NationalLibraryOfScotland/medical-history-of-british-india huggingface dataset:
20 THE INDIAN JOURNAL OF VETERINARY SCIENCE AND ANIMAL HUSBANDRY [ II, I
The egg-shell subsequently splits up into an outer and an inner shell. In some
cases, e. g., Ascaris, the segmentation of the ovum starts after the eggs are laid by
a female. In most of the Strongylidæ a few segments are formed in the uterus and
the eggs are laid in this condition. In both these cases the embryos are formed in
the eggs after their deposition and the species are classed oviparous. In some
cases, parasitic generation of Strongyloides stercoralis, for example, a complete
embryo is formed within the egg while the latter is still in the uterus and is ready
to be hatched soon after the egg is laid; in such cases the species are called
ovoviviparous. In other cases, e.g., Dracunculus medinensis and Trichinella
spiralis, the embryos are hatched in the uterus and the species are called viviparous.
During the growth of the embryos of nematodes usually four moults or ecdyses
take place, at which the outer cuticle along with the lining of the mouth, œsopha¬
gus and rectum is cast off. Before each ecdysis a new cuticle is formed under-
neath the old one and some changes occur in the structure of the embryo. Four
moults divide the larval life into different stages, thus four ecdyses entail five
stages in the life-history. Usually after the second moult the larva enters an
infective stage, which, in some cases, may be reached in an egg before hatching.
Frequently the second moult takes place after the ensheathed larva is transferred
to its host.
The various ways in which the life-history of nematodes is completed and the
manner in which the definitive host is infected may be described briefly as
follows :—
I. Without an intermediate host.
(a) In Ascaris an egg passed out in the fæces of the host develops in it an
infective larva. This is taken by a new host along with food or drinking water.
The larva is hatched in the alimentary canal and undergoes a course of migration
in the body of the host before settling down in the intestine and developing into
an adult.
In Oxyuris equi the mode of development and transference to the host is the
same as in Ascaris, but the larvæ instead of undergoing a course of migration through
the various organs of the host settle down at once in the alimentary canal and
develop into adults.
(b) In Strongyloidea usually when the eggs are laid they contain only some
segments. These pass out with the fæces of the host. After some time the larvæ
of the first stage emerge out of them. These reach the infective stage after two
moults. The infective larvæ reach the definitive host either with food or with
drinking water. In some cases, e.g., in Ancylostoma, the larvæ may enter the host
HELMINTH PARASITES AFFECTING DOMESTIC ANIMALS
21
through the skin and reach the alimentary canal through the circulatory or
lymphatic system. In the genus Trichonema the infective larvæ after reaching
the alimentary canal penetrate the mucus membrane where they produce nodules
and undergo development up to sexual maturity; after this they again enter the
lumen of the alimentary canal and pass the rest of their existence. In the genus
Strongylus, on the other hand, the infective larvæ after reaching the alimentary
canal undergo a course of migration through some organs of the body, such as the
liver, lungs and circulatory system before they settle down finally in the alimentary
canal and develop into adults.
(c) In Strongyloides stercoralis the parasitic female deposits ova containing
embryos. From these the larvæ hatch in the fæces or in the soil, and they may
develop either into infective larvæ or give rise to a bisexual generation, whose progeny
develop into infective or " filariform " larvæ. The definitive host gets the infec-
tion either through the mouth or the skin. In the latter case the larvæ reach the
alimentary canal either through the vascular or lymphatic system as in
Ancylostoma.
II. Without an intermediate host.
(a) In Spiruridæ and Dracunculidæ an intermediate host is usually required
for the completion of the life-history. This is generally an Arthropod or a lower
vertebrate. The second stage larva becomes encapsuled in its organs or its body
cavity and there undergoes a second moult and is converted into a third stage or
infective larva. The final host becomes infected by the ingestion of the inter-
mediate host containing an infective larva.
In Filariidæ the larvæ are present in the vascular system of the host and are
taken up by some blood-sucking insect along with its feed. They migrate from the
stomach of the insect to the muscles where they grow to the infective stage, after
which they pass to the proboscis. The final host becomes infected when bitten by
such an insect.
(b) In Trichinella spiralis the fertilized female penetrates the intestinal wall and
enters the lymph spaces. Here it produces a large number of embryos which pass
to the voluntary muscles and become encapsuled there. These when ingested by
another suitable host develop to sexual maturity in its intestine.
TECHNIQUE.
Collection. The helminth parasites are notorious for infecting all the soft organs
of the body and body cavities of their hosts. Great, care must therefore be exercised
when searching for them. Large parasites offer no difficulty in being detected and
22 THE INDIAN JOURNAL OF VETERINARY SCIENCE AND ANIMAL HUSBANDRY [ II, I
can be picked up with ease from the site of infection. Very often, however, they
inhabit the minute passages of an organ such as bile-ducts or alveoli of the lungs
or are embedded in an organ in a very obscure manner. In such cases several
parallel cuts must be made with a knife in the organ which should afterwards be
pressed gently with the hands, when the parasites will be squeezed out of their
resting places. The organ should then be washed in a basin of water. The con-
tents should be allowed to stand for about five minutes, when the parasites will
sink to the bottom of the basin. The supernatant liquid having been decanted off,
the parasites may be collected. Those which produce nodules in the connective
tissue may be collected after cutting open the nodules, and exposing the parasite.
Sometimes, however, especially in the case of larvæ, the nodules produced are very
minute and extreme care should be exercised in teasing out the larvæ from them.
Frequently, however, when collecting parasites of the alimentary canal of a
large animal their presence remains unnoticed on account of the bulky contents of
the gut. In such cases the gut should be cut into several portions and the contents
of each portion emptied into a large enamelled basin and mixed with water. The
contents should then be passed through three sieves having increasingly fine
meshes, the sieve with the finest mesh being used last. The material arrested in
the sieves should be carefully searched for larger parasites. The filtrate after the
last sieve should be collected in a basin and allowed to stand for about five
minutes. After decanting off the supernatant liquid, careful search should be made
for parasites in the residual contents of the basin. Having dealt with their con-
tents, the portions of the gut should be washed vigorously in a basin of water.
The washings should be allowed to stand, as before, for five minutes, and after
decanting off the supernatant liquid, search should be made for parasites in the
remaining portion.
The parasites in all cases should be collected in a dish containing normal saline
solution. If ordinary tap water is used instead of normal saline solution, the cuticle
of many nematodes will burst and the worms rendered unfit for a detailed examina-
tion.
Examination in the living condition. The value of examining worms in the
living condition cannot be over-estimated. In the case of trematodes and those
nematodes in which the cuticle is very thin, nearly the whole of their anatomy can
be distinguished under the low power of a microscope if examined after the applica-
tion of a cover slip on the parasite. Some details, such as the excretory system
of trematodes, especially the movements of the cilia, cannot be studied after the
parasites have been killed. Similarly, among the nematodes, a few structures such
as the various papillæ and the rays of the genital bursa are seen to greater advan-
tage in the living condition.
HELMINTH PARASITES AFFECTING DOMESTIC ANIMALS
23
Fixing. This term is applied to the killing of the living tissues in such a
manner that the details of their structure will be retained as closely as possible to
that found in the living condition. The parasitic worms fall under two phyla:
the PLATYHELMINTHES consisting of Trematoda and Cestoda and the NEMA¬
THELMINTHES comprising Nematoda and Acanthocephala. The fixatives used for
these two naturally different groups of animals also fall into two distinct categories.
Before fixing, the parasites should be shaken vigorously in normal saline solu-
tion, so as to free them completely of the mucus or other foreign matter adhering
to them.
1. Fixatives for PLATYHELMINTHES.
(a) For ordinary examinations and toto-preparations, the best fixative is 5 per
cent. formalin which should be warmed to steaming point when used. The worms
fixed in this fluid may either remain in it, or be transferred to 70 per cent. alcohol
after washing away the whole of the formalin in running water.
(b) For histological examinations, however, one of the following fixatives may
be used :—
(i) Zenker. (2 gms. potassium dichromate, 1 grm. sodium sulphate, 100 c. c.
distilled water, 5 gms. mercuric chloride and 5 c. c. acetic acid.) The
specimens to be fixed in this fluid should be allowed to remain in it
for twenty-four hours and then washed in running water for another
twenty-four hours or more. In order that the specimens shall be free
from mercuric chloride they should be transferred to a very dilute
solution of tincture of iodine and kept in it for a couple of hours;
after which they should be washed in 70 per cent. alcohol and pre-
served in it.
(ii) Bouin. (Saturated solution of picric acid 75 parts; 40 per cent.
formalin 25 parts; acetic acid 5 parts.) The specimens to be fixed
in this fluid should remain in it for at least 18 hours; after which
they should be transferred to 50 per cent. alcohol, then to 70 per cent.
alcohol in which they may be preserved.
(iii) Mann's fluid. (Picric acid 10 gms. ; mercuric chloride 25 gms., 40 per
cent. formalin 300 c. c. and water 1,000 c. c.) The specimens to be
fixed should be kept in this fluid for at least 12 hours. They should
then be washed in running water for 24 hours or more, and afterwards
transferred to a dilute solution of tincture iodine as in the case of
Zenker, washed in 70 per cent. alcohol and preserved in it.
24 THE INDIAN JOURNAL OF VETERINARY SCIENCE AND ANIMAL HUSBANDRY [ II, I
2. Fixative for NEMATHELMINTHES.
These should be fixed in a steaming
mixture of 70 per cent. alcohol and 5 per cent. glycerine. The specimens after being
fixed in this fluid may be left in it.
The tissues intended for section cutting should be fixed in Zenker, Bouin or
Mann's fluid as described above or in 10 per cent. formalin, after which they should
be washed in running tap water for twenty-four hours.
Flattening. In the case of large PLATYHELMINTHES it has been observed
that the specimens undergo contraction when transferred to a fixative. This is
avoided in the case of cestodes, as recommended by Meggitt, by taking hold of the
tail end of the worm by means of a pair of forceps, immersing the worm in a beaker
containing a fixative, and then drawing it along the edge of the beaker. By repeating
this process a few times, complete flattening and extension of the worm is ensured.
The large trematodes can be fixed in their natural form by placing them between two
glass slides and tying the latter with a fine cotton thread. The slides should then
be transferred to a fixative. Special care should be exercised so as not to damage
the worms by putting too much pressure upon them. Thick cestodes, e.g., the
species of Anoptocephala, may also be fixed in this manner.
Preservation. Although the method of preservation is known to every student
of Biology and hints on it would appear to be superfluous, the writer wishes to
record here an experience of his own. Specimens are usually preserved in 70 per
cent. alcohol and are stored in glass tubes with corks. When the alcohol comes
in contact with the cork, the tannin of the latter is dissolved in it and stains the
specimens black. This black coloration of the specimens renders them incapable
of being stained by the usual agents, and the internal anatomy remains obscure in
such cases. In order to avoid this, it is better to store the specimens in stoppered
phials, but in cases where the storage in the corked tubes cannot be avoided, it is
advisable to use 5 per cent. formalin as the preservative. In order to bleach the
specimens which have become black, the following procedure, recommended by
Baylis, may be adopted. Soak the specimens for 3 to 4 days in pure beech-wood
creosote, then transfer for the same period to 5 per cent. hydrochloric acid in 70 per
cent. alcohol.
Staining. This may be done successfully in the case of trematodes and
cestodes. In the case of nematodes and acanthocephala it is rendered impossible
on account of the thick cuticle that usually covers the body. The internal anatomy
of these is studied by clearing them in one of the media given later. Small speci-
mens of nematodes and acanthocephala may, however, be stained successfully
by any one of the methods given below, provided a few punctures are previously
made in the cuticle.
HELMINTH PARASITES AFFECTING DOMESTIC ANIMALS.
25
1. * Acetic alum carmine. Dilute solutions of this stain should be used. The
exact degree of dilution is determined by adding to distilled water just sufficient
concentrated stain until the mixture attains the colour of port wine. The speci-
mens to be stained may be left in it for 24 to 48 hours.
2. *Delafield's hæmatoxylin. This stain also should be used in very dilute
solutions. The exact degree of dilution is determined by adding to a tube of
distilled water a few drops of concentrated stain until the mixture becomes violet
in colour. The specimens to be stained may be left in this solution for 24 hours.
Differentiation. The specimens stained in any one of the methods are not, as
such, good for mounting. The excess of stain of certain organs should therefore
be removed by acidulated water (water 100 c.c. and 0.5 c.c. hydrochloric acid) or
acidulated alcohol (70 per cent. alcohol 100 c.c, hydrochloric acid 0.5 c.c.). This
removal of the excess of stain from certain organs so as to distinguish them from
others is called " differentiation ".
It is advisable to differentiate hæmatoxylin stained specimens in acidulated
water for they can be converted to brilliant blue coloration by washing them after
differentiation with ordinary tap water. The trace of alkali usually present in
tap water brings about this change from red to blue. The specimens stained with
acetic alum carmine should be differentiated in acidulated alcohol.
Permanent mounting. The specimens after staining and differentiating in the
manner stated above should be dehydrated successively in the graded alcohols (30,
50, 70, 90 per cent. alcohols and absolute alcohol); the grades of alcohol already
covered before differentiation should be avoided. They should then be cleared
in clove oil, cedar-wood oil or xylol until they become perfectly transparent, and
then mounted in canada balsam.
Section cutting. For studying the histology of the worms recourse has to be
made to section cutting. Occasionally the specimens may be inextricably buried in
the tissues of the host or the specimens may be so thick, e.g., various Amphistomes,
that section cutting is indispensable, in order to distinguish even the gross features
of the anatomy. It is therefore thought necessary to describe the process briefly.
The fixed material should be dehydrated in graded alcohols, cleared in clove
oil, cedar-wood oil or xylol and embedded in a paraffin bath. The time required
for embedding in the paraffin bath will vary according to the thickness of the
specimens. After embedding, the specimen should be placed in melted paraffin
poured into a rectangle composed of two metal L-pieces. After orientating it
* For preparation see A. Bolles Lee's Microtomist's Vade mecum, revised by Gatenby & Cowdry
(1928).
26 THE INDIAN JOURNAL OF VETERINARY SCIENCE AND ANIMAL HUSBANDRY [ II, 1
properly, the paraffin should be cooled suddenly by immersion in cold water. The
paraffin, containing the specimen is easily extricated in the form of a block after
cooling. If required, the block may be trimmed and sections of it cut with a micro-
tome. They should then be floated in luke-warm water on a slide smeared with
Mayer's albumen (50 c.c. glycerine, 50 c.c. white of an egg and 1 gm. of sodium
salicylate mixed and filtered). When the sections have become completely flat, the
slide should be placed for 24 hours in an oven maintaining an even temperature of
40° C., when the water will have completely evaporated and the sections will be
found well fixed to the slide. The slide should then be immersed in a trough of
xylol in order to dissolve the whole of the paraffin. It should then be passed down
through absolute alcohol, 90, 70, 50, 30 per cent. alcohols and distilled water (ten
minutes in each medium being sufficient). The slide should then be placed in a
dilute solution of hæmatoxylin (of the same strength as was used for staining in
toto) for 24 hours, differentiated in acidulated water, dehydrated in graded alcohols,
cleared in xylol and mounted in canada balsam. If it is desired to have a double
stain, the slide after staining with hæmatoxylin as described above should be kept
for some time in eosin solution, after which it should be washed, dehydrated,
cleared and mounted as in the former case.
Clearing. It has already been remarked that the nematodes and acantho¬
cephala cannot be stained. Their anatomy can, however, be well elucidated if
they are cleared in some suitable medium. The clearing agents usually used are
beech-wood creosote, carbolic acid and lactophenol (40 c.c. glycerine, 20 c.c. carbolic
acid, 20 c.c. lactic acid and 20 c.c water). Of these three, lactophenol clears the
specimens to a moderate degree and is useful in some cases. Carbolic acid can be
used as it is, at higher temperatures only. In order to make it serviceable at lower
temperatures, it is usually mixed with a varying quantity of 70 per cent.alcohol.
Beech-wood creosote is a very good clearing agent, but if the specimens are kept
in it for a long time they are turned black. In order that the specimens may be
quickly cleared it is advisable to dehydrate them in 70 per cent.alcohol before
transferring them to a clearing medium. Spines and hooks in Platyhelminthes are
much more clearly observed after clearing the specimens in any one of the above
media.
Rapid examination of trematodes and cestodes. It will be seen from the above
that the processes of fixing, staining and mounting require at least a few days for
completion. The examination has therefore to be necessarily delayed by that
period. It is, however, sometimes desired to examine the specimens in a very
short period. For this purpose the " rapid method " recommended by Meggitt
[1924] may be followed. 97 parts of saturated solution of carmine in 45 per cent.
HELMINTH PARASITES AFFECTING DOMESTIC ANIMALS
27
acetic acid and 3 parts of saturated solution of ferric acetate in glacial acetic acid
are prepared immediately before use and mixed. The living specimens may be
transferred to this fluid and allowed to remain there from 5 to 30 minutes. They
may then be dehydrated in absolute alcohol for 5 minutes, cleared and mounted.
The whole process does not require more than 45 minutes even in the case of a
thick specimen.
Urine examination. Eggs of Schistosoma and Dioctophyme are discharged in
urine. In order to detect these, the urine is centrifuged and the sediment examined
on a slide under a microscope, when the characteristic eggs will be seen.
Fæces examination. It is a well known fact that the eggs of parasitic worms
harboured by the host are passed out with the fæces. By examining these the
kind of worms infesting the animal can be determined. The procedure adopted for
examining the eggs differs according as they are operculated and heavy or light.
(a) For operculated and heavy eggs. Dilute the fæces with 10 to 20 times
its volume of water. Pass the solution through a fine sieve, centri-
fuge the solution and examine the sediment.
(b) For lighter eggs. The method recommended by Sheather [1923] is
found very useful:—
1. Emulsify the fæces with water until the mobility approximates
to that of water.
2. Strain through wire gauze having 30 meshes in a linear inch.
3. Mix with an equal volume of sugar solution (sp. gr. 1,200).
4. Centrifuge for 2 to 3 minutes and allow the solution to stand for
10 minutes.
By this treatment all the eggs float on the surface of the mixture and can be
taken out by lowering a cover-slip on the top of the fluid by means of plastocene
attached to a glass rod. The cover-slip should be placed on a slide, and the eggs
can then be seen under a low power of a microscope.
Egg counts. For economic purposes it is necessary to count the number of eggs
per gram of fæces. For this Chandler's [1925] modification of Stoll's method is the
best. Three grams of fæces should be diluted with 87 c. c. of water. The mixture
should then be well shaken. 0.3 c. c. of the mixture should then be spread on a
slide and the number of eggs in it should be counted under a microscope. This
number multiplied by 100 gives the egg concentration per gram of fæces.
Preservation of fæces. It is sometimes found that the fæces examination has
to be deferred for some days and occasionally for some weeks. In such cases it is
28 THE INDIAN JOURNAL OF VETERINARY SCIENCE AND ANIMAL HUSBANDRY [ II, I
necessary to preserve the fæces for the examination. Southwell [1930] recommends
the following procedure for this :—
1. Make the fæces of the consistency of porridge by adding water.
2. Add to it 10 times its volume of 5 per cent. formalin after bringing it
to boiling point.
3. Allow to stand after shaking well.
4. Pour off the supernatant liquid and add 5 per cent. cold formalin.
In the above method 5 per cent. formalin may be substituted by 10 per cent.
formalin and water by normal saline solution.
Procedure for larvæ contained in blood.
Collection. Embryos of Filariidæ and Trichinella are present in the circula-
tory system of the host. In such cases the blood from a vein is mixed with twenty
times its volume of physiological salt solution and kept in a refrigerator.
Fixation. A drop of blood is spread on a slide and is quickly dried with a
spirit lamp flame. It can also be fixed by dipping the slide in steaming 70 per
cent.alcohol or for special purposes by dipping in a solution of mercuric chloride.
Fixation and hæmoglobinization can be done simultaneously by transferring the
slide to acetic alcohol (glacial acetic acid 1 part and absolute alcohol 3 parts).
Staining. The slide after fixation and hæmoglobinization should be kept in
a dilute solution of Giemsa's stain for half an hour or more until the desired
degree of staining is obtained. The dilution that gives good results is by addition
of a drop of the stain to a c.c. of distilled water. Hæmatoxylin should be used for
staining the sheath.
Vital staining. For determining the details of the anatomy of filarial larvæ
and for studiyng the larval forms of trematodes and ciliated hexacanth larvae of
Pseudophyllidea, vital staining is very important. For this dilute solutions of
neutral red, methylene blue, Giemsa or cresyl blue should be used.