The Laboratory Diagnosis of Malaria - Semantic Scholar

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THE LABORATORY DIAGNOSIS OF MALARIA.* jy

By R. KNOWLES,

LIEUTENANT-COLONEL, I.M.S.,

Professor of Protozoology, Calcutta School of Tropical Medicine.

When the Secretary asked me to prepare something for this meeting, I offered him the choice between a

paper on the historical aspects of the treatment of

malaria, and a clinical demonstration on laboratory methods in the diagnosis of malaria. He chose the latter. The subject may seem a most elementary one, but almost every day of the week I receive letters from all over India asking for information with regard to it.

Correspondents write, " I cannot get Leishman's stain to work in the mofussil. Why is this? " Or,

" Please give me details of the method of diagnosis of malaria by culture." Or,

" Must the water used in Romanowsky staining be distilled; will not tap water do?" Now I think that any surgeon will admit that his success in surgery depends upon the most meticulous attention to details of technique, and the same is true of all laboratory procedures.

In order not to take up too much of your time, Dr. B. M. Das Gupta has kindly consented to demons- trate the various procedures concerned, as I describe them; whilst in my department next door we have got out a demonstration showing the complete life cycle in man of Plasmodium falciparum as seen in Bass' culture. The latter illustrates such points as that the early gametocytes of this species are spherical bodies, and that they only become crescentic in shape as they reach maturity and pass into the peripheral circulation; the very marked development of black hsemozoin in all developing forms?usually in the form of a con-

densed eccentric cluster; and the mature schizont rosettes of this species, which fill the whole of the erythrocyte and contain approximately 22 merozoites. There are four procedures here to be considered, and

they are:? (i) The preparation, staining and examination of thin

blood films. {ii) The preparation, staining and examination of

thick blood films. (Hi) The use of thin and thick blood films on the

same slide. (iv) The diagnosis of malaria by cultural methods. Before proceeding further, let me emphasise the

utmost importance of using only perfectly clean and well polished slides in making films. Frosted slides, scratched slides?such as medical storekeepers delight to supply?are useless. If you entrust the cleaning of old used slides to the laboratory sweeper, he will sit down on the floor with a bowl of water and a supply of saji mutti (crude carbonate of soda), and scrub the slides clean. This ruins their surface, as it scratches them; it should never be permitted. The simplest way of

cleaning old slides is to boil them for some hours in an emulsion of country soap or soft soap in water; they must then be very thoroughly washed in running tap water for some hours to remove all trace of alkali; then stored in rectified spirit in a glass jar with a ground glass stoppered lid. Before use they are removed from the spirit, polished with a clean, old, much washed cotton handkerchief, and four or five are laid flat on a clean piece of paper by the patient's bedside.

Thin Blood Films.

It is important to select the spreader with care. The best spreader is a glass slide with a smooth, even end,

* Being an address delivered to the Calcutta Branch of the British Medical Association on Friday, the 13th February, 1931.

272 THE INDIAN MEDICAL GAZETTE. [May, 1931.

as tested by running it across the ball of the thumb. Or a hypodermic needle may be used instead. In using the spreader do not pick it up by the ends, as this may deposit dirt from the operator's fingers on the spreading edge, but hold it by the sides. As a pricker, I always use a clean, sterilised hypodermic needle; or

a surgical needle may be used, or Wright's glass pricker, made by drawing out in the flame a capillary glass tube and breaking off the end. It is not usually necessary to mop up the patient's skin with alcohol and iodine, but it is necessary to flame the needle. Boyd (1930) in his Introduction to Malariology says that it is better to take the blood from the lobe of the patient's ear, as patients may sometimes faint when the finger is

pricked; all that I can say is that in 26 years' experi- ence of pricking patients' fingers, I have never known a patient to faint. Do not prick the pulp of the finger, as this is painful, or the lateral aspect by the side of the nail. Select instead the dorsal aspect just below the root of the nail. If, as often happens in malaria, the patient is perspiring profusely, it may be necessary to cleanse and dry this part with a cotton-wool swab soaked in spirit. If the patient is very amende, make him forcibly flex the fingers on to the palm of the

hand; this will ensure getting a good supply of blood. In extremely anaemic subjects it may be necessary to

apply a tourniquet, especially if thick films are also wanted. Thin rubber pressure tubing is the best form of tourniquet to apply. Having pricked the finger with a short deep stab, take

the clean slides from the paper one by one, invert them, and bring the slide just into contact with the issuing blood. This gives a small?not large?drop of blood about half an inch from the end of the slide. Apply the spreader at an angle of 45 degrees; wait until the blood runs by capillary attraction between the spreader and the slide, and then draw the film. It is very important that the blood should follow the spreader, and that the spreader shall not push the blood in front of it (vide Fig. 1).

If the blood is pushed in front of the spreader, the spreader passes over the cells and greatly distorts the parasites. Even in a properly drawn film the very delicate hair-like rings of P. falciparum are liable to considerable distortion, whilst the delicate cytoplasmic bridges in the growing trophozoites of P. vivax may be broken, this giving rise to the appearance of detached blue staining masses of parasite cytoplasm within the erythrocyte. The cigarette paper method of preparing films

recommended in some books is not advisable; it may fill the film with paper fibres and other foreign bodies. Four further points about the thin film. In extreme

dry heat, e.g., in Iraq in the middle of the hot weather with atmospheric temperatures going up to 125?F., films taken in the middle of the day are apt to show considerable distortion of the parasites, and render the

identification of species difficult; under such conditions it is best to take the films after sunset or in the early

morning, except in cases of emergency. In intense wet

heat, such as prevails in the Vizagapatam Agency Tracts in the rains, the instant that the film has been drawn it must be held over the flame of a spirit lamp t?

fix it; if this is not done the film becomes hsemolysed, thus rendering the identification of parasites difficult. In malaria survey work, method is essential. Each child in rotation should be given a number, his name entered in the roll opposite that number, the fi/nl numbered to correspond, and put into a vertical filing cabinet opposite the corresponding number. If a colleC" tion of blood films is being made in the field to be

taken to a base laboratory to be stained subsequently) it is essential to fix the films at once, either by exposure for 3 minutes to methyl alcohol, or for 10 minutes to absolute alcohol. Staining may be deferred until the

base is reached, but old unfixed films stain extremely badly. Further, in survey work, blood films must be

protected from flies; flies are passionately fond ot

eating up blood films, and if you want to see what a

contaminated blood film can look like, allow a fly to crawl over it and then stain and examine it; the fil01 will be found full of bacteria, fungi, yeasts, extraneous protozoal C3?sts, and may even show helminthic ova. The simplest method of labelling the film is to take

a sharply pointed pencil or a needle and write the

patient's name across the film, using the dried film as

if it were a piece of frosted glass. Films should neither be so scanty that they can

hardly be seen, nor so thick that they look as if the

doctor had used the slide to stem a profuse haemorrhage-

Thick Blood Films. It is extremely desirable in all cases to supplement

the examination of the thin film by examination of a thick blood film. Again and again examination of thick films will detect parasites, where they are missed in the thin film. The only difficulty about thick films is that, if only ring forms are present, it is sometimes difficult to determine the species concerned. This, however, does not apply to growing trophozoites, schizonts, 91 gametocytes. Personally, I always examine both thin and thick films from every patient. The best method of preparing the thick film is that

advocated by James (1920). Inverting the clean slide

bring it to just touch the drop of blood issuing fronj the finger, and take four small drops of blood on 1

at the corners of a half inch square. The drops should be small and not large. With a rounded needle nex

pool the four drops into an even thick film covering the half inch square. Fuddling should be avoided, and the film must not be too thick. The procedure ]S

illustrated in Fig. 2.

Such a film takes two hours to dry #t room tempei&~ ture, or one hour in the 37?C. incubator. Drying ca0 be hastened by leaving the film under a rapidly moving fan. If the film be taken at the patient's bedside,

1

is best to cover it (e.g., with a saucer) to keep it froo1

F/a /.

Fig. 1.?Preparation of the Thin Blood Film.

Fig. 1.?Preparation of the Thin Blood Film.

Fig. 2.?Preparation of the Thick Blood Film. (After James, 1920.)

Fig. 2.?Preparation of the Thick Blood Film. (After James, 1920.)

May, 1931.] LABORATORY DIAGNOSIS OF MALARIA: KNOWLES. 273

contact with flies and dust, and to leave instructions that it be sent to the laboratory three hours later. If possible, it is best to take the thick film one day, and stain it the next.

Thin and Thick Films on the Same Slide.

This method is advocated by Sinton (1925) and it is special value in survey work. A thin film is spread

?n two-thirds of the slide, and a thick film at the other end. A transverse line across the slide between the two films is drawn with a blue grease pencil (vide Fig. 3).

Sinton (1925) gives the following particulars with Tegard to the value of thick blood films as contrasted with thin ones in survey work:?

P. vivax.

Thin Thick films. films.

Parasites in 100 fields.

Parasites found in 1 7.7 2 minutes.

lime required to 6.6 1 find parasites.

P. falciparum.

Thin films.

Thick films.

1

1

23

16

22

1

The Romanowsky Stains. The stains universally employed for the staining of

blood films are the Romanowsky ones. It is true that these stains probably do not give exactly true pictures ?f the parasites concerned, as they are not cytologicallv Sood stains. On the other hand, they have been so

Universally employed in all malaria work for thirty years that very few workers are familiar with the ttialaria parasites as stained by any other method. The Principle upon which these stains depend is that Medicinal (not pure) methylene blue contains a number ?f oxidation products, the most important of which is Methylene azure. When watery or alcoholic solutions

medicinal methylene blue and of eosin are mixed together, a series of loosely combined chemical bodies are formed, such as methylene-blue eosinate, methylene- azure eosinate, etc. These different compounds possess different affinities for different cell structures, and thus differential staining results, although but a single stain lst used. The red blood corpuscles stain a transparent Pink or orange colour; the nuclei of leucocytes, shades

violet; eosinophile granules in the coarsely granular e?sinophile leucocytes, red; neutrophile granules in the Polymorphonuclear leucocytes, yellow to lilac; mast Cell granules, deep violet; blood platelets, purple; the cytoplasm of malaria and other blood-inhabiting Protozoa, a bright

"

Cambridge" blue, and their chromatin a bright ruby red. I do not propose here to deal with the original

Romanowsky method. I would like to mention, however, that Colonel Christophers is very fond of it.

is economic and very suitable for laboratories where large numbers of films have to be stained dailyIn India, the two varieties of Romanowsky stains univer- sally employed are Leishman's and Giemsa's stains, and we may deal with these in turn.

L

In using both Leishman's and Giemsa's stains it is absolutely essential to use distilled water of neutral or very faintly alkaline reaction to dilute the stain. It is precisely this point in all probability which gives laboratory workers in the mofussil so much trouble with Leishman's stain, and which leads to so many of them abandoning it in favour of fixation with methyl alcohol and staining with Giemsa's stain. Further, the distilled water must be perfectly fresh, for stale samples of distilled water are very apt to become contaminated with Bodo, fungi, yeasts, etc., which may get into the stained film from this source and confuse the worker. Fresh, doubly distilled water is the best.

If there is any. doubt about the distilled water in use, proceed as follows:? Place 5 c.c. of the distilled water in a test tube and

add to it 5 drops of a 0.04 per cent, watery solution of bromo-cresol purple as indicator. If the water is faintly acid?as is usually the case?it will turn a pale yellow colour. In this case add very cautiously drop by drop a 1 in 1,000 watery solution of sodium carbonate. The water turns faintly alkaline and turns a purple-violet colour. At this point stop at once, and now use the water thus prepared for diluting the stain. (The faint trace of bromo-cresol purple present will not affect the staining.) Excess of soda solution must not be used. With regard to washing the films after staining,

ordinary tap water can usually be used. The Calcutta tap water always gives a definitely alkaline, purple, reaction with the bromo-cresol purple indicator.

Leishman's Stain. This stain is invaluable, if properly prepared and

properly used. It is both a combined fixative and stain, as it is made up with methyl alcohol. Its use, however, demands the most meticulous attention to details if good results are to be obtained. The ingredients used must be the very purest possible, Griibler's or Merck's powdered Leishman's stain, and Merck's methyl alcohol purissimum, free from any trace of acetone. Proceed as follows:? Scrupulously clean a 150 c.c. ground glass stoppered

bottle, a 100 c.c. graduated glass cylinder, and a glass (not porcelain) pestle and mortar. When they are

absolutely clean give them a rinse out with pure methyl alcohol. Weigh out 0.15 gramme of the Leishman's powder, and measure out 100 c.c. of methyl alcohol purissimum in the graduated cylinder. Put the weighed out Leishman's powder in the glass mortar, add a little methyl alcohol from the cylinder, and grind. Pour the dissolved stain from this into the glass bottle. Add more methyl alcohol to the residue, grind again, and again pour the dissolved stain into the bottle. Repeat this again and again until every particle of the stain has gone into solution and the whole 100 c.c. of methyl alcohol have been used up. The most essential

step in the procedure is to ensure absolutely complete solution of the stain. Next incubate the stain by placing it for 24 hours (or overnight, but not for longer than 24 hours) in the 37?C. incubator. Or the bottle

may be placed in a warm, dark cupboard for this period. This ripens the stain and gives greatly improved staining results. For use the stain should be poured into perfectly

clean 30 c.c. drop bottles. The stock solution should be kept in a dark cupboard. As thus prepared, Leishman's stain will keep for at least a fortnight in the hot weather in Calcutta. For the general practitioner, who

'

has only an occasional film to stain, the use of

Burroughs Wellcome & Co.'s "Soloid" Romanowsky (Leishman's) stain may be recommended. If fresh, the " Soloids" give an admirable stain, but if old results

are inferior; in fact one would like to recommend that the " Soloids" should be dated, as is done with sera and vaccines. To prepare the stain a drop bottle is made perfectly clean and then rinsed out with Merck's methyl alcohol pimssimum. Three of the "

Soloids," i.e., 0.045 gramme, are dropped into the bottle, and 30 c.c. of Merck's methyl alcohol purissimum added. The stopper is turned so that no fluid can escape from the bottle and complete solution of the

" Soloids "

Fig. 3.?Thin and Thick Blood Films on the same slide. Sinton's method. Fig. 3.?Thin and Thick Blood Films on the same slide. Sinton's method.

274 THE INDIAN MEDICAL GAZETTE. [May, 1931.

brought about by shaking. It is advisable to incubate the bottle afterwards overnight in the 37?C. incubator. To use Leishman's stain, proceed as follows:? 1. Lay the slide, blood film surface upwards, on a

staining rack. A convenient way to make such a rack is to fix two pieces of glass tubing parallel with one another across a Petri dish or basin with plasticine. This answers very well where large numbers of films have to be stained, and avoids mess. The rack must be dead level.

2. Drop Leishman's stain from the drop bottle on

to the film until its whole surface is covered. The

methyl alcohol in the stain fixes the film. This takes half a minute only, and the readiest way to measure this interval of time is to count up to 25 slowly mentally. Less than half a minute fails to fix the slide, and to

leave the stain on longer undiluted results in deposit of stain on the slide.

3. At the end of half a minute drop on to the slide double the corresponding number of drops of pure distilled water. (This distilled water should have been previously tested with bromo-cresol purple, as detailed above.) By tilting the Petri dish or the end of the slide allow the stain and water to mix thoroughly. Stain for about 10 to 15 minutes. Less than 15 minutes will not bring out stippling in the infected cells; whilst staining for more than 20 minutes is apt to result in deposit on the slide. If the stain has been properly prepared and diluted a thin golden scum will rise to the surface of the fluid.

4. Fill a bowl or beaker with water; this may be either distilled or tap water. Take the slide, still covered with the stain, and plunge it into the bowl of water so that the stain floods off. (Films stained by the Romanowsky methods should never have the stain drained off, as this is apt to result in deposit.)

5. Transfer to a small Petri dish full of water (dis- tilled or tap). Rock the dish gently until the film, which is bluish-green, commences to turn pink. This takes about a minute or less. At this stage remove

the slide from the Petri dish, and place it, film side downwards, leaning against a vertical wall or the ver- tical edge of the table, to dry. Films should never be blotted, as this is liable to introduce all sorts of foreign bodies. In examining thin films for malaria parasites, a most

useful lens is the l/7th inch oil immersion " fluorite"

objective.' This gives very clear definition, a clear field, and with it one can cover ground rapidly. If a

suspicious object be found, the l/12th inch oil immersion objective may be substituted for the l/7th inch or a

higher eyepiece employed. It is always good practice, before using the oil

immersion objective, to glance through the film with the l/6th inch dry objective to see that the nuclei of the leucocytes are deeply stained. If this is the case any malaria parasites present will presumably be well

stained.

Methyl Alcohol Fixation and Giemsa's Stain. This is preferred by many workers to Leishman's

stain, as it is a method which is very free from deposit. Again, the materials used must be of unimpeachable quality. The methyl alcohol should be Merck's purissimum, free from any trace of acetone. The preparation of Giemsa's stain is troublesome, and for the ordinary laboratory worker it is best to purchase ready made up undiluted Giemsa's stain. The writer has always used that supplied by the Central Research Institute at Kasauli for many years, and has found it admirable. Ready made Giemsa's stain prepared by Griibler & Co. can also be purchased from the Scientific Supplies Co., College Street Market, Calcutta. Methyl alcohol rapidly loses its fixative power in the

wet tropics. This has been shown by Ivnowles and Senior-White (1930). A batch of blood films was taken from a patient suffering from benign tertian malaria one day during September, and kept in a desiccator. A phial of Merck's methyl alcohol purissimum was opened and was kept on the laboratory bench. Each day films were taken from the batch in the desiccator, were fixed

with the same opened specimen of methyl alcohol, and. were stained with the same brew of Giemsa's stain. On the 1st and 2nd days the films stained perfectly. On the 3rd day the staining was fairly satisfactory, but it

failed to bring out Schiiffner's dots. On the 6th day the staining had become weak. On the 7th day and later it was hopeless; on the 10th day the parasites failed to take the stain at all, and only pigment could be seen. The moral of this is that methyl alcohol purissimum for use in the wet tropics should be put up in small, 5 c.c., phials, as the contents of the phial can only be relied upon for 5 or 6 days. To use Giemsa's stain:? 1. First fix the film. This can be done by covering

it for 3 to 5 minutes with pure methyl alcohol, or by dipping it for 10 minutes into ordinary absolute ethyl alcohol. Then wash thoroughly in water.

2. Dilute the Giemsa's stain, 1 part with 14 parts of distilled water. This is most readily done by

measuring out 10 or 15 c.c. of distilled water and dropping into it the corresponding number of drops from a drop bottle of the undiluted stain. The distilled water must be absolutely neutral or on the very faintly alkaline side of neutrality as tested by the bromo-cresol purple indicator as detailed above.

3. Place the slide film surface upwards in a Petri

dish, and flood with the stain. Stain for half an hour or longer. The more dilute the stain and the longer it

is allowed to act, the better the result. The stain may be made more dilute, the Petri dish covered to keep dust out, and the film stained overnight or for 12 to

18 hours. 4. As soon as the slide is stained, remove it from

the Petri dish, plunge it into a beaker full of tap water to flood off the stain. Then soak in a bath of fresh distilled (or tap) water in a Petri dish until the fita begins to turn from a bluish-green to a pink colour. At this stage remove the slide and place it to dry by leaning it, film side downwards, against a vertical surface. Do not blot.

The, Combined Use of Leishman's and Giemsa's Stains- This gives admirable results. The procedure is as

follows:? 1. Place the slide, film surface upwards, on a staining

rack. Cover it with undiluted Leishman's stain for 30 seconds.

2. Dilute the Leishman's stain with double the quan- tity of diluted Giemsa's stain (one drop to each c.c.)- Mix thoroughly.

3. Stain for 15 minutes or so. Then flood off the stain as in Leishman's method and differentiate as usual in a Petri dish of water.

Shute's Stain.

This method is given by S. P. James (1929), and is of special value in bringing out stippling in the infected red blood corpuscles. Details are as follows:?

" The stain is made with pure methyl alcohol (' free from acetone') and crystals of Leishman's stain. The reaction of the methyl alcohol must be tested before

making up the stain. This is done with the aid of an outfit for determining the hydrogen ion concentration, the indicator used being phenol red, and the range of

standard tubes being from pH 6.6 to pH 8.0 (Baird & Tatlock's outfit No. P. 2759). Into the test tube made of cordite glass which is supplied with the outfit, pipette 5 c.c. of the methyl alcohol to be tested. With a

separate pipette add 0.5 c.c. of a 0.01 per cent, solution of phenol red. Shake, and after a moment or two,

compare the tint with that in the standard tubes

provided. Every brand of methyl alcohol which we

have tested in this way gives a slightly acid reaction- After adding the indicator to 5 c.c. of the brand which we use, the tint corresponds nearly always with that of the standard tube marked 6.8, but sometimes with that marked 6.6. It is our practice to discard supplies of methyl alcohol which are as acid as is indicated by the tube marked 6.6, and we have had to do so even with some supplies which makers put up in hermetically sealed tubes ' for use in microscopic staining.'"

May, 1931.] LABORATORY DIAGNOSIS OF MALARIA: KNOWLES. 275

,, make up the stain, rinse a glass stoppered bottle * oroughly with some of the methyl alcohol that will

e used for the stain, and then put in 0.15 gramme of eishrnan's crystals (usually called

' Leishman's Powder'). Add 100 c.c. of the methyl alcohol. Shake r?m time to time during the next 24 hours, after which Period nearly all the crystals will be dissolved, and he stain will be ready for use. For several reasons ls unwise to make the solution in a pestle and mortar,

01 to filter it as is usually recommended in the text- books."

Next deal with the distilled water which will be

l'r j *n staining process and for washing the stained 'des. "We work with a 1-litre flask of distilled water

. ich has been treated as follows: Shake up the water

j.11 the flask and wash out a 5 c.c. pipette with water rom it; test 5 c.c. of the water in the same way as Was described for testing the methyl alcohol. Probably the water will be found to be at least as acid as is indicated bv the standard tube marked 6.6. Add to the water three or four drops of a saturated filtered solution of lithium carbonate, shake the flask to ensure thorough mixing, and repeat the test. Continue the Procedure of adding one or two drops of the lithium carbonate solution and of testing until the water in the nn.sk becomes exactly of an alkalinity indicated by observing that, after adding 0.5 c.c. of the phenol red solution to 5 c.c. of the water, the resulting colour batches the colour of the solution in the standard tube

parked 7.2. This is the degree of alkalinity that must be reached when the methyl alcohol is of an acidity represented by the tube marked 6.8."

To stain a blood film, drop four drops of the staining solution on the film, rock for ten seconds, add twelve orops of the distilled water, and mix thoroughly by tilting and rocking. We do not use a glass rod for huxing the water and the stain on the slide, because a rod often carries specks of dust or of cotton fibre ^hich are transferred to the slide; but a good deal of Practice in tilting and rocking the slide is required in order to obtain quick and complete mixing of the water and the stain without spilling some of it off the slide,

on the fingers, and without allowing any of the stain ^9 dry on the film before the water has reached it. r"0r. do we employ the usual practice of making a

barrier with a wax pencil across the proximal end of be slide, because, when this is done, particles of

^ethylene blue from the pencil invariably become

jnixed with the stain and alter the result. By using , our drops of stain and twelve drops of water, the stain diluted three times, which, we think, gives the best 1 esults. Four drops of stain, carefully applied, are

Quite sufficient to cover a film and, when twelve drops ?t water are added, the amount of fluid on the slide ls easy to manipulate so that none spills off the slide 0r reaches the fingers. We time the ten seconds during ^'bich the stain alone is on the film by a watch with a large second hand." For routine work the film should be stained for 30

^nutes, but in cases of infection with P. malarice a

Period of 45 minutes is desirable to bring out the very ne Ziemann's stippling in the infected cells.

, On the termination of staining, the stain must not e poured off the slide before beginning to wash the bn; a good stream of distilled water must be applied

?t once so that all the stain and deposit is flushed off

^ the first moment. Washing in the stream of distilled ?ftater should be continued for fifteen seconds by the Watch." We have used Shute's stain a good deal since

^ol. James' paper appeared. It is an admirable method

jn demonstrating stippling in the infected _ cells, but

ooth the chromatin and cytoplasm of malaria parasites stain go deeply by it that it is sometimes difficult to

identify the various parasite stages seen._ For routine

^agnostic work we prefer the combined use of

leishman's and Giemsa's stain as detailed above.

Difficulties with Thin Stained Films.

1. Deposit.?This is probably the commonest diffi- culty with Leishman's stain. It is especially apt to occur if the film has been stained too long, or if too long an interval has been allowed to elapse before diluting the stain with distilled water. To remove deposit from a stained film, first remove the cedar wood oil from the slide with xylol. Next let the film dry completely. Flush the slide?for an instant or two only?with rectified spirit and instantly immerse the slide in distilled water to remove the alcohol. The flushing with spirit must be practically instantaneous. The use of a rocker upon which to rest the slide

whilst staining will prevent the formation of deposit. There are several such patterns of rocker on the market, or a home made one can easily be made.

2. Staining too blue.?After removing the cedar wood oil, as above, immerse the slide for a few seconds (only) in 1 in 5,000 acetic acid in

_ water, then immediately transfer to distilled water.

3. Staining too weak.?If the batch of stain used is giving weak results, deeper staining can often

_ be

obtained by the use of 1 in 2,000 watery potassium carbonate solution to dilute the stain with, instead of distilled water.

4. Permanent preparations.?Films stained by any of the Romanowsky stains as a rule fade very rapidly, the first thing to disappear in old stained slides being the stippling in the red corpuscles. In order to make permanent preparations several writers recommend taking the stained film up through the graded alcohols to xylol and mounting in "neutral" Canada balsam. The writer has never had any success with this method, since however rapidly one passes through the alcohols a great deal of the stain is dissolved out, whilst he has never yet seen a truly neutral solution of Canada balsam. A much better?although crude?method is to let the stained film dry completely in the 37?C. incubator, and then cover it either with " Euparal "??a mounting medium prepared by Flatters & Garnett, 309, Oxford Road, Manchester, or with Gurr's Neutral Mounting Medium?prepared by G. T. Gurr, 136, New King Road, Fulham, London, S.W.6; then place on it a perfectly clean thin long cover slip 2X1 inches in size. Such preparations will keep without fading for at least six months, usually for a year.

5. Re-staining old slides.?In the writer's experience there is no really satisfactory method of re-staining old slides. Daniels recommends the following, though we have not found it very successful. Treat the film for a few minutws before staining with a mixture of 3 to 5 drops of glacial acetic acid to 1 oz. of absolute alcohol. Wash extremely thoroughly in neutral distilled water, to remove every trace of acid. Stain with Leishman's or Giemsa's stain in the usual manner. Old blood films take on a deep blue staining of the red corpuscles, instead of the normal orange-pink. To some extent this can be got rid of by flooding the slide with a 1 per cent, watery solution of acid sodium phosphate, but the

results are never too good.

Staining the Thick Film.

For his combined thin and thick film on the same

slide, Sinton recommends the following method:?

The thin film is first fixed by dipping it into either

methyl alcohol for 3 minutes or absolute alcohol for 10 minutes. It is then allowed to dry completely.

_ The

entire slide, both thin and thick films, is then cautiously flooded with Giemsa's stain (diluted, 1 drop to each

c.c.), and staining completed in the usual manner.

In the writer's experience, in endemic kala-azar areas, this procedure plasmolyses any Leishmania donovani which may be present, and is not of help in differentiat- ing kala-azar from malaria. In such areas it is

necessary to fall back on some method which both fixes The bottle should be of hard glass (green glass).

276 THE INDIAN MEDICAL GAZETTE. [May, 1931.

and dehtemoglobinises the thick film. The following is advocated by Knowles and Das Gupta (1924):?

1. Lay the film on a staining rack and gently flood the slide with the following mixture:?

Glacial acetic acid; 2.5 per cent, solution in distilled water .. .. 4 parts.

Tartaric acid, crystalline; 2 per cent.

solution in distilled water .. 1 part.

This mixture keeps indefinitely, and it is better to

keep the two solutions mixed, as fungi are apt to grow in the tartaric acid solution. It should be kept in a

stoppered bottle. 2. This solution dehsemoglobinises the film, and the

process should be watched. An ordinary thick film

quickly dehsemoglobinises, but films with thicker patches may take a little longer. The dehajmoglobinised film should show a grey-white colour.

3. As soon as dehsemoglobinisation is complete, drain off the fluid by gently tilting the slide. Flood the slide with methyl alcohol, and allow this to remain on for five minutes. The film is now dehsemoglobinised and fixed.

4. Drain off the methyl alcohol and wash the film

very thoroughly with (neutral or very slightly alkaline) distilled water. Every trace of acid must be got rid of.

5. Stain the film with Giemsa's stain, one drop to

each c.c., for 20 minutes or longer. Differentiate in the usual way with distilled water. Do not blot the

film, but let it dry by slanting it against a vertical surface, film side downwards.

If careful attention is paid to details in this technique, the results are excellent. Further, in the endemic kala- azar areas it gives some 67 per cent, of positive findings in cases of kala-azar, and is of special value in

differentiating between the two diseases.

The Diagnosis of Malaria by Culture.

Cultivation of the malaria parasites in vitro was first accomplished by Bass and Johns (1912). The original technique was simplified by J. G. and D. Thomson (1913, 1913a), whilst we now use a slight modification of the Thomsons' technique. Between the years 1921 and 1930 a great change has occurred in the methods employed in the diagnosis of malaria in the Proto-

zoology Department of the Calcutta School of Tropical Medicine. From 1921 to 1924 we relied chiefly upon the prolonged search of thin films. From 1924 to 1928 this was supplemented in each case by the examination of thick blood films in addition from each patient. From the beginning of 1929, however, we have used Bass' culture as a routine method of diagnosis in

every case. The method is so simple, so easy of appli- cation, and requires so little equipment that it should

always be employed in the laboratory in all difficult and obscure cases. The chief advantages of the cultural method in

malaria are the following:? 1. It will enable one to diagnose cases in which the

parasites are so scanty that they cannot be detected either in thin or thick films.

2. In cases where only ring forms are encountered in the film and one is uncertain of which species is present, a culture will enable one to diagnose the

species with certainty. 3. It is of special value in those all too common

cases where the patient has taken a 5-grain dose of quinine before sending for the doctor. This dose is sufficient to prevent one finding parasites in the thin (and often even in the thick) film, but quite insuffi- cient to control the fever. Under such circumstances, however, a Bass' culture will be positive.

4. It will enable one to detect the real frequency of mixed infections.

5. The life history of P. falciparum (as it occurs in man) can be studied in such cultures, and such forms as developing trophozoites, schizonts, and young gametocytes?which are only but rarely encountered in peripheral blood films?can readily be studied in cultures.

Details of the technique are as follows:? 1. A 50 c.c. thick walled (Erlenmeyer) flask is taken

and 20 to 30 glass beads are introduced into it. It

is plugged with cotton-wool, and the whole autoclaved. 2. A 50 per cent, watery solution of Merck's dextrose

purissimum is prepared, and sterilised by steaming for half an hour daily on each of three consecutive days.

3. A 5 c.c. syringe is sterilised. This can be accom- plished either by washing it out in turn with olive oil heated to 160?C., with sterile 1 per cent, sodium carbonate solution, and then with sterile normal saline; or by washing the syringe through repeatedly with a

mixture of 1 part of lysol to 9 of alcohol, then with rectified spirit, then repeatedly with normal saline. The

syringe must be sterile; it may contain traces of normal saline, but no trace of water should be present, as this will plasmolyse the parasites.

4. Take 5 c.c. of blood from the patient's vein and transfer it to the flask, being very careful that air

bubbles do not form. Next by gentle rotation of the flask on the laboratory bench by hand defibrinate the blood.

5. Sterile miniature test tubes, 12-J; X cms. in

dimensions, are next taken. Into the bottom of each introduce one drop of the dextrose solution with a

sterile capillary pipette. With a sterile capillary pipette aspirate off the defibrinated blood from the flask, and introduce it into the miniature test tubes, so that there is a column of blood about 21 cms. in depth in each. Plug with flamed cotton-wool.

6. Warm the upper part of the tube in order to

expel air. Whilst it is still warm, seal by fitting over its mouth a rubber teat. This gives partial anserobiosis.

7. Set the test tubes vertically in plasticine and incubate at 37?C. (Growth will also occur at room

temperature, but more slowly.) By the next morning the column of blood will have

settled in the test tube into three layers; a layer of clear plasma above; then a very thin leucocyte layer; with the red corpuscles below. With a capillary pipette aspirate material from the upper surface of the deposit of red corpuscles, prepare films in the usual manner, and stain by Leishman's or Giemsa's stain in the usual manner.

Cultures should be examined, if necessary, at 12, 24 and 48 hours after they have been put up. This method has still one further advantage. If one

wishes to test whether a given line of treatment in malaria has or has not eradicated all parasites from a patient's system?(as contrasted with merely clinical " cure ")?blood culture is the most delicate test that

can be applied. If a culture of 5 c.c. of the patient s blood fails to show parasites, it is evidence very con-

siderably in favour of the malaria infection having been exterminated.

The Leucocyte Count in Malaria.

At one time considerable reliance was placed on the total and differential leucocyte count in malaria. To-day, however, with the introduction of the thick film and cultural methods, the leucocyte count has

largely fallen into disuse. D. Thomson (1911), on a most careful examination of

the whole question of the leucocyte count in malaria, comes to the following conclusions:?

1. Fever plus leucocytosis plus an increased per- centage of polymorphonuclears is not malaria. It may mean sepsis.

2. Fever with leucopsenia and a large hyaline mono- nuclear increase?e.g., 12 to 15 per cent, (in the absence of kala-azar)?is strong confirmatory evidence of

malaria. 3. A persistently high large mononuclear percentage,

with from time to time a leucocytosis, should arouse the suspicion of malaria.

4. Leucocytosis per se does not necessarily exclude malaria. Stephens and Christophers (1904) state that a figure

of over 15 per cent, of large hyaline mononuclears (in the absence of kala-azar) is diagnostic of malaria, whilst

May, 1931.1 LABORATORY DIAGNOSIS OF MALARIA: KNOWLES. 277

if 20 per cent, be encountered (in the absence of kala- azar) further search of the films will usually show malaria parasites. Knowles (1920) contrasts the

leucocyte findings in kala-azar and malaria as follows.

Total leucocyte count.

Leucopsenia

Percentage of large mononu-

clears.

Percentage of Polymorphonu- clears.

Kala-azar.

Usually under 3,000.

Constant and progressive.

20 per cent, or

more.

Usually less than 50 per cent.

Malaria.

Between 3,000 and 5,000.

Fluctuating.

10 to 16 per cent., not usually over 20 per cent.

Usually more than 50 per cent.

Whilst leucopsenia is the general rule during the lebrile phases of an attack of malaria, leucocytosis may l)e associated with severe infections with P. falciparum, Especially during the earlier rigors. In Calcutta, the chief value of the total and differ-

ential leucocyte count is in differentiating influenza and dengue from malaria. In influenza the film will often show some evidence of a polymorphonuclear leuco- cytosis; in dengue leucopsenia?which reaches its Maximum about the fourth day of the disease?is in evidence, whilst there is a complete or almost complete absence of coarsely granular eosinophile leucocytes. The finding of hsemozoin pigment in the cytoplasm the large hyaline mononuclear leucocytes is evidence

that the patient is or has been suffering from malaria. I he observer must satisfy himself, however, that what ls present is true haemozoin, and neither the azure

granules of chromatinic origin so frequently seen in the ^ytoplasm of the large hyaline mononuclears, nor dust. -Dust, when present on the slide, may be on the surface ?f such a cell, but will also be present between the ?ells.

The Personal Factor in Diagnosis. The last point to which I should like to refer before

concluding, is that of the personal factor in diagnosis, by which I mean the training and experience of the observer. This point has been dealt with on experi- mental lines by Knowles and Senior-White (1930). -I1 our "tests" were carried out, as follows:?

Test 1.?Eight microscopes were set out, and under each a single malaria parasite was focussed in the field; *?r the most part a single ring. Forty-five students irom the Calcutta D.T.M. class of 1928-29 volunteered '?r the test. Each passed in turn to each of the eight microscopes, was allowed three minutes at it, and recorded his diagnosis of the species present. No talking or communication was allowed, and at the end of the test the writer collected all papers. The slide ^'as not moved. The result was an accuracy of only 41 per cent. Now

these students were near the end of their class; they had had no end of malaria films for study during the Previous six months, and were just about to go up for their examination for the Diploma. The results were s? disappointing that a second test was decided upon.

Test 2.?Seven microscopes were set out, and under each a malaria blood film. The same students as

before volunteered. Each passed to each microscope in turn, was allowed five minutes at it, and was allowed to use the mechanical stage, and to see as much of 'he film as he could in that time. Otherwise, conditions ^'ere the same as in Test 1. The percentage of accuracy now rose in this test to 80 per cent. " e may conclude that the accuracy of the

" brigade

"

0r "

divisional laboratory" standard of worker is

Approximately SO per cent.

Test 3.?In order to check these results, we decided to repeat tests 1 and 2 with " experts"?or at least volunteers who have been diagnosing malaria species during a lifetime of laboratory work in the tropics. The volunteers for tests 3 and 4 were: Lieut-Col. H. W. Acton, c.i.e., Lieut-Col. R. Knowles, i.m.s.; Dr. P. A. Maplestone, Dr. C. Strickland, Dr. L. E. Napier?all of the Calcutta School of Tropical Medicine ; Mr. R. Senior-White, Malariologist, Bengal-Nagpur Railway; Capt. B. S. Chalam, Malariologist, Eastern Bengal Railway; Dr. T. N. Sur, Offg. Professor of

Pathology, Calcutta Medical College; and Dr. B. M. Das Gupta, Assistant Professor of Protozoology, Calcutta School of Tropical Medicine.

Test 3 consisted in a repetition of test 1, with these volunteers. Here there can be no question of "

accuracy," since it is a matter of the opinion of one "

expert" with regard to species as against that of

another. The percentage of "

agreement" in test 3 was 63 per cent.

Test 4 consisted of a repetition of test 2 with the "

expert" volunteers. The percentage of " agreement

"

with regard to the species present now rose to 93 per cent. The moral from these four tests is obvious. It is the

usual custom in laboratories to diagnose the species of malaria parasite present on the first or first and second parasite forms encountered, and then to discard the slide. Such a procedure will give a wrong diagnosis of species as often as not. Some standard method of examining blood films should be adopted by all labo- ratory workers. As few such workers are willing to spend more than 10 minutes on the examination of a

case, we suggest that not less than 8 minutes should be spent on searching the thin film, together with at least 2 minutes on a thick blood film from the same patient. In all cases it is desirable that a Bass' culture should also be taken. In conclusion, I should like-to emphasise again that

accuracy in the laboratory diagnosis of malaria is very largely dependent upon the most meticulous attention being paid to details of technique.

References.

Bass, C. C., and Johns, F. M. (1912). The cultiva- tion of malarial plasmodia (Plasmodium vivax and Plasmodium falciparum) in vitro. Journ. Exp. Med., XVI, 567. Boyd, M. F. (1930). An Introduction to Malariology.

Harvard University Press, Cambridge, Massachusetts.

James, S. P. (1920). Malaria at Home and Abroad. London. James, S. P. (1929). A note on the Shute technique

for staining malaria parasites with Leishman's stain and on the stippling in infected red blood corpuscles which it reveals. Trans. Roy. Soc. Trop. Med. and Hyg., XXIII, 269. Knowles, R. (1920). A study of kala-azar. Indian

J ourn. Med. Res., VIII, 140.

Knowles, R., and Das Gupta, B. M. (1924). The

diagnosis of kala-azar by examination of thick blood films. Indian, Med. Gaz., LIX, 438. Knowles, R., and Senior-White, R. (1930). Studies

in the parasitology of malaria. Indian Med. Res. Mem. No. 18. (Supplementary to the Indian J ourn. Med. Res.).

Sinton, J. A. (1925). Notes on the "thick-film" method of examination for malarial parasites. Indian J ourn. Med. Res., XII, 537.

Stephens, J. W. W., and Christophers, S. R. (1904). ?

The Practical Study of Malaria and other Blood Parasites. Second Edition. London.

Thomson, D. (1911). The leucocytes in malarial

fever; a method of diagnosing malaria long after it is

apparently cured. Ann. Trop. Med. and Parasit., V, 83

Thomson, J. G., and Thomson, D. (1913). The culti- vation of one generation of benign tertian malarial parasites (Plasmodium, vivax) in vitro by Bass' method. Ann. Trop. Med. and Parasit., VII, 153.

278 THE INDIAN MEDICAL GAZETTE. [May, 1931.

Thomson, J. G., and Thomson, D. (1913a). The growth and sporulation of the benign and malignant tertian malarial parasites in the culture tube and in the human host. Ann. Trop. Med. and Parasit., VII, 509. Proc. Roy. Soc., B, LXXXVII, 77.