as follows.

Stains

Stain A (Polychromed Methylene Blue)

Methylene blue	1.3 g
Disodium hydrogen phosphate (Na₂HPO₄.12H₂O)	12.6 g
Potassium dihydrogen phosphate (KH₂PO₄)	6.25 g
Water	500 ml

Dissolve the methylene blue and the disodium hydrogen phosphate in 50 ml of water. Then boil the solution in a waterbath almost to dryness to ‘polychrome’ the dye. Add the potassium dihydrogen phosphate and 500 ml of freshly boiled water. After stirring to dissolve the stain, set aside the solution for 24 h before filtering. Filter again before use. The pH is 6.6–6.8.

Alternatively, azure B may be added to the methylene blue in the proportion of 0.5 g of azure B to 0.8 g of methylene blue, and the combined dyes are then dissolved directly in the phosphate buffer solution.

Stain B (Eosin)

Eosin	1.3 g
Disodium hydrogen phosphate (Na₂HPO₄.12H₂O)	12.6 g
Potassium dihydrogen phosphate (KH₂PO₄)	6.25 g
Water	500 ml

Dissolve the phosphates in warm freshly boiled water and then add the dye. Filter the solution after letting it stand for 24 h.

Method

Fix the film for 10–15 s in methanol. Pour off the methanol and drop on the slide 12 drops of diluted Stain B (1 volume of stain to 4 volumes of water). Immediately, add 12 drops of Stain A. Agitate the slide to mix the stains. After 1 min, rinse the slide in water, then differentiate the film for 5 s in phosphate buffer at pH 6.6, wash the slide in water, and then place it on end to drain and dry. Two-stage stains of this type are also available commercially.

Mounting of Coverglass

When thoroughly dry, cover the blood film with a rectangular No. 1 coverglass, using for this purpose a mountant that is miscible with xylol (e.g. DPX Mountant, Merck). For a temporary mount, cedarwood oil may be used.

The coverglass should be large enough to overlie the whole film, so that the edges and the tail of the film can be examined. If a neutral mounting medium is used, the staining should be preserved for many years if kept in the dark. Although it is probable that stained films keep best unmounted, there are objections to this course: it is almost impossible to keep the slides free from dust and from being scratched and, in the absence of a coverglass, the observer is tempted to examine the film solely with the oil-immersion objective, a practice that is to be deprecated because it is important to have a general overview of the film before studying specific cells.

Examination of wet blood film preparations

The examination of a drop of blood sealed between a slide and coverglass is sometimes of considerable value. The preparation may be examined in several ways: by ordinary illumination, by dark-ground or by Nomarski (interference) illumination. Chemically clean slides and coverglasses (see p. 623) should be used,* and the blood should be allowed to spread out thinly between them. If the glass surfaces are free from dust, the blood will spread out spontaneously, and pressure, which is undesirable, should not be necessary. The edges of the preparation may be sealed with a melted mixture of equal parts of petroleum jelly and paraffin wax or with nail varnish.

Red Cells

Rouleaux formation is typically seen in varying degrees in wet preparations of whole blood and must be distinguished from autoagglutination. The distinction is sometimes a matter of considerable difficulty, particularly when, as not infrequently happens, rouleaux formation is superimposed on agglutination. The rouleaux, too, may be notably irregular in haemolytic anaemias characterized by spherocytosis, whereas the clumping caused by massive rouleaux formation of normal type may closely simulate true agglutination. This pseudoagglutination owing to massive rouleaux formation may be distinguished from true agglutination in two ways:

1. By noting that the red cells, although forming parts of larger clumps, are mostly arranged side by side as in typical rouleaux.

2. By adding 3–4 volumes of 9 g/l NaCl to the preparation. Pseudoagglutination owing to massive rouleaux formation should either disperse completely or transform itself into typical rouleaux. The addition of saline to blood that has undergone true agglutination may cause the agglutinates to break up somewhat, but a major degree of it is likely to persist and typical rouleaux will not be seen.

Anisocytosis and poikilocytosis can be recognized in ‘wet’ preparations of blood, but the tendency to crenation and the formation of rouleaux tend to make observations on shape changes rather difficult. Such changes can best be studied in a wet preparation after fixation. For this, dilute a sample of freshly collected heparinized or EDTA-anticoagulated blood in 10 volumes of iso-osmotic phosphate buffer, pH 7.4 (see p. 622) and immediately fix with an equal volume of 0.3% glutaraldehyde in iso-osmotic phosphate buffer, pH 7.4. After standing for 5 min, add 1 drop of this suspension to 4 drops of glycerol and place 1–2 drops on a glass slide that is then sealed.¹¹

Pitting occurs normally in less than 2% of the red cells; an increase of more than 4% is an indication of splenic dysfunction. The pits are readily identified by Nomarski illumination or electron microscopy when they have the appearance of small crater-like indentations on the cell surface.¹²

Sickling of red cells in ‘wet’ preparations of blood is described in Chapter 14.

Crystals of haemoglobin C can be demonstrated by incubating a sample of blood with an equal volume of 30 g/l sodium chloride for 4 h at 37°C.¹³ In blood from patients with haemoglobin C disease this induces formation of intracellular haemoglobin C crystals, large, clear structures that are well shown when the preparation is then stained by any Romanowsky stain.¹³ They can also be demonstrated in red cells from patients with compound heterozygosity for haemoglobins S and C.

Cryoglobulinaemia

To identify cryoglobulinaemia, put a drop of blood from an EDTA sample that has been kept at room temperature onto a glass slide, cover it with a glass coverslip, and examine it by phase contrast microscopy. The cryoglobulin will be seen as large clear deposits of amorphous material or as refringent precipitates that disappear when the slide is warmed to 37°C.

This is a useful test when an automated blood count gives anomalous results with spuriously elevated white blood cell and platelet counts.¹⁴

Leucocytes

The motility of leucocytes can be readily studied in heparinized blood if the microscope stage can be warmed to about 37°C. Usually, only the granulocytes show significant progressive movements. However, the examination of living neutrophils in plasma is not useful in day-to-day routine haematological practice. Specialized microscopy techniques applicable to leucocytes are discussed in the 8th edition of this book.

Separation and concentration of blood cells

A number of methods are available for the concentration of leucocytes or abnormal cells when they are present in only small numbers in the peripheral blood. Concentrates are most simply prepared from the buffy coat of centrifuged blood. However, most methods affect to some extent subsequent staining properties, chemical reactions and viability of the separated cells.

Making a Buffy Coat Preparation

Centrifuge an EDTA blood sample in a plastic tube for 5–10 min at 1200–1500 g. Then remove the supernatant plasma carefully with a fine plastic pipette and with the same pipette deposit the platelet and underlying leucocyte layers onto one or two slides. Mix the buffy coat in a drop of the patient’s plasma and then spread the films. Allow them to dry in the air and then fix and stain in the usual way.

When leucocytes are scanty or if many slides are to be made, it is worthwhile centrifuging the blood twice; first, about 5 ml are centrifuged and a second tube is then filled from the upper cell layers of this sample.

As an alternative to centrifugation, the blood may be allowed to sediment by placing the tube vertically on the bench without disturbance, with or without the help of sedimentation-enhancing agents such as fibrinogen, dextran, gum acacia, Ficoll (Pharmacia) or methylcellulose.¹⁵ Bøyum’s reagent¹⁶ (methylcellulose and sodium metrizoate) is particularly suitable for obtaining leucocyte preparations with minimal red cell contamination.

Utility of the Buffy Coat

It is well known that atypical or primitive blood cells circulate in small numbers in the peripheral blood in health. Thus, atypical mononuclear cells, metamyelocytes and megakaryocytes may be found. Even promyelocytes, blasts and nucleated red cells may occasionally be seen but only in very small numbers. Efrati and Rozenszajn ¹⁷ described a method for the quantitative assessment of the numbers of atypical cells in normal blood and gave figures for the incidence of megakaryocyte fragments (e.g. mean 21.8 per 1 ml of blood) and of atypical mononuclear cells and metamyelocytes and myelocytes. In cord blood, the incidence of all types of primitive cells is considerably greater.¹⁸

In disease, abnormal cells may be seen in buffy coat preparations in much larger numbers than in films of whole blood (Fig. 4.3). Another example, for instance, is megakaryocytes and immature cells of the granulocyte series found in relatively large numbers in disseminated carcinoma.¹⁹ Megaloblasts, if present, may help in the diagnosis of a megaloblastic anaemia. Ring sideroblasts may be seen in patients with sideroblastic anaemia; their presence can be confirmed with a Perls’ stain. Haemophagocytosis, which is more often observed in the bone marrow, may also sometimes be demonstrated in buffy coat preparations.²⁰ Erythrophagocytosis may be conspicuous in cases of autoimmune haemolytic anaemia (Fig. 4.4). In systemic lupus erythematosus (SLE) a few LE cells may be found, but this is not the best way to demonstrate LE cells; moreover, the detection of LE cells for the diagnosis of SLE has been supplanted by immunological tests for the detection of antinuclear or anti-DNA antibodies.

Figure 4.3 Buffy coat film. From blood of a patient with pancytopenia showing two hairy cells, an erythroblast and a bare nucleus. Only rare cells resembling hairy cells were seen in the conventional film, whereas the buffy coat film showed many such cells.

Figure 4.4 Film of a buffy coat. Erythrophagocytosis in autoimmune haemolytic anaemia.

Buffy coat films can be useful for the detection of bacteria, fungi or parasites within neutrophils, monocytes or circulating macrophages; they also can help find cells that may be present in very small numbers (e.g. hairy cells in hairy cell leukaemia). With the availability of monoclonal antibodies reactive with epithelial and other tumour cells, immunocytochemical techniques can now be applied for the identification of infrequent neoplastic cells.

Separation of Specific Cell Populations

It is now possible to identify specific cell populations by flow cytometric immunophenotyping, and the need for separation of mononuclear cells from blood has diminished. However, differences in density of cells can also be used to separate individual cell types, using gradient solutions of selected specific gravity.^16,²¹ This is also a useful method for use in leucocyte imaging with radio-isotope-labelled neutrophils (see p. 388). A simple convenient technique has been described for layering the blood or bone marrow over the density preparations.²² The median density values for the main haemopoietic cells are as follows:

Erythrocytes	1100
Eosinophils	1090
Neutrophils	1085
Myelocytes	1075
Lymphocytes	1070
Monocytes	1064
Myeloblasts	1062
Platelets	1035

Bacteria and fungi detectable in blood films

Ehrlichiosis is a tick-borne fever in which clusters of small organisms may be seen in Romanowsky-stained blood smears. The detection of organisms within neutrophils or monocytes is important for its diagnosis. Other bacteria and fungi are occasionally detected within neutrophils or monocytes or extracellularly.

Parasites detectable in blood, bone marrow or splenic aspirates

There are now a number of screening tests for diagnosing malaria based on the detection of malarial antigens (see Chapter 6). However, the essential method for a definitive diagnosis remains the finding of parasites in a blood film and the identification of the species by morphology.^23,²⁴ Only brief outlines of the microscopic diagnoses are given in this chapter. For more detailed accounts, readers are referred to a parasitology textbook. In addition to the plasmodia that give rise to malaria, the other important parasites to be found in the blood are leishmaniae, babesiae, trypanosomes and microfilaria.

In addition to standard thin films, thick films are extremely useful when parasites are scanty. These should be prepared and examined as a routine, although identification of the species is less easy than in thin films and mixed infections may be missed. If 5 min are spent examining a thick film, this is equivalent to about 1 h spent in traversing a thin film. Once the presence of parasites has been confirmed, a thin film should be used for determining the species and, in the case of Plasmodium falciparum, for assessing the severity of the infection by counting the percentage of parasitized cells (excluding cells containing only gametocytes).

Low levels of parasitaemia detected by immunological tests may be missed by microscopy and proficiency testing studies have demonstrated the need for all laboratories, and especially those lacking expertise, to take part in external quality control programmes and to refer problematic cases to more experienced centres.^25,²⁶

Thick blood films are also useful for the detection of microfilaria. When they are used for this purpose, it is important to scan the entire film using a low-power objective, or parasites may be missed. Examination of wet preparations of blood can be used for diagnosis of microfilariae and has the advantage that the parasites are easily detected because they are moving. A stained film is necessary for confirmation of species. Wet preparations are also useful for the detection of trypanosomes and the spirochaetes of relapsing fever. The presence of small numbers of trypanosomes or spirochaetes is revealed by occasional slight agitation of groups of red cells. Examination of a stained film confirms their nature.

Examination of blood films for parasites

Making Thick Films

Make a thick film by placing a small drop of blood in the centre of a slide and spreading it out with a corner of another slide to cover an area about four times its original area. The correct thickness for a satisfactory film will have been achieved if, with the slide placed on a piece of newspaper, small print is just visible.

Allow the film to dry thoroughly for at least 30 min at 37°C. If it is necessary to hurry the procedure, the slide can be left near, but not touching, a light bulb where the temperature is 50–60°C, for about 7 min; the quality of the film may deteriorate if it is overheated. Films which are not completely dry may wash off in the stain.

Staining Thick Films

Field’s method of staining ^9,¹⁰ is quick and usually satisfactory for thick films, but the method is not practical for staining large numbers of films; for this purpose the Giemsa, Leishman or azure B–eosin Y methods are more suitable. Careful attention to pH is critical for satisfactory staining of parasites.