Chapter XLV: Blood Examinations in the Diagnosis of Tropical Diseases (1)

In a short chapter on such a large subject only the more important methods and findings can be considered. As regards interpretation of blood findings in various tropical diseases one may note in the recent work of Schilling-Torgau the difficulties which at present beset the subject. Until some universal agreement as to standard methods of technique and in particular complete accord as to the characteristics of the diagnostic cells can be arrived at, conflicting reports as to findings must of necessity be obtained.

In taking up this subject it has seemed convenient to divide it into 4 heads: (1) The microscopical examination of fresh preparations or stained blood smears; (2) blood culture methods; (3) serological examinations, and (4) other practical methods of haematological study.

In the companion volume on laboratory work I have endeavored to
take up rather in detail the various methods and techniques but in
this chapter I shall only give single methods or point out short
cuts in well-recognized ones or make suggestions as to new methods
of blood study which may eventually aid us in diagnosis.

Those who work in temperate climates cannot realize the
difficulties which beset the tropical laboratory worker from the
lack of proper assistance, damaging effects of heat and moisture on
stains and media and, of greater importance, the impairment of that
driving energy so necessary for the carrying out of complicated
methods. A short and simple method has a far greater value in the
tropics than at home.

BLOOD PREPARATIONS

To obtain blood, except for blood cultures, use either a platino-iridium hypodermic needle which can be sterilized in the flame, a small tenotome, or a surgical needle with cutting edge.

Needles should be sterilized by boiling since flaming dulls the
edge. A steel pen with one nib broken off or the glass needle of
Wright may also be used. To make a glass needle, pull straight
apart a piece of capillary tubing in a very small flame. Tap the
fine point to break off the very delicate extremity. Scarcely any
pain attends the use of such a needle. In puncturing either the tip
of the finger or lobe of the ear a quick piano-touch-like stroke
should be used. The ear is preferable, as it is less sensitive
and there is less danger of infection. Before puncturing, the
skin should be cleaned with 70% alcohol and allowed to dry. It is
advisable to sterilize the needle before using it.

Note that in order to secure in the specimen a cell count that
corresponds to that obtaining in the circulation as a whole it
is necessary to massage the ear vigorously prior to making the
puncture. Subsequently there should be no manipulation of the part,
the blood examined being that which exudes freely. This procedure
renders more likely the finding of blood parasites.

The first drop of blood which exudes should be taken up on the paper of the Tallquist haemoglobinometer, using subsequent ones for the blood pipettes and smears. If it is necessary to make a complete examination, it is rather difficult to draw up the blood in the pipettes, dilute it, and then get material for fresh blood preparations and films without undue squeezing, which is to be avoided. Of course, fresh punctures can be made. Ordinarily, complete blood examinations are not called for. It is only a white count or a differential count or an examination for malaria that is required.

As a practical point it is very rare that a red count is indicated.
There is one point not sufficiently recognized by physicians and
that is that a routine blood examination is not apt to be as
carefully conducted as one calling for a specific feature. Without
disparaging the necessity of routine examination of urine as well
as blood it is a fact that the internist who knows what he wants
gets better results from the laboratory man.

THE MICROSCOPICAL EXAMINATION OF FRESH PREPARATIONS OR STAINED BLOOD SMEARS

As regards haemocytometry it may be stated that in the tropics the counting of red cells is required more frequently in comparison to white ones than is the case in temperate climates where probably 100 white counts are necessitated as against 1 red count. This is on account of the frequency of secondary anaemias in the tropics.

The idea that time may be saved by making a white and red count
from the same preparation is not borne out practically so that it
is better to make white and red counts separately.

As a diluting fluid for red counts a normal salt solution,
preferably about 0.9%, answers perfectly and if desired may be
tinged with neutral red, methyl green or gentian violet to bring
out white cells. When available, however, I prefer a 2½% aqueous
solution of potassium bichromate for red cell counts.

=Rulings.=—The most desirable rulings are those of Türck, Zappert
and Neubauer.

In these the entire ruled surface consists of nine large squares,
each 1 mm. square. These are subdivided, and in the central large
square are to be found the small squares used for averaging the
red cells. These small squares are 1/20 mm. square and are arranged
in nine groups of 16 small squares by bordering triple-ruled lines.
As the unit in blood counting is the cubic millimeter, if one
counted all the white cells lying within one of the large squares
(1 mm. square), he would have only counted the cells in a layer
one-tenth of the required depth, so that it would be necessary to
multiply the number obtained by 10. This product, multiplied by
the dilution of the blood, would give the number of white cells in
a cubic millimeter of undiluted blood. The Neubauer ruling is the
most satisfactory.

=Bürker Haemacytometer.=—Some workers prefer the _Bürker
haemacytometer_. In this there are two ruled wedge-shaped pieces of
glass, separated at their bases, which take the place of the ruled
disc of the Thoma apparatus. Two oblong pieces of glass are on
either side of the ruled wedges and are 0.1 mm. higher, thus taking
the place of the shelf. Clamps fix a cover-glass on these shelves
giving a space 1/10 mm. over the ruled surfaces. The blood is run
in by capillarity from the mixing pipette. I gave up this type of
counter because the clamps made manipulation awkward.

=Thoma-Levy Chamber.=—In the Thoma-Levy modification of the Bürker
apparatus the central portion of the slide is cut away and in this
depression is cemented a rectangular strip of glass, divided by a
central channel. Each half of this strip of glass has a Neubauer
ruling on it so that one can make a white count on one side and a
red one on the other, simply touching the tip of the red pipette
to the space separating the under surface of the cover-glass from
the ruled rectangular slips on one side and then with the white
pipette repeating the same on the other side. An advantage of the
Thoma-Levy is that the original thickness of the slide makes the
shelf on which the cover-glass rests instead of the support being
strips on either side of the ruled surfaces and cemented to the
slide. The Neubauer ruling is undoubtedly the most satisfactory of
the haemacytometer rulings, its rulings being simpler than those of
the Türck system. The unit square in all these haemacytometers is
the small square for counting red blood cells, 1/20 mm. square.

=To Make a Red Count.=—Having a fairly large drop of blood,
apply the tip of the 101 pipette to it and, holding the pipette
horizontal, carefully and slowly draw up with suction on the rubber
tube a column of blood to exactly 0.5. The variation of 1/25 of an
inch from the mark would make a difference of almost 3%. If the
column goes above 0.5, it can be gently tapped down on a piece of
filter-paper until the 0.5 line is cut. Now insert the tip of the
pipette into some diluting fluid, and revolving the pipette on its
long axis while filling it by suction, you continue until the mark
101 is reached.

A variation of 1/25 of an inch at this mark would only give an
error of about 1/30 of 1%. This gives a 1-200 dilution. After
mixing thoroughly by shaking for one or two minutes, the fluid
in the pipette below the bulb is expelled (this of course is
only diluting fluid). A drop of the diluted blood of a size just
sufficient to cover the disc when the cover-glass is adjusted, is
then deposited on the disc and the cover-glass applied by a sort
of sliding movement, best obtained by using forceps in one hand
assisted by the thumb and index-finger of the other.

In red counts we use exclusively the small 1/20 mm. squares which
are in groups of 16 bounded by triple-ruled lines.

The depth of fluid over the ruled surface is 1/10 mm., hence each
of these small squares is 1/10 × 1/20 × 1/20 = 1/4000 of a c.mm.,
so that it takes 4000 such spaces to equal the unit for blood
counting (1 c.mm.). My practice in making red counts is to count
the red cells in five of the groups of 16 small squares. This in
normal blood is about 100 for the 16 squares. After counting 5
groups of 16 we have counted the red cells of 80 small squares
which is 1/50 of 4000 (the number in the 1 c.mm. unit). For this
reason 50 × 200 (the blood dilution) = 10,000, so that it is only
necessary to multiply the number of red cells found in 5 groups of
16 small squares by 10,000 in order to obtain the number of red
cells per c.mm. For more accurate determination the process can be
repeated with a second or third drop of the diluted blood, which
would give an average from 160 or from 240 small squares.

=To Count White Cells.=—Draw up the blood in the white pipette
to the 0.5 line. Then, still holding the pipette as near the
horizontal as possible, because the column of blood tends to fall
down in the larger bore, draw up by suction a diluting fluid which
will disintegrate the red cells without injuring the whites. The
best fluid is 0.5% of glacial acetic acid in water. This makes the
white cells stand out as highly refractile bodies. Some prefer to
tinge the fluid with neutral red or gentian violet. The 0.5 mark is
preferred because it takes a very large drop of blood to fill the
tube up to the 1 mark and if there is much of a leucocytosis a 1 to
10 dilution is not sufficient.

The blood having been drawn up to 0.5, we have a dilution of 1 to
20.

Making a preparation, exactly as was done in the case of the red
count, we count all of the white cells in one of the large squares
(1 sq. mm.). The cross ruling greatly facilitates this. Note the
number. Then count a second and a third square. Strike an average
of the large squares counted and multiply this by 10, as the depth
of the fluid gives a content equal to only 1/10 of a c.mm. Then
multiply by the dilution.

EXAMPLE.—First large square 50; second large square 70; third
large square 60. Average 60. Then 60 × 10 × 20 = 12,000, the number
of leucocytes in 1 c.mm. of blood. In order to save time the count
is preferably made with a low power (⅔-inch objective) as the
leucocytes stand out like pearls. It is more accurate, however,
to use a higher power, so that pieces of foreign material may be
recognized and not enumerated as white cells.

If one will accustom himself to comparing the distribution of the leucocytes in a well-made stained dried-blood film, prepared according to Ehrlich’s cover-glass method, with that in a haemacytometer preparation, he can readily acquire an experience which will enable him to determine with considerable accuracy the degree of leucocytosis by the examination of a stained, cover-glass preparation alone.

After making a blood count, the haemacytometer slide should be
cleaned with soap and water and then rubbed dry, preferably with
an old piece of linen. As the accuracy of the counting chamber
depends upon the integrity of the cement, any reagent such as
alcohol, xylol, etc., and in particular, heat, will ruin the
instrument. The pipettes should be cleaned by inserting the ends
into the tube from a vacuum pump, as a Chapman pump. First draw
water or 1% sod. carbonate solution through the pipette, then
alcohol, then ether, and finally allow air to pass through to dry
the interior. If the interior is stained, used 1% HCL in alcohol.
If a vacuum pump is not at hand, a bicycle pump or suction by mouth
will answer.

PREPARATIONS FOR THE STUDY OF FRESH BLOOD

Many authorities prefer a fresh blood specimen to a stained dried smear in the study of parasites of the blood. In malaria in particular there is so much information as to species to be obtained from a fresh specimen that the employment of this method should never be neglected. While waiting for the film to stain one has five or six minutes which could not be better spent than in examining the fresh specimen which only requires a moment to make.

=Manson’s Method.=—Have a perfectly clean cover-glass and slide.
Touch the apex of the exuding drop of blood with the cover-glass
and drop it on the center of the slide. The blood flows out in a
film which exhibits an “empty zone” in the center. Surrounding this
we have the “zone of scattered corpuscles,” next the “single layer
zone” and the “zone of rouleaux” at the periphery. It is well to
ring the preparation with vaseline. When desiring to demonstrate
the flagellated bodies in malaria, it is well to breathe on the
cover-glass just prior to touching the drop of blood.

=The Method of Ross= is very easy of application and gives most
satisfactory preparations. Take a perfectly clean slide, and make
a vaseline ring or square of the size of the cover-glass. Then,
having taken up the blood on the cover-glass, drop it so that
its margin rests on the vaseline ring. Gently pressing down the
cover-glass on the vaseline makes beautiful preparations which
keep for a very long time. If it is desired to study the action
of stains on living cells, this method is also applicable. A very
practical way to do this is to tinge 0.85% salt solution containing
1% sodium citrate (the same as is used in opsonic work) with
methylene azur, gentian violet, or methyl green. With a capillary
bulb pipette, take up one part of blood, then one part of tinted
salt solution. Mix them quickly on a slide and then deposit a
small drop of the mixture in the center of the vaseline ring and
immediately apply a cover-glass and press down the margins as
before. This method will be found of great practical value.

PREPARATION AND STAINING OF DRIED FILMS

When preparations are desired for a differential count, Ehrlich’s method of making films is to be preferred, as the different types of leukocytes are more evenly distributed. In making smears by spreading, there is a tendency for the polymorphonuclears to be concentrated at the margin while lymphocytes remain in the central part of the film.

=Cover-slip Films.=—In _Ehrlich’s method_ we have perfectly clean
dry cover-slips. Take up a small drop of blood without touching the
surface of the ear or finger. Drop this cover-glass immediately on
a second one and as soon as the blood runs out in a film, draw the
two cover-slips apart in a plane parallel to the cover-glasses.
Ehrlich uses forceps to hold the cover-glasses to avoid moisture
from the fingers, but I find I can work more quickly and
satisfactorily with the fingers alone. The method shown in Fig. 150
is a very convenient one. In making malarial smears it is better to
wash the finger or ear with soap and water to get rid of all grease
and dirt. Then dry thoroughly before puncturing. Alcohol is not so
efficient.

Slides and spreaders should be absolutely clean and grease-free.
Scrubbing with soap and water, thorough rinsing and drying, then
subjecting the slide to the flame to make it grease-free is
satisfactory.

For removing dirt and grease from skin, a mixture of acetone, 40;
alcohol, 60; is the best and quickest means. A bottle is kept on
hand, with the puncturing needle embedded in the stopper.

For cleaning a slide, nothing equals Bon Ami. Rub up some with the
wet finger, rub the slide with the lather until there is a friction
squeak; let dry; polish with a clean, dry cloth. This is far
better than soap and water, alcohol, ether and flaming combined.
Note how a drop of water spreads on a glass so treated.

=Smears on Slides.=—Of the various methods of spreading films on slides, that described by Daniels is quite satisfactory. In this the drop of blood is drawn along and not pushed along. The films are even, can be made of any desired thickness by changing the angle of the drawing slide, and there is little liability of crushing pathological cells. Take a small drop of blood on the end of a clean slide. Touch a second slide, about ½ inch from end, with the drop and as soon as the blood runs out along the line of the slide end, slide it at an angle of 45° to the other end of the horizontal slide. The blood is pulled or drawn behind the advancing edge of the advancing slide. An angle less than 45° makes a thinner film; one greater, a thicker film.

Instead of a slide a square cover-glass may be used and if the edge
be smooth it makes a more satisfactory spreader than the slide.

Instead of the Daniels method I prefer to take up the drop of blood
on the slide on which the smear is to be made, about ½ inch from
the end. Then apply the spreader slide and so soon as the drop runs
along the end of the spreader slide proceed as above described.
This method is shown in Fig. 150.

=Spreaders.=—Of the various methods of making smears by means of
cigarette paper, rubber tissue, needles, etc., the best seems to be
to take a piece of capillary glass tubing and use this instead of a
needle in making the film. There is one advantage about the strip
of cigarette paper touched to the drop of blood and drawn out along
the slide or cover-glass, and that is that it is almost impossible
not to make a working preparation by this method.

THICK-FILM METHODS

Such methods are of the greatest practical value in searching
for malarial parasites when they are in very small numbers in
the peripheral circulation, in finding trypanosomes, relapsing
fever spirochaetes and filarial embryos. Ruge’s method so brings
out the polymorphonuclears that such a technic can be used for
opsonic index. Many workers prefer the _Ross thick-film method_ in
examining for malaria. In this about one-half of a drop of blood
is smeared out over a surface about equal to that of a square
cover-glass and allowed to dry. It is then flooded with 1/10
of 1% aqueous solution of eosin for about fifteen minutes. The
preparation is then gently washed with water and then treated with
a polychrome methylene-blue solution. After a few seconds this is
carefully washed off and the preparation dried and examined.

James smears out an ordinary drop of blood so that it makes a
circular smear about ¾ inch in diameter. This may be easily
accomplished with a spatulate toothpick. When dry, treat the blood
smear with alcohol containing HCl (Alcohol 50 cc., HCl 10 drops)
until the haemoglobin is dissolved out. Then wash thoroughly in
water for five or ten minutes. Allow to dry and then stain as
ordinarily with the Wright or Giemsa stain.

=Ruge’s Method.=—The best thick-film method is that of Ruge. After the blood has dried well gently move the slide about in a glass containing a 2% solution of formalin to which has been added 1% of glacial acetic acid. After laking is completed, as shown by disappearance of brown color, treat the slide in the same way in a glass of tap water to remove all traces of acid. Next wash gently in distilled water and stain with dilute Giemsa (1 drop to 1 cc. of water) for twenty to thirty minutes. Wash in water and allow to dry without heat or blotting paper. Some workers prefer to stain the dried thick smear for one hour in a jar containing dilute Giemsa stain (1 to 40) without previous fixation or dehaemoglobinization. At present, I make my thick films by taking up a large loopful from the exuding drop of the puncture wound.

This is deposited at one end of the slide and from it three or four more daubs are made in succession toward the other end of the slide. These daubs are quickly smeared out before coagulation takes place in the first daub.

With all thick-film methods it is extremely important to have
thorough drying of the smear before dehaemoglobinizing or staining.
This ordinarily requires one or two hours in the air or twenty to
thirty minutes in the incubator. It is particularly important in
working with such smears, although holding for ordinary smears, to
protect them from flies, ants, etc., as such insects will eat up
the smear in a few minutes if left exposed.

=Fixation of Film.=—In Wright’s, Leishman’s, and other similar
stains the methyl-alcohol solvent causes the fixation. In staining
with Giemsa’s stain, or haematoxylin and eosin, separate fixation
is necessary. For Giemsa either absolute alcohol (ten to fifteen
minutes) or methyl alcohol (two to five minutes) answers well.

For haematoxylin and eosin, heat gives the best results. The
best method is to place the films in an oven provided with a
thermometer. Raise the temperature of the oven to 135°C. and then
remove the burner. After the oven has cooled, take out the fixed
slides or slips.

One of the handiest methods is to drop a few drops of 95% alcohol
on the slide or cover-glass. Allow this to flow over the entire
surface; then get rid of the excess of alcohol by touching the edge
to a piece of filter-paper for a second or two. Then light the
remaining alcohol film from the flame and allow the burning alcohol
to burn itself out.

=Staining Blood-films.=—As separate staining with eosin and methylene blue rarely gives good preparations and as the modifications of the Romanowsky stain recommended are easy to make and employ, and give much greater information, the separate method of staining is not recommended.

_Wright’s Method._—The stain is made by adding 1 gram of methylene
blue (Grubler) to 100 cc. of a ½% solution of sodium bicarbonate in
water. This mixture is heated for one hour in an Arnold sterilizer.
The flask, containing the alkaline methylene-blue solution should
be of such size and shape that the depth of the fluid does not
exceed 2½ inches. When cool, filter the methylene blue solution,
and add 500 cc. of a 1 to 1000 eosin solution (yellow eosin, water
soluble). Add the eosin solution slowly, stirring constantly
until the blue color is lost and the mixture becomes purple with
a yellow metallic lustre on the surface, and there is formed a
finely granular black precipitate. Collect this precipitate on
a filter-paper and when thoroughly dry (dry in the incubator
at 38°C.) dissolve 0.3 gram in 100 cc. of pure methyl alcohol
(acetone-free). Wright lately has recommended using 0.1 in 60 cc.
methyl alcohol. This constitutes the stock solution. For use filter
off 20 cc. and add to the filtrate 5 cc. of methyl alcohol.

A _modification by Balch_ is very satisfactory. In this method
instead of polychroming the methylene blue with sodium bicarbonate
and heat, the method of Borrel is used. Dissolve 1 gram of
methylene blue in 100 cc. of distilled water. Next dissolve 0.5
gram of silver nitrate in 50 cc. of distilled water. To the silver
solution add a 2 to 5% caustic soda solution until the silver oxide
is completely precipitated. Wash the precipitated silver oxide
several times with distilled water. This is best accomplished
by pouring the wash-water on the heavy black precipitate in the
flask, agitating, then decanting and again pouring on water.
After removing all excess of alkali by repeated washings, add the
methylene-blue solution to the precipitated silver oxide in the
flask. Allow to stand about ten days, occasionally shaking until
a purplish color develops. The process may be hastened in an
incubator. When polychroming is complete, filter off and add to the
filtrate the 1 to 1000 eosin solution and proceed exactly as with
Wright’s stain.

In _Leishman’s method_ the polychroming is accomplished by adding
1 gram of methylene blue to 100 cc. of a ½% solution of sodium
carbonate. This is kept at 65°C. for twelve hours and allowed to
stand at room temperature for ten days before the eosin solution is
added. The succeeding steps are as for Wright’s stain.

_In all Romanowsky methods_ distilled water should be used. If not obtainable, the best substitute is rain-water collected in the open and not from a roof.

_Method of staining:_

1. Make films and air dry.

2. Cover dry film preparation with the methyl-alcohol stain for one
minute (to fix).

3. Add water to the stain on the cover-glass or slide, drop
by drop, until a yellow metallic scum begins to form. It is
advisable to add the drops of water rapidly in order to eliminate
precipitates on the stained film. Practically, we may add 1 drop of
water for every drop of stain used.

4. Wash thoroughly in water until the film has a pinkish tint.

5. Dry with filter-paper and mount.

Red cells are stained orange to pink; nuclei, shades of violet;
eosinophile granules, red; neutrophile granules, yellow to lilac;
blood platelets, purplish; malarial parasites, blue; chromatin,
metallic-red to rose-pink.

_Giemsa’s Modification of the Romanowsky Method._—This is one of the most perfect of the modifications. The objection is that greater time in staining films is required than with the Wright or Leishman method and the stain is very expensive.

Take of Azur II eosin 0.3 gram. Azur II 0.08 gram.

Dissolve this amount of dry powder in 25 cc. of glycerine at 60°C.
Then add 25 cc. of methyl alcohol at the same temperature. Allow
the glycerine-methyl alcohol solution to stand overnight and then
filter. This is the stock stain. To use: Dilute 1 cc. with 10 to 15
cc. of distilled water. If 1 to 1000 potassium carbonate solution
is used instead of water it stains more deeply. These same dyes,
mixed with methylene violet, are now obtainable commercially as a
powder ready for solution in methyl alcohol.

The alkaline diluent is used to obtain the coarse stippling in malignant tertian (Maurer’s clefts). Having fixed the smear with methyl alcohol for one to five minutes, pour on the diluted stain, and after fifteen to thirty minutes wash off and continue washing with distilled water until the film has a slight pink tinge. For _Treponema pertenue_ stain from one to twelve hours.

=Haematoxylin Staining.=—While the Romanowsky methods are more
satisfactory for differential counts and for the demonstration of
the malarial parasites, and especially for differentiating species,
yet by reason of the liability to deterioration in the tropics of
methylene blue the haematoxylin methods may be preferable. Many
workers in blood-work and cytodiagnosis prefer the haematoxylin.

1. Fix the film either by heat, with methyl alcohol for two minutes
or with Whitney’s fixative. Heat is to be preferred.

2. Stain with Meyer’s hemalum or Delafield’s haematoxylin for from
five to fifteen minutes according to the stain. Frequently three
minutes will be found sufficient. To make the hemalum, dissolve
0.5 gram of haematin in 25 cc. of 95% alcohol. Next dissolve 25
grams of ammonia alum in 500 cc. of distilled water. Mix the two
solutions and allow to ripen for a few days. The stain should be
satisfactory in two or three days.

3. Wash for two to five minutes in tap water to develop the
haematoxylin color.

4. Stain either with a 1 to 1000 aqueous solution of eosin or with
a one-half of 1% eosin solution in 70% alcohol. The eosin staining
only requires fifteen to thirty seconds.

5. Wash and examine.

DIFFERENTIAL COUNT

In making a differential count I would recommend the following from the directions of Schilling-Torgau. It will be remembered that considerable interest was raised a few years ago in what was termed the Arneth index. In this the more normal, more mature, better resisting polymorphonuclears were considered to have 3 or 4 lobes to the nuclear structure, even occasionally 5. The immature cells had only one or at most two lobes to the nucleus. The index was obtained by adding the percentages of cells showing 1 and 2 lobes to ½ the percentage of those with 3 lobes. As will be understood a high percentage of these immature cells was unfavorable in prognosis. These cells are graded from left to right, I, II, III, IV, V, as to separate masses in the nucleus, so that when the percentage is shoved or displaced to the left it indicates an increase in the immature cells.

Schilling-Torgau divides his polymorphonuclears into: (1) The
myelocyte which is always of course a pathological cell. (2)
The immature form polymorphonuclear. In this there is a close
resemblance to the neutrophile myelocyte but there is a nuclear
indentation instead of the round nucleus of the myelocyte. It is
this cell which often puzzles us as to whether to regard it as a
true myelocyte. It is the meta-myelocyte of many authorities. (3)
Between the mature or segmented polymorphonuclear and the immature
one or metamyelocyte we have what may be designated the band-form
nucleated one. These show the type of nucleus which one is familiar
with in the nucleus of the transitional. (4) The mature, multilobed
or segmented nucleus of the typical polymorphonuclear.

It would seem that if all tropical workers would agree upon some single method of recording differential counts it would be advantageous.

Under the blood findings in liver abscess, in a paragraph to follow
in this chapter, I give suggestive counts indicating the value of
Schilling-Torgau’s method.

In the differential count he not only divided up the
polymorphonuclears but makes no separation of small from large
lymphocytes. Although I have always divided lymphocytes into large
and small ones I believe it unnecessary and unpractical and shall
henceforth group all such cells in one grouping. The statement that
large mononuclears and transitionals are cells of a similar origin,
type and significance has always been my view.

SCHEME OF SCHILLING-TORGAU
-------------------------------------------------+-------+---------------
| Normal| Percentage
Type of Cell |Percen-|Moderate Sepsis
| tage |(W. C. 14,000)
-------------------------------------------------+-------+---------------
1. Mast cells | 1 | 1.0
2. Eosinophiles | 3 | 1.5
{ a. myelocytes | 0 | 0.5
3. Neutro- { b. immature forms (metamyelocytes) | 0 | 5.0
philes { c. band-form (Stabkernige) | 4 | 13.5
{ d. multilobed (Segmentkernige) | 63 | 64.0
4. Lymphocytes | 23 | 10.5
5. Large mononuclears and transitionals | 6 | 4.0
-------------------------------------------------+-------+---------------

BLOOD CULTURING

Among tropical diseases, only malta fever, kala-azar and plague demand this method of diagnosis, although there are met commonly in the tropics many cosmopolitan diseases in which blood culturing is a principal diagnostic procedure. There are many ways of carrying out the cultivation of organisms from the blood but the one which may be strongly recommended is the following. The blood is obtained from a vein, the overlying skin of which has been painted with tincture of iodine to insure a sterile skin surface.

A stout hypodermic needle is attached to about 6 inches of rubber
tubing which in turn is pushed over a downward bent glass tube
which passes through a doubly perforated rubber stopper. A second
glass tube, which also passes through the stopper, is bent upward
to be attached to a second piece of rubber tubing for use in
suction by the mouth. The glass tubes project about ½ inch below
the under surface of the rubber stopper and above are about 2½
inches including the bent arm. This system of tubing and stopper is
readily sterilized by boiling in a pan or instrument sterilizer.
As a receptacle for the blood we employ Erlenmeyer flasks of 100
cc. capacity, containing 25 cc. of salt solution with 1% of sodium
citrate, for prevention of coagulation. Blood that contains 0.2%
of sodium citrate will not coagulate so that a 0.5% solution could
be used instead of the usual 1% one. These citrated salt solution
flasks are plugged with cotton, sterilized and kept on hand ready
for immediate use, so that we only have to sterilize the stopper
and tubing by boiling and flame the neck of the flask when removing
the cotton plug to insert the stopper of the system. By suction we
can take any amount of blood desired. I usually count the drops
of blood as they fall into the citrated salt solution allowing
16 drops to the cc. In this way we may take from 10 to 25 cc. of
blood at the bedside and then later on in the laboratory, when it
is convenient, inoculate various media from the flask. For plates
add 2 or 3 cc. of this citrated blood to 6 or 8 cc. of melted
agar at 45°C. The blood mixture can also be added to various
sugar bouillons for fermentation reactions. Finally we place the
receiving flask in the incubator and culture it as well as the
other media.

=Clot Cultures.=—A very simple method is to take blood with a Wright U-tube. Then centrifuge and use the serum for agglutination tests and the clot, emulsified in some liquid medium, for the blood culturing. For paratyphoid culturing _bile media_ are preferable, just as for typhoid.

=Lyon Blood Tube.=—Quite recently I have been using the blood
tube recommended by Lyon. To make it, heat a 5- or 6-inch section
of ¼ inch tubing in the centre and draw out as for making 2
bacteriological pipettes. Divide and seal off the large end in the
flame. Next seal off the capillary end. Then apply a very small
flame to a point on the large end just before it begins to taper
to the capillary part. The heat causes the heated sealed-off air
inside to force out a blow hole. To use: Break off the sealed
capillary end and allow the capillary end to suck up blood from a
drop just as with the Wright tube. I consider this tube superior to
the Wright one.

=N. N. N. Medium.=—In culturing blood for protozoa the N. N. N.
medium is usually employed. Novy and MacNeal originally used a 12½%
meat infusion containing 2½% agar, 2% peptone, 1% normal sodium
carbonate solution and ½% salt. To one part of this agar, melted
and cooled to 60°C., they added twice the amount of defibrinated
rabbit’s blood. In the N. N. N. medium, as modified by Nicolle,
there is beside the blood only salt and agar—no peptone or meat
extractives.

Citrated salt solution was the medium used by Rogers in the
cultivation of splenic juice from kala-azar patients.

THE TAKING OF BLOOD FOR SEROLOGICAL TESTS

This can be done with the Wright tube, pipetting off the clear serum after centrifuging. We usually draw blood from a vein by use of the system of stopper and tubing described under blood culturing but employing an empty, sterile centrifuge tube.

Agglutination Tests

There are two methods of testing the agglutinating powers of a serum—the microscopical and the macroscopical or sedimentation method.

=For the microscopical method= draw up serum to the mark 0.5 of
the white pipette. Then draw up salt solution to the mark 11. This
when mixed gives a dilution of 1 to 20. One loopful of the diluted
serum and one loopful of a bouillon culture or salt solution
suspension of the organism to be tested gives a dilution of 1 to
40. One loopful of the 1-20 diluted serum and 3 loopfuls of the
bacterial suspension give a dilution of 1-80. These two dilutions
answer in ordinary diagnostic tests. The red pipette with a 1-100
or 1-200 dilution may be used where dilutions approaching 1-1000
are desired. Having mixed the diluted serum and the bacterial
suspension on a cover-glass, we invert it over a vaselined concave
slide and examine with a high power dry objective (⅙ inch). It
is simpler to make a ring of vaseline to fit the cover-glass and
make the mixture of diluted serum and culture in the centre of
this ring or square. Then apply the cover-glass, press it down on
the vaseline ring and examine as with the ordinary hanging drop.
In making dilutions it is preferable to use salt solution, as
the phenomenon of agglutination requires the presence of salts.
Ordinarily, thirty minutes is a sufficient time to wait before
reporting the absence of agglutination. Agglutination is more
rapid at body temperature than at room temperature. In reporting
agglutination, always give time and dilution. It is absolutely
necessary that a control preparation be prepared in every instance;
that is, one with the bacterial culture alone or with a normal
serum of the same dilution as the lowest used. Some normal sera
will agglutinate in 1 to 10 dilution, and group agglutinations (as
paratyphoid with typhoid serum) may occur in 1 to 40 or possibly
higher. It is very unusual for sera to agglutinate any other
bacteria then the specific one in dilutions as high as 1-80.

=Macroscopic Agglutination.=—For the macroscopical or
sedimentation test, take a series of small tubes (⅜ × 3 inches) and
deposit 1 cc. of salt solution in each of the series. Now, having
taken an empty test-tube, drop 4 drops of serum in it and then
add 12 drops of salt solution. This approximately gives 1 cc. of
a 1 to 4 dilution of the serum. It is more exact to make the 1 to
4 dilution with a graduated pipette. With a rubber-bulb capillary
pipette, which has been graduated to hold 16 drops or 1 cc., draw
up the contents of the tube containing the 1 to 4 serum and add it
to the next tube containing 1 cc. of salt solution. This gives 2
cc. of a dilution of 1 to 8. Now mix thoroughly by drawing up and
forcing out with the bulb pipette, and then withdraw 1 cc. and add
to the next tube containing 1 cc. of salt solution. This gives a
dilution of 1 to 16. Having mixed as before, again withdraw 1 cc.
of the mixture and add it to the 1 cc. in the next tube. We now
have a dilution of 1 to 32. Again withdrawing 1 cc. and adding it
to the fourth tube containing 1 cc. of salt solution we have a
dilution of 1 to 64. In tube 1 there is now 1 cc. of a dilution of
the serum of 1 to 8; in tube 2, there is 1 cc. of a dilution of 1
to 16; in tube 3 of 1 to 32. Tube 4 contains 2 cc. of 1 to 64. The
dilutions can be carried on in the same manner to any extent that
may be desirable. In cholera agglutinations we may run up to 1 to
5000 or thereabouts. Of course, where such dilutions are employed,
we generally start with 2 cc. of 1 to 50 in the first tube. When
we have completed the series, each tube having 1 cc. of diluted
serum, and the last 2 cc., we remove with the pipette 1 cc. from
the last tube and discard it by ejection from the pipette leaving
1 cc. in the last tube. Now adding 1 cc. of a culture of typhoid
or any other organism, we have the dilution of the serum in each
tube doubled. Tube 1 now contains a serum in dilution of 1 to 16,
acting on the bacteria; tube 2 of a 1 to 32; tube 3 of a 1 to 64.
Now place these tubes in the incubator and, after two to five hours
or overnight, we examine for the clearing up of the supernatant
fluid. If the serum in a certain dilution agglutinates, the clumps
gravitate to the bottom and the upper part becomes clear. If so
desired, these dilutions may be carried on to 1 to several hundred
in the same way. It is safer to work with dead cultures instead of
living ones. To prepare, take a twenty-four-hour agar slant culture
of typhoid or paratyphoid and emulsify in salt solution (about 6
cc. to a slant).

By adding 0.1 of 1% of formalin to the typhoid emulsion and placing
in the ice-box the cultures will be found sterile in about three
days. The emulsion should be shaken twice daily while undergoing
sterilization in the ice-box. Such cultures are not easily
contaminated and appear to retain their agglutinable qualities
for several months. The macroscopic methods are preferable with
such dead cultures. For our Dreyer emulsions we use a two-billion
suspension of typhoid or para-typhoid organisms in 1 cc. of the
formalinized culture.

_Combination of Microscopical and Macroscopical
Methods._—Microscopic: Prepare dilutions of serum as above
described and take from each or several of the series, a loopful
of the diluted serum. For control use a loopful of salt solution.
Place on a cover-glass and add loopful of bouillon culture of the
living organisms. Make hanging drop preparation, report after one
hour at room temperature. Use ⅔ inch lens for examination.

Macroscopic: Add to each of the series, including the control, an
equal amount of an emulsion of killed organisms.

The method of using a slide with two vaselined rings, one
containing an emulsion in the specific serum and the other in salt
solution, is of great practical value. This method is described
under cholera.

_Complement Fixation._—Complement fixation tests have been
employed in the diagnosis of several tropical diseases but do not
seem to be at present sufficiently reliable or practical with the
exception of that for yaws and tularaemia. The chief difficulty
with complement fixation tests for suspected sera is to obtain a
reliable antigen. Should we later on be able to prepare bacterial
antigens as satisfactory as Noguchi’s acetone-insoluble antigen
is for the Wassermann test there may be a field for such tests in
tropical pathology.

OTHER PRACTICAL METHODS OF HAEMATOLOGICAL STUDY

Haemoglobin Estimation

The standard method now is the estimation of the oxygen capacity of the blood, using some gas apparatus, such as Van Slyke’s. Otherwise, the most accurate instrument for this purpose is the Miescher modification of the v. Fleischl haemoglobinometer.

The apparatus is expensive, requires considerable time and care in
the making of estimations, and is exclusively an instrument for a
well-equipped laboratory.

=Sahli’s Haemometer.=—A simple and apparently very scientific
instrument which has been recently introduced is the Sahli
modification of the Gower haemoglobinometer. Instead of the tinted
glass, or gelatin colored with picrocarmine to resemble a definite
blood dilution, Sahli uses as a standard the same coloring matter
as is present in the tube containing the blood. By acting on blood
with 10 times its volume of N/10 HCl, haematin hydrochlorate is
produced, which gives a brownish yellow color. In the standard
tube, which is sealed, a dilution representing 1% of normal blood
is used. To apply this test, pour in N/10 HCl to the mark 10 on
the scale of the graduated tube. Add to this 20 cubic millimeters
of the blood to be examined, drawn up by the capillary pipette
provided. So soon as the mixture assumes a clear bright dark-brown
color, which requires about ten minutes, add water drop by drop
until the color of the tubes matches. The reading of the height of
the aqueous dilution on the scale gives the Hb. reading. The tubes
are encased in a vulcanite frame with rectangular apertures. This
gives the same optical impression as would planoparallel glass
sides.

The most accurate readings are obtained with artificial light in a
dark room but almost as satisfactory comparisons can be obtained
with natural light from a window. It is advisable to turn the ruled
side around so that one may match colors without being influenced
in his determination by the scale.

The apparatus must be kept in a dark place as strong light will
change the color of the standard tube. It is recommended that the
N/10 HCl be preserved with chloroform.

The Dare instrument is excellent.

Pappenheim has recently proposed an instrument in which the
blood is converted into haematin hydrochloride as for the Sahli
apparatus. Instead of matching a standard tube, with a dilution
made drop by drop in the second tube, the new method employs a
wedge-shaped glass vessel showing graduations of the brown colored
blood, the treated blood being matched against the wedge-shaped
container (Autenreith-Koenigsberger Haemocolorimeter).

=Tallquist’s Haemoglobin Scale.=—This is a small book of specially
prepared filter-paper with a color-scale plate of ten shades of
blood colors. These are so tinted as to match blood taken up on
a piece of the filter-paper and are graded from 10 to 100. So
soon as the blood on the filter-paper has lost its humid gloss,
the comparison should be made. This is best done by shifting the
blood-stained piece of filter-paper suddenly from one to the other
of the holes cut in each shade—the piece of filter-paper being
underneath the color plate. At least a square centimeter of the
filter-paper should be stained by the blood. Daylight coming from
a window to the rear or at the side should be used in making the
comparison. The error with this method is probably not over 10%
after a little experience. If the colored plate is not kept in the
dark, the tints tend to fade.

NORMAL BLOOD

In considering what may be termed normal blood, it must be borne in mind that the normal varies for men, women, and children:

Hb. Red Cells Leucocytes

Men, 90 to 110%, 5 to 5½ million, 7500.
Women, 80 to 100%, 4½ to 5 million, 7500.
Children, 70 to 80%, 4½ to 5 million, 9000.

COLOR INDEX

This is obtained by dividing the percentage of the haemoglobin by the percentage of red cells, 5,000,000 red cells being considered as 100%.

To obtain the percentage of red cells it is only necessary to
multiply the two extreme figures to the left by two. Thus if a
count showed the presence of 1,700,000 red cells the percentage
would be 34 (17 × 2 = 34). If the Hb. percentage in this case were
50, then the color index would be 50 ÷ 34, or 1.4.

In normal blood the color index is, approximately, 1.

In anaemias we have three types of color index: 1. The pernicious
anaemia type which is above 1. Here we have a greater reduction in
red cells than we have of the haemoglobin content of each cell.
For example, in a case of pernicious anaemia we have 2,000,000 red
cells (40%) and 60% of haemoglobin, 60 ÷ 40 = 1.5. 2. The normal
type, when both red cells and haemoglobin are proportionally
decreased, as in anaemia fallowing haemorrhage. 3. The chlorotic
type. Here there is a great decrease in haemoglobin percentage,
but only a moderate decrease in the number of red cells. Hence the
color index is only a fraction of 1. For example, in a case of
chlorosis we have 40% of haemoglobin and 4,000,000 red cells, 40 ÷
80 = 0.5.

One can judge fairly well the approximate color index by noting the
character of the staining of the red cells. This is faint in bloods
of low color index and deeper than normal in cells in a case with
high color index.

TESTS FOR AGGLUTINATION AND HAEMOLYSIS OF THE RED CELLS (TRANSFUSION)

Transfusion of blood has become a method of greatest value in many types of anaemia.

In the selection of a donor for blood for transfusion it is always necessary to try his red cells against the serum of the recipient as well as the patient’s red cells against the serum of the donor, in order to prove the absence of haemolyzing or agglutinating bodies.

Certain persons have isohaemolysins in their blood which dissolve
the red cells of other persons and in paroxysmal haemoglobinuria
autohaemolysins may be present which can destroy the patient’s
own red cells. This autohaemolysin seems operative only when a
low temperature is followed by a high one. When haemoglobinaemia
exists the liver converts it into bile pigment, causing bilious
stools and jaundice. If one-sixth of the red cells are destroyed
haemoglobinuria results.

In the following tables, two groupings of blood are given. Both are quoted in text-books, and both are in common use. Although that of Moss is more generally followed in France, England and the United States, the obvious desirability of having one classification universally employed, in order to avoid confusion and the possibility of serious accidents, has led to the recommendation that, on the basis of priority the grouping of Jansky be adopted.

In 1907, Jansky described the following four groups.

Group 1, the serum of which agglutinates the corpuscles of Groups
2, 3 and 4, while the cells are not agglutinated by any serum.

Group 2, the serum of which agglutinates the corpuscles of Groups
3 and 4, but not those of Groups 1 and 2, while the corpuscles are
agglutinated by the serum of Groups 1 and 3, but not by those of
Groups 2 and 4.

Group 3, the serum of which agglutinates the cells of Groups 2
and 4, but not those of Groups 1 and 3, while the corpuscles are
agglutinated by the serum of Groups 1 and 2, but not by those of
Groups 3 and 4.

Group 4, the serum of which does not agglutinate any corpuscles,
while the corpuscles are agglutinated by the serum of all other
groups.

In 1910, Moss made the following classification:

Group 1, the serum of which does not agglutinate any corpuscles,
while the corpuscles are agglutinated by the serum of Groups 2, 3
and 4.

Group 2, the serum of which agglutinates the corpuscles of Groups 1
and 3, while the corpuscles are agglutinated by the serum of Groups
3 and 4.

Group 3, the serum of which agglutinates the corpuscles of Groups 1
and 2, while the corpuscles are agglutinated by the serum of Groups
2 and 4.

Group 4, the serum of which agglutinates the corpuscles of Groups
1, 2 and 3 while the corpuscles are not agglutinated by any serum.

At the present time it is accepted that the four groups considered
include all adult persons; i.e., that the classification is
complete.

=Before transfusing= carry out the following tests:

From a vein take about 1 cc. of blood in a centrifuge tube
containing 1% of sod. citrate salt solution; then shift the stopper
of the blood system to a dry centrifuge tube and draw into it about
3 or 4 cc. of blood. Throw down the citrated blood, pipette off the
supernatant fluid and wash the sediment with normal saline.

Again pipette off the saline after centrifuging and make a 10%
emulsion of the red-cell sediment in normal saline.

Centrifuge the coagulated blood in the other tube and collect the
serum which separates from the clot.

Carry out these procedures for both donor and recipient.

Tests: 1. In a small test-tube deposit 1 drop of the donor’s 10%
red-cell emulsion and then add 4 drops of the recipient’s serum.

2. Treat similarly 1 drop of the recipient’s red-cell emulsion with
4 drops of the donor’s serum.

3. Treat 1 drop of donor’s red-cell emulsion with 4 drops of his
serum.

4. Treat 1 drop of recipient’s red-cell emulsion with 4 drops of
his serum. Finally add 1 cc. of salt solution to each of the four
tubes, shake gently and place in incubator for two hours.

5. Treat one drop of donor’s red-cell emulsion with four drops of
salt solution.

6. Treat one drop of recipient’s red-cell emulsion with four drops
of salt solution.

Tubes 5 and 6 are controls of saline.

Tests 3 and 4 should fail to show either agglutination or
haemolysis. If agglutination or haemolysis appears in tubes 1 or 2,
the donor is not satisfactory; but if agglutination appears in tube
2 only, he may be used in an emergency.

Some prefer to keep the tubes overnight in ice-box after the
preliminary examination following incubation.

_Lee’s Technique._—For the regular carrying out of this method one
should keep on hand the sera of individuals belonging to groups 2
and 3 (Moss). To carry out the tests prepare a suspension of the
donor’s red cells by dropping 2 or 3 drops of his blood into 1 cc.
of citrated salt solution. Deposit a platinum loopful of standard
serum 2 on a slide and emulsify in it a loopful of the donor’s
red-cell suspension. A concave slide with two concavities is
convenient, the serum-cell emulsion being made on the cover-glasses
which are to be inverted over the vaseline ringed concavities. The
agglutination can be observed with a high power magnifying glass or
the ⅔-inch objective. Agglutination, when it occurs, is usually
complete in five to fifteen minutes. Repeat test with serum 3. If
both test sera agglutinate the donor’s cells he belongs to group
one. If neither agglutinate, to group four.