OTHER SOURCES OF PATHOLOGIC MATERIAL.
1. =Autopsies on Animals.= In the case of the small animals used
ordinarily for laboratory purposes, such as the mouse, rat,
guinea-pig, rabbit, cat and dog, the animal is put upon its back and
fastened to the autopsy-board either by small nails driven through
the extremities or by slip-knots of string or rope passed over the
latter. Autopsy board and holders designed especially for the purpose
can be obtained from makers of laboratory apparatus. A main-incision
is made in the anterior or median line from the chin to the
genitalia, and the skin stripped back from the thorax on each side
to expose the ribs. The thoracic cavity is then opened by cutting
the ribs with the bone-shears or bone-forceps, and the sternum and
cartilages are removed. The neck, thoracic and abdominal organs may
then be removed _en masse_ and examined outside the body, or the
organs may be removed singly and examined in succession, following in
general the same methods of procedure as in the autopsy on the human
body, adapting the methods given above to differences in anatomic
structure and size. For the opening of the skull and spinal canal the
bone-forceps alone may be used, or in the case of larger animals the
saw may be needed. Anatomic considerations should govern the method
of opening the skull. Directions for the performance of autopsies on
inoculated animals will be found in textbooks on bacteriology; and
veterinary methods of autopsies on the larger domestic animals are
given in textbooks on veterinary pathology. In all cases of autopsies
on animals full protocols should be kept, following the general order
of the autopsy, altered to suit the individual case.
2. =Surgical Operation.= A very large part of the material obtained
for pathologic examination is removed by the surgeon for diagnostic
purposes. The question of the surgical technique employed may be left
to the surgeon, but as far as the pathologic aim is concerned certain
principles should be followed, if the object of the examination is to
be secured. Unfortunately these principles are not recognized by the
great majority of practitioners, and pieces of tissue to be examined
are taken at haphazard from the surface or from necrotic areas, to
be run through by the pathologist, only to find that no diagnosis
is possible, either because the portion of tissue removed did not
extend deeply enough or is wholly necrotic. Great care and judgment
should be exercised in the choice of the portions to be removed for
diagnosis. The part removed must be characteristic of the condition
present. It is necessary not only to ascertain the character of
the pathologic change but also the nature of the reaction in the
surrounding tissue. A neoplasm may show the histologic structure
of an adenoma, but at its periphery may be found infiltrating the
neighboring tissues as an adenocarcinoma. If the piece of tissue
for examination is removed from the central part or surface of the
tumor, an incorrect diagnosis may be given. This is especially
true in the case of rectal and uterine polypi, papillomata of the
mouth and penis, horny warts, etc. The rule to be followed in all
cases is that the excised portion must be at the boundary-line of
the neoplasm or morbid process, and extending across it so as to
include both pathologic and surrounding normal tissues. The cut
must be deep enough to extend into living tissue, and in the case
of epithelial surfaces to go below the basement membrane. It should
be made at right angles to the surface. Necrotic, softened, or
degenerating portions should be avoided, unless a portion of this is
removed in addition for the purpose of ascertaining the nature of
the degenerative changes present. The scraping away of superficial
scabs, exudates, etc., should never be practiced for purposes of
diagnosis. Time is saved if a satisfactory excision be made the first
time, and to secure this the tissue must be living, the cut must be
deep enough, and the portion removed must fully represent the nature
of the condition present. When organs are removed, as in the case of
the appendix, uterus, tubes, ovaries, mamma, etc., several portions
of tissue representing different structures of the organ should be
secured for the examination.
Tumors and other pathologic specimens received from the surgeon
should be fully described as to _size_, _form_, _weight_,
_consistence_, _color_, _relation to surrounding tissues_
(encapsulated, well-defined borders, growth by infiltration or
expansion, zone of inflammation, etc.), _character of cut-surface_
(color, moisture, translucency, smooth or elevated, homogeneous,
character of cell-scraping, evidences of structure, etc.).
Accompanying all pathologic material sent to the pathologist should
be concise and accurate notes giving the name, sex, age, nationality,
occupation and status of the patient, anything in the individual
or family history bearing upon the condition, the source of the
specimen, its exact location and relations, manner of growth and
character of operation. The pathologist should have full data upon
which to construct his diagnosis. A very common idea among surgeons
is that the specimen alone should be sufficient for the pathologist,
and that other data are not necessary for the formation of his
opinion. Many other considerations than the mere histologic picture
presented by a specimen enter into the formulation of a pathologic
diagnosis, if it is to bring to the aid of the surgeon all that a
pathologist’s knowledge and experience can give. This is particularly
true when the pathologist, as is usually the case, is asked to give a
prognosis. Both in hospital service and in private surgical practice
it is best to have printed history forms to be filled out and to be
sent to the pathologist with each specimen.
Another factor seriously interfering with the efficiency of the
pathologist’s work is the failure of the surgeon to see that the
material removed for diagnosis is properly taken care of before it
reaches the pathologist. Tissues removed for examination should
never be allowed to dry. They should not be exposed to the air, but
should either be placed at once in a fixing fluid or covered with
damp cloth. Curettings should be placed for a moment upon a pad of
gauze to remove the excess of blood, and the fragments of tissue
are then picked up and put into the fixing solution. When sent by
mail or express fresh tissues should be wrapped in damp cloth and
then in rubber cloth; or if the distance is great they should be put
into fixing fluids. A sufficient quantity of the latter should be
used, or decomposition may take place before the specimen reaches
the pathologist. All material for bacteriologic examination should
be removed under proper precautions, put into sterilized vessels,
properly sealed and sent to the pathologist under proper precautions.
PART II.
THE TREATMENT OF THE MATERIAL.
=INTRODUCTION.= The material obtained by autopsy, surgical operation,
curettage, excision, spontaneous discharge, animal experimentation,
etc., may be examined microscopically in the fresh state, or prepared
for microscopic examination by methods of fixation, hardening and
imbedding. The methods necessary for such histologic studies of
pathologic material are given in the following pages, arranged as
far as possible in their logical order. Only those methods are
given that, in the light of the writer’s experience, yield the best
results, from the standpoints of economy of time, labor, and expense,
and perfection of result. The number of histologic methods contained
in the literature is so great that it is out of the question for
the student or practical worker to try out all of them. To give all
of these methods would create confusion. I have attempted to avoid
this by giving in full detail only those which in our laboratory
experience have yielded the best results. So many methods represent
but slight variations of some original method, and in the great
majority of cases these variations add so little or nothing of value
to the original method that in such cases the latter alone is given
in full, with references only to the variations of the method. The
individual equation plays such a large part in the judgment of
laboratory methods that allowance has been made for this when certain
variations or alterations in original methods have been strongly
recommended by expert laboratory workers.
The purpose of the microscopic examination is the revealment
of pathologic changes too small to be recognized by the naked
eye, and the securing of a diagnosis that cannot be made
macroscopically, as well as the confirmation of diagnoses based upon
the gross appearances. Aside from these more immediate practical
considerations, the microscopic examination of tissues is concerned
with the solution of etiologic and pathologic problems, and the
extension of our knowledge of disease. The aim of pathologic
technique is the fixation of tissues for microscopic examination
in such a manner that all of the morphologic and chemical elements
and constituents of the tissue are perfectly preserved, so that
with differential staining methods they are all brought out with
sufficient contrast to be readily and correctly identified. In a
certain number of methods this ideal is attained, and to Weigert,
more than to any other worker in the field of pathologic technique,
are we indebted for such ideal methods.
The choice of methods will depend upon the source and nature of the
material, the object of the examination, the time-element and the
degree of responsibility involved. The cellular elements of all
pathologic fluids, secretions and excretions should be examined
in the fresh state as well as in fixed preparations. For the
demonstration of various chemical and morphologic features that are
lost or altered by processes of fixation and imbedding, and when a
rapid diagnosis is required, the examination of the material in its
fresh state or by the freezing method is indicated. When the freezing
method cannot be employed because of the changes in cells and tissue
produced by it, when very thin or serial sections are desired, when a
rapid diagnosis is not required, and when a very careful and minute
study is desired, with the application of various staining methods,
then the material should be fixed, hardened and imbedded and cut
upon a microtome. Whatever method is chosen, it must be borne in
mind, particularly in practical diagnostic work, that the portions
chosen for microscopic examination must represent the characteristic
anatomic structures of the tissue or organ, that living tissue be
included, that the pathologic condition be represented both in its
fully-developed state and at the transition-border between it and the
healthy tissue, and that when sections are cut the block or tissue
must be so oriented as to give the most comprehensive view of the
tissue and the pathologic process. To accomplish this fully it is
often necessary to make a number of blocks representing different
areas of the material, and to cut these in different planes.
CHAPTER XVII.
THE LABORATORY OUTFIT.
For practical diagnostic work or for pathologic research various
instruments and utensils are necessary, although the expense of
fitting up a working pathologic laboratory is not as great as it
is often thought to be. The most expensive item, as well as the
most important, is the =microscope=. This should be of the best
make, and should be carefully selected and tested before the final
purchase. As a rule the German makes, Zeiss, Leitz, and others,
are to be preferred to the American instruments, in spite of the
higher cost due to the duty imposed. I have found the German
microscopes uniformly good and standing the wear and tear of a
teaching laboratory much better than the American-made stands. I have
never seen a poor Zeiss or Leitz objective, but cannot say the same
thing of other makes. On the other hand, I have seen some American
objectives that were as good as any German ones, but there are not
many such. If one is going to buy an American microscope it should
be bought on the same principle that one would buy a violin or a
piano, wholly on its individual merits; and these can be ascertained
only by having the instrument carefully examined and tested by an
expert. Most laboratory workers will agree that the Zeiss instruments
are the best; they are also the most expensive. For all practical
purposes a Leitz stand costing ninety to one hundred dollars is
quite good enough. A medium-sized continental stand, with rack and
pinion and micrometer screw for coarse and fine adjustment, a triple
nose-piece with dust-protector, Abbé condenser, iris diaphragm,
plane and concave mirrors, three objectives (low, high and 1/12
oil-immersion), and two eye-pieces, a low and a high, form a complete
outfit that answers all practical requirements. The new type of
stand with curved arm and large stage, permitting the examination of
all parts of a Petri dish or glass plate, and with the mechanism of
the fine adjustment protected from any strain when the instrument
is lifted by the arm is especially recommended. I have also found
the black-finish very practical. The entire outfit need not be
purchased at once; the stand with its accessories and a low power
may first be purchased, and the higher-power objectives obtained
later. One of the first luxuries is a movable adjustable stage. A
very good and relatively cheap one is made by the Spencer Lens Co.,
of Buffalo. If a Zeiss stand is purchased the objectives A, D, and
1/12 oil-immersion, and oculars 2 and 4 best meet the requirements.
Of the Zeiss apochromatic series, the objectives 16.0, 8.0, 4.0, and
oil-immersion 2.0 mm., apert. 1.30, and oculars 4, 6 and 8 are most
serviceable. The apochromatic objectives and the compensation-oculars
are expensive, and need not be used for ordinary work, but are
indispensable for photographic work. The Leitz objectives, 3, 6, or
7, and 1/12 oil-immersion, with oculars 2 and 4, and the equivalent
objectives of the Spencer Lens Co. or Bausch and Lomb will answer
all ordinary needs. For the purpose of microscopic measurements an
ocular micrometer is necessary. This may be obtained as a separate
eye-piece, or as a round piece of glass with measured divisions
marked upon it that may be put into an ocular. The value of the scale
must be determined for every lens and tube-length by estimating the
number of its parts covering one part of a stage-micrometer marked in
hundredths of a millimetre.
[Illustration: FIG. 50.—A satisfactory outfit for the working
laboratory. Continental stand, medium-sized, with large stage, three
objectives, etc.]
An instrument of any one of the above-mentioned makes, carefully
selected and tested, should last its owner a life-time if proper
care is taken of it. It should receive the same careful attention
accorded a good violin or piano. It should be protected from dust,
action of chemicals, heat, sunlight, and rough usage. When carried
it should be supported in such a way that its weight is not thrown
upon the thread of the adjusting screws. The adjustment, draw-tube
and iris diaphragm should be carefully oiled at intervals, using the
least possible amount of the best microtome oil. It is not necessary
here to enter into the construction and theory of the microscope,
as this knowledge has usually been obtained before the pathologic
laboratory is reached. Experience has shown me, however, that it is
always necessary to remind students, even those experienced for some
time in the use of the microscope, of certain fundamental principles
in the adaptation of microscopic technique to pathologic work. The
following rules are of value:—
1. Use a _low-power_ objective for all work except for the study of
bacteria, microparasites and finer cell-structures. The aim should
be to obtain as much of a bird’s-eye view of the “geography” of
the section as possible. Contrast plays a very important part in
pathologic diagnosis; and it is lost in high-power work, so far as
the relations of cells and tissue-elements and pathologic products
are concerned. The student almost invariably enters the pathologic
laboratory with a fixed “high-power habit,” and he is usually greatly
surprised to learn how much he misses with the high-power and how
much he can see with the low-power. A motto used many years in my
laboratory, “Low-power objective and high-power cortex,” is of
greater educational value than may appear at first sight. A slide
should be examined first with the naked-eye, as it is held against
a window or light; then it should be examined under the low power.
Rarely will it be necessary to use a high-power except for the
purposes mentioned above.
2. Weak eye-pieces should be used; strong ones darken the field
and tire the eyes. Only in the case of apochromatic lenses and
compensation oculars can the strong ones be used without darkening
the field too much.
3. In the use of higher powers see that the tube is drawn out to
the proper length, as indicated in the directions sent with the
instrument.
4. For pathologic work an Abbé condenser is essential. It should be
pushed up to its proper position beneath the stage, and the plane
mirror should be used with it, reflecting the light from a cloud if
possible. Daylight is always the best light. When this cannot be
obtained an incandescent or Welsbach lamp with ground-glass globe
can be used. The concave mirror should then be used. The yellowish
tint of artificial light may be avoided by the use of a piece of blue
glass placed beneath the condenser, or a vessel of copper sulphate
solution may be interposed. Especial lamps designed to meet the
requirements are offered by the trade.
5. The iris-diaphragm should be adjusted by the same hand that moves
the slide, usually the left one. With unstained preparations the
diaphragm should be nearly closed; when using the oil-immersion it
should be fully opened, as is also the case when stained preparations
are studied with the lower power. With higher magnification the
aperture is diminished somewhat, although color-effects are best
shown with open diaphragm. In the study of pigments the diaphragm
should be fully closed for a few moments to see if the pigment
shows any color by reflected light. It is then examined by open
diaphragm. In the study of sections the mirror and diaphragm should
be manipulated in various ways to bring out all of the detail of the
preparation, and should be adjusted to suit each preparation.
6. Objectives must never be screwed down until they strike the slide
or stage. The higher-powers are frequently ruined in this way. When
running the objective down always examine from the side to see that
there is no danger of its striking the stage. In the use of the
oil-immersion place the drop of oil upon the slide or cover-glass,
and lower the objective by turning the coarse adjustment until the
oil spreads out between the lens and the glass; then focus with
the fine adjustment until a well-defined field is obtained. The
oil-immersion lens should not be allowed to stand many consecutive
hours in the oil. The oil should be cleaned from the lens by wiping
the latter with lens-paper or a soft cloth; if the lens is sticky the
paper or cloth may be moistened with benzol. The lens itself should
never be _wet_ with benzol, xylol, alcohol or any cleaning-fluid,
because of the danger of softening the balsam in which the lenses are
imbedded.
7. Use the mechanical stage only for differential blood-counting, or
when the entire section is to be gone over carefully, or when certain
details are found with difficulty and it is desirable to mark them
for future reference. An immense amount of time is lost in the use
of the mechanical stage for ordinary work. By moving the slide with
the fingers of the left hand resting upon the stage an entire section
may be gone over in a few seconds without missing any part of it;
to accomplish the same thing with the mechanical stage requires much
more time.
It is an excellent plan for the student to purchase his microscope
when entering the medical school and to use his own instrument
throughout his course. It is the one instrument without which no
physician can afford to enter practice; and the student who uses his
own microscope before graduation will continue to use it afterward.
The microscope obtained, the remaining expenditure necessary for
the fitting-up of a practical working laboratory of clinical and
pathologic diagnosis need not be very great if one’s financial
condition does not warrant spending with a free hand. It is possible
with a little labor and ingenuity to make at home, or to show the
local tinsmith how to make, a large part of the necessary apparatus,
such as sterilizers, paraffin-ovens, drying ovens, thermo-regulators,
etc., at a slight cost. Students of mine have made these things out
of old tin cans and glass tubing; one student at a cost of less than
three dollars constructed a microtome on which practical working
sections could be cut. For the celloidin method no apparatus except
the microtome is necessary, as the process of imbedding is carried
on in bottles or dishes. These points are mentioned to offset the
prevalent idea that a large expenditure is a necessity in installing
a practical working laboratory.
In a large diagnostic laboratory, or in one intended for teaching and
investigation, there are numerous accessories necessary to modern
microscopic technique. For the observation of living objects a _warm
stage_ is needed. The simple electrical apparatus devised by Ross is
the most convenient form, as it can be slipped on and off the slide
without changing the focus. It can be attached to any electric light
circuit and requires no attention.
For drawing from the microscope the improved form of the _camera
lucida_, or the latest model of the _Edinger drawing-apparatus_ are
recommended. Both of these instruments have recently been greatly
improved. The _Zeiss microphotographic apparatus_ is by far the
best for microphotographic work. For the _polarization-microscope_,
_microspectroscope_, and the complicated and expensive _ultra-violet_
and _dark-field_ apparatus the worker is referred to the Zeiss
catalogues. A simple and practical _dark-field_ method for the
illumination of bacteria, spirochætes and ultramicroscopic particles
suspended in fluids requires only a strong illumination and the use
of a _Zeiss_, _Leitz_, or _Reichert dark-field condenser_; or the
very simple “India-ink” method may be used for the demonstration of
spirochætes. (See Staining of Spirochætes.) Especial instruments
for easily finding a certain field are obtainable, and are of great
convenience in marking slides for photographic purposes.
[Illustration: FIG. 51.—A good practical microtome for paraffin and
celloidin work. Well-adapted to needs of students and beginners.]
Next to the microscope the most important instrument in pathologic
work is the =microtome=. One that can be used for either celloidin or
paraffin work, and that can also be utilized as a freezing microtome,
should be selected in private work, when economy is desired. The
majority can be used for either paraffin or celloidin, and one of the
Becker models can be easily attached to the carbonic acid holder for
the cutting of frozen material. The Bardeen freezing microtome is
relatively inexpensive, very satisfactory, and can be easily attached
to the carbonic acid tanks. It can be recommended for freezing work.
The “slide” type, either that in which the knife-holder is moved by
the hand directly or by a crank turned by the hand, is advised for
ordinary diagnostic work. For students and beginners the crank is of
great advantage, as the knife is securely held and cannot jump. The
best microtomes are made by Schanze of Leipzig, Jung of Heidelberg,
Becker of Göttingen, the Cambridge Scientific Co., the International
Instrument Co., and the Bausch & Lomb Co. For either celloidin or
paraffin work the medium or large Schanze slide-microtome or the
Minot’s precision microtome are recommended; for cutting serial
sections in paraffin the latest modification of the Minot automatic
rotary microtome is especially adapted. The best =microtome knives=
are made by Walb of Heidelberg. A long, heavy knife is to be
preferred to a light one. For the freezing microtome a knife of the
type of the blade of a carpenter’s plane set in a wooden handle
should be used, Hones and strops of the best quality are necessary.
=Paraffin-ovens= and =drying-ovens= of suitable size, and constructed
preferably of copper, are necessary for paraffin work. The
water-space about the oven should be sufficiently large, and the
temperature should be controlled by a thermo-regulator. Various
models are offered in the trade, but they can be made more cheaply
by the local tinsmith. The thermo-regulator can also be home-made by
anyone who has the necessary training in glass-blowing usually given
in courses in bacteriology.
=Various instruments= and =utensils=, such as _razors_,
_double-bladed knives_, _forceps_, _spatulas_, _section-lifters_,
_needles_, _scalpels_, _scissors_, _glass rods_ and _tubing_,
_test-tubes_, _graduates_, _flasks_, _funnels_, _bottles_,
_staining dishes_, _reagent bottles_, _rubber tubing_,
_water-bath_, _tripods_, _centrifuge_, _Bunsen burners_, _asbestos
pads_, _gauze_, _filter-papers_, _absorbent paper_, _labels_,
_oil_ and _wax colored pencils_, _slides_, _cover-glasses_,
_slide-boxes_, _camel’s-hair brushes_, _glass droppers_, _platinum
wire_, etc., are required for the pathologic laboratory, and can
be chosen to suit the individual needs. The solid watch glasses
make very good small staining dishes; the enamelled trays and
glass dishes used in photographic work are especially adapted
to the plate-method, particularly the size used for the 4 × 5
plate. Tea-strainers or small sieves can be used for staining
a large number of celloidin sections; and there are different
types of staining-dishes designed for the staining of slide- and
cover-glass preparations in number. Slides should be of the best
quality, colorless and with ground edges, and of medium thickness.
Cover-glasses should be square or oblong, round covers having but
little use in pathology. For ordinary work the No. 2 square, ¾
inch, is recommended; for work with the higher-power dry objectives
a thinner cover must be used. Slides and covers may be cleaned by
placing them in a solution of equal parts of one per cent sulphuric
and chromic acids and then rinsing in distilled water. A good
cleaning fluid is also made of one part acetic acid to three of
80 per cent alcohol. To clean old mounts melt the balsam by heat
or dissolve in turpentine, separate slides and covers, boil in
ten per cent lysol for half an hour, or for ten minutes in the
sulphuric-chromic-acid mixture, rinse thoroughly, dry with cloth
having no lint.
The laboratory should be supplied with running water, a sink
large enough for washing out specimens, numerous stop-cocks, and
a drip-board. Distilled water in abundance must be available. The
laboratory-table should have an alcohol- and xylol-proof finish,
black on the whole being the most practical color. A portion of the
table should be covered with glass beneath which there is laid a
sheet of white paper.
CHAPTER XVIII.
THE EXAMINATION OF FRESH MATERIAL.
I. METHODS OF EXAMINATION.
Pathologic material is examined in the fresh state when it is desired
to make a diagnosis in the shortest time possible, or when the
processes of fixation and hardening produce such alterations in the
morphology and chemic constitution of the cells that these features
can be recognized only in the unfixed, fresh state. So far as the
saving of time is concerned it is possible to take material removed
during an operation, examine it in the fresh state, and return a
diagnosis to the surgeon, while the patient is still on the table
under the influence of the anæsthetic. It is not possible to do this
with all tissues or with all pathologic conditions; but, when it can
be done, the advantages of such a rapid diagnosis, in the saving of
time, labor, expense and danger to the patient, are obvious. The best
idea of the cell is also gained by its study in a fresh condition.
Vital phenomena and certain morphologic features, as cilia, can be
observed only in fresh material. Many of the chemic constituents
of cells (glycogen, fat, mucin, pseudomucin, albumin-granules,
cholesterin, etc.) are either lost, or are so changed by processes
of fixation or hardening that they can no longer be recognized. The
majority of specific chemic tests can be made in fresh tissues only.
Moderate and slight degrees of fatty degeneration and cloudy swelling
are easily recognized in the fresh state; in fixed and hardened
preparations they may not be recognized at all. Particularly in the
case of the heart-muscle is it necessary to make an examination
of the fresh material when the diagnosis of these conditions is
concerned. Further, a greater or less degree of shrinking is caused
by many of the agents used in fixing and hardening, and this is
avoided by the examination of the fresh material. In the case of
pathologic fluids (sputum, urine, féces, etc.) an examination of
the sediment in the fresh state should always be made as a matter
of routine. The formed elements of these fluids are best determined
by this means. In the case of tissues, a diagnosis made by means of
scrapings, smears, teased bits of tissue, frozen sections, etc.,
should always be controlled by the examination of fixed and hardened
material.
In the examination of fresh material the following methods are
employed: _Sedimentation_, _smears_, _scraping_, _crushing_,
_teasing_, _maceration_, _sections_, _shaking or penciling_,
_digestion_, _intravital staining_, injection, _the warm stage_ _and_
“_cultivation_.”
1. =Sedimentation.= The formed elements of pathologic fluids (urine,
sputum, pus, blood, exudates, transudates, cyst-contents, etc.) are
examined by collecting the sediment of such fluids from the bottom
of a sedimenting glass or bottle, by means of a capillary pipette
controlled by the finger. While the sediment is passing up into the
tube the pipette should be moved about the bottom of the vessel so as
to get some of the sediment from all parts. When the fluid is rich in
cellular elements sedimentation is not necessary; a drop of the fluid
is placed upon the slide; if too thick it is diluted with physiologic
salt-solution or serum. If poor in cellular elements the fluid must
be centrifugalized by means of a water- or electric-centrifuge; and
a drop of the sediment in the centrifuge tube is then removed by the
pipette and placed upon the slide, and covered with a cover-glass.
To facilitate the low-power examination of such sediments parallel
streaks upon the slide may be made across its entire length, and
examined without the use of cover-glasses. To apply the various
reagents mentioned below it becomes necessary to use a cover-glass as
directed.
2. =Smears.= A clean fresh cut is made into the organ or tissue, and
a clean slide or cover-glass is drawn across the surface. Without
permitting the smear to dry a drop of salt-solution or any desired
reagent is put upon it, and it is then examined. This method is
especially applicable to the study of the cells of the spleen,
bone-marrow, lymphnodes, etc. Permanent balsam-mounts may be made
of such smears by fixing with heat or alcohol and ether, staining,
drying and mounting.
3. =Scraping.= A fresh cut is made into the organ or tissue, and
the excess of blood absorbed by a pad of absorbent paper devoid of
lint. A clean scalpel held at an angle of 45° is then drawn with
some force back and forth over the cut-surface until its blade
collects a sufficient amount of “tissue-juice” made up of the cells
of the tissue. This is then put upon a slide, and covered with
salt-solution or any desired reagent, and is then examined. This
method is used especially for cellular infiltrations, soft tumors,
and parenchymatous organs (spleen, lymphnodes, bone-marrow, liver,
etc.), and for the inner wall of cysts (echinococcus-cysts, cysts
lined with ciliated epithelium).
4. =Crushing.= A small bit of the tissue is cut out with the scissors
or scalpel, and placed upon the slide. A cover-glass is then placed
upon it and pressed down so firmly that the bit of tissue is spread
out in a thin film or layer beneath the cover-glass. Reagents are
introduced beneath the cover-glass, as desired. (See below.) This
method is used in the examination of the lung, kidneys and brain for
fatty embolism, and of the brain and spinal-cord for “fat-granule”
cells, pigment, calcified ganglion-cells, Negri bodies, etc. It is
also frequently used in the bacteriologic examination of tissues
(crushing of tubercles, etc.).
5. =Teasing.= A small bit of the tissue is cut out and placed
upon the slide or in a staining-dish and covered with physiologic
salt-solution. It is then teased with fine needles until divided
into its ultimate elements. Hard tumors (mature connective-tissue
tumors, fibrosarcomata, etc.), muscle (examination for trichina),
nerve-trunks, etc., are best examined in this way.
6. =Maceration.= In the case of some tissues the ultimate histologic
elements are so firmly held together that they cannot be separated
without the aid of a _macerating-_ or _dissociation-fluid_. (See
below.) =Digestion= is also used for the same end. The tissue should
be as fresh as possible and cut into small bits, which are placed
in the maceration-fluid in watch-glasses or staining-dishes for
twenty-four hours or longer. The macerated bits are then teased
until the finest elements are separated. In the case of very minute
elements the teasing may be carried on under a hand-lens or the
stereoscopic binocular microscope. During the process of maceration
all parts of the macerating tissue must be kept covered with the
macerating-fluid, or the uncovered portions will become hardened.
7. =Section-cutting.= Sections of fresh tissue may be made with
the _curved shears_, _simple razor_, _double-bladed razor_ and the
_freezing-microtome_.
_a._ _Curved Shears._ The tissue is put upon a stretch, and from the
surface a thin, flat section is cut out with the scissors. With care
a fairly thin section may be obtained in this way. It may be examined
by pressing it upon a slide beneath a cover-glass, or it may be
treated as a frozen section.
_b._ _Simple Razor._ As a part of their required laboratory training
medical students should learn how to make working sections with a
simple razor. Such a technical knowledge is sure to be of practical
use at some time or other. With a little practice sections
sufficiently thin for ordinary diagnostic work can be cut. If the
piece of tissue is large it may be held in the hand; but when small
or soft it may be placed in a matrix of hardened liver, pith,
potato, apple, firm lard or butter, paraffin, etc., and cut at the
same time with the latter. Both blade and tissue should be wet with
physiologic salt-solution. The blade, which must be very sharp,
should be _drawn_ through the tissue by a shoulder movement, with the
wrist-joint fixed. As the sections are cut they are floated off of
the razor-blade into physiologic salt-solution, and thence treated as
desired.
_c._ _Double-bladed Knife._ This consists of two parallel blades
arranged so that the space between them can be changed by means
of screws. For firm material the blades should be close together;
for soft material farther apart. The blades should be dipped into
physiologic salt-solution to fill up the space between the blades.
The instrument is then drawn through the tissue, and the section
between the blades floated out in salt-solution. Some workers use
this double-bladed knife to make a perpendicular cut into the tissue,
then turning it to either side to cut the lower edge of the section
and removing the blades with the section between them. Both the
single- and the double-bladed razors require practice for successful
section-cutting. Further, it must be borne in mind that sections of
fresh tissue obtained by these methods are unsuited for complicated
staining methods and can be used only for the simplest staining
processes. Nevertheless, in a fairly large proportion of pathologic
conditions it is possible to secure a diagnosis by these methods.
_d._ _Freezing Microtome._ Much more satisfactory sections can
be obtained by freezing the fresh tissue and cutting it upon the
=freezing microtome=. Various types of these instruments are in
the market. The freezing is accomplished by the use of _ether_,
_ethyl-chloride_ or _fluid carbonic acid gas_. The ether and
ethyl-chloride freezing-microtomes consist essentially of a metal
plate or hollow box on which the tissue rests and against the
under-surface of which a spray of ether or ethyl-chloride is forced
by means of a rubber bulb connected with a supply of the freezing
agent in a bottle. Such freezing attachments can be attached to any
microtome, but special instruments as Jung’s “student’s freezing
microtome,” Cathcart’s, Bausch and Lomb’s or Becker’s ether-freezing
microtome can be recommended for this purpose. (See Fig. 52.) The
tissue to be frozen must be of small size and not more than 3-4 mm.
thick. It is placed upon the freezing plate in a drop of water, white
of egg or thick gum-arabic, and pressed firmly against the plate. The
spray must not be too constant or strong, but should be given with
regular pauses of about a second to allow the ether to evaporate. The
tissue must be firmly frozen, but not so hard that it crumbles. It is
usually difficult to freeze the upper part of the tissue hard enough
to give good sections, and this half-frozen tissue must be trimmed
off with the knife until good sections are secured. As the sections
are cut they are removed from the knife-blade with the finger and
put into physiologic salt-solution that has been recently boiled to
drive off the air, so that, in thawing, there may be no formation
of air-bubbles in the tissue to cause artefacts. If put into strong
alcohol diffusion-currents may damage the tissue; a succession of
graded solutions should, therefore, be used if the sections are to be
fixed and hardened. The further treatment of frozen sections is given
below. The relative slowness of the method of freezing, its greater
cost, the necessity of the frequent replacement of the rubber bulbs
and tubing, and the greater amount of trouble required by the use of
ether or ethyl-chloride are disadvantages that can be avoided by the
use of the carbonic acid freezing microtome. Where much work is done
by the freezing method the use of the latter is advised.
[Illustration: FIG. 52.—Type of freezing-microtome, for the use of
ether or ethyl-chloride. Cathcart model.]
[Illustration: FIG. 53.—Carbonic-acid freezing-microtome. Becker
model.]
[Illustration: FIG. 54.—Bardeen freezing-microtome attached to
carbonic-acid gas-cylinder.]
The _carbonic acid outfit_ consists of a microtome arranged so that
it can be connected with a cylinder of compressed carbonic acid
gas, as shown in the Aschoff-Becker or Bardeen models. (Figs 53 and
54.) The cylinders or drums containing a charge of 15-30 lbs. of
the liquid carbonic-acid are furnished by the trade at reasonable
rates. It is not necessary to buy the drums, as they are replaced
by full ones as needed. They are provided with valves and can be
fastened in an upright position or laid flat upon a table, in any
way convenient for the attachment to the microtome. Connection is
made between the valve of the cylinder and the object-holder of the
microtome by means of strong rubber tubing which should be securely
wired at both ends, or by a flexible metallic tube. The latter is
preferable, as the rubber-tubing often bursts, or is so stretched
by the pressure of the gas that it must be frequently replaced.
Object-holders with flexible metallic tube attachment for use in an
ordinary microtome are supplied by Bausch and Lomb and other firms.
My personal experience makes me prefer the Bardeen freezing microtome
(see Fig. 54) to all others. It is cheap and can be easily attached
to the valve of the gas-cylinder, which is fastened horizontally
to the table-top, as it has a screw-thread fitted to the uniform
thread of the drum-valve. The object-holder can be raised or lowered
between the glass-tracks on which the knife runs, so that sections
of a definite thickness can be obtained. The most satisfactory knife
is that of the type of a plane-bit set in a wooden handle. (See Fig.
55.) It must be well honed and stropped. The tissue, which should
not be more than 5 mm. thick, is placed upon the object-holder in a
drop of water, albumin fixative, or saturated solution of gum-arabic,
and pressed firmly against the plate as the gas is turned on slowly
and evenly. Freezing is usually accomplished in one-half to one
minute. The best results are obtained by turning the gas on for about
15-20 seconds, then turning it off for several seconds; the tissue
will continue to freeze; if not hard enough the gas is turned on
again for 15 seconds and then turned off. If necessary this may be
repeated until the tissue is properly frozen. The interruption of the
gas-flow prevents over-freezing and thereby lessens the amount of
change in the cells. When sufficiently frozen the gas is then turned
off; the knife, with bevel-edge down, is held in the right hand with
its handle between the palm and the ball of the thumb with the back
of the hand uppermost; and the edge of the knife set at an angle of
45° to the tracks, along which it is rapidly pushed back and forth,
shaving the sections from the frozen tissue as the latter is pushed
up above the level of the tracks by means of the micrometer screw
turned with the left hand. The screw should have been adjusted at the
proper height before freezing, so that no time is lost in getting
the object-holder up to the height for cutting. By holding the elbow
and fore-arm closely against the body and pushing from the elbow
with wrist-joint fixed the shaving of sections can be accomplished
very quickly, so that with one freezing several hundred sections can
be obtained. The microtome-screw should not be turned by the left
hand until the knife on its return has cleared the tissue. A little
practice in co-ordinating movements is necessary for expert work. The
sections should be allowed to collect upon the knife-blade until a
large number have been cut; they are then swept off the blade by the
finger into cold freshly-boiled physiologic salt-solution. Sections
of tissue that have been previously fixed can be put into 60 per cent
alcohol, where the sections will unroll and straighten out perfectly
flat. When the method of freezing by alternately turning on and off
the gas, as given above, is followed, and the amount of freezing in
the interval noted, there is no danger of over-freezing, and the
breaking and crumbling of sections, with the production of marked
artefacts, is avoided. When over-freezing has occurred the block may
be partly thawed out by the finger. But _over-freezing_, as well as
_repeated freezing and thawing_, may cause so much damage to fresh
tissue that a diagnosis cannot be obtained from the frozen sections.
After thawing out in the physiologic salt-solution the sections
of fresh tissue obtained by freezing are treated according to the
methods given in the next section of this chapter.
[Illustration: FIG. 55.—Knife for Bardeen freezing-microtome.]
It must be emphasized here that the process of freezing is an
active one, and alters the relation of cell-structures. With many
fresh tissues the changes resulting from the freezing are so great
that no diagnosis can be made. It seems necessary here to warn
against the routine employment of the rapid method of freezing and
staining fresh tissues in the diagnosis of material obtained by
surgical operation. It has become a fad with some surgeons to make
a pathologic diagnosis by the freezing method while the patient is
on the table. Consequently, as the result of diagnoses made by the
rapid freezing and staining method, many mistakes are made, even
by supposed experts in this line. Particularly in the diagnosis of
sarcoma is it easy to make mistakes because of the altered aspect
of the cells caused by freezing. Normal lymphnodes, tonsils and
inflammatory infiltrations may look like spindle-cell sarcoma in
the sections prepared by the rapid freezing and staining method;
and the exact nature of many other pathologic conditions cannot
be accurately determined from such sections. On the other hand, a
certain number of pathologic conditions can always be recognized in
sections obtained in this way, and this fact justifies the employment
of the method when properly controlled. _In all cases in which the
pathologic condition is not clearly evident in sections obtained
by the rapid freezing and staining method no diagnosis should be
given._ In such cases the tissue should be fixed and then cut upon
the freezing-microtome, or imbedded in celloidin or paraffin and then
cut. Even when the patient is upon the table the tissue removed can
be put into a 10 per cent formol solution for a few minutes and then
frozen directly in gum without washing out the formalin. The longer
the time that can be used for this short preliminary fixation the
better the sections will be and the less the production of artefacts
by the freezing. The process of fixation can be hastened by warming
the fixing-fluid. I advise this short fixation before freezing in
all cases of operative diagnostic work when the diagnosis is wanted
as soon as possible. For all other work with the freezing microtome,
when the question of time is not so important, fixation with
formol for 12-24 hours should be carried out. This combination of
formol-fixation and the freezing method permits the early diagnosis
of autopsy and operation material, makes possible the demonstration
of fat and other substances altered or dissolved out by the imbedding
methods, and is a convenient way of selecting tissues, requiring more
complicated staining processes. (See also Page 239.) The further
treatment of sections of fresh tissue obtained by freezing will be
found in the second section of this chapter.
8. =Penciling or Shaking.= For the demonstration of the stroma or
reticulum either fresh or fixed sections may be placed upon a slide
in an abundance of fluid and gently penciled with a fine, blunt
camel’s hair brush until the fluid becomes cloudy. The cloudy fluid
is washed away and replaced by fresh as long as cells are given
off. The same results may be obtained by shaking the sections in a
test-tube until the cells are shaken out of the stroma. The removal
of the cells from the section is shown by its greater transparency.
One of the practical applications of these methods in diagnostic work
is the differentiation between alveolar round-cell sarcomata and
carcinomata.
9. =Digestion.= For the demonstration of stroma, parasites, etc., the
tissues may be digested with gastric or pancreatic ferments until the
required elements are freed. A freshly prepared pepsin in 0.2 per
cent HCl in the incubator for 3-5 hours will digest fibrin in fresh
clots. Sections of fixed tissues may be imbedded in paraffin, cut,
and digested on the slide with Grübler’s pancreatin according to the
method of Flint.
10. =Intravital and Supravital Staining.= Various methods have been
advised for the intravital staining of cell-granules. Intravenous or
intraperitoneal injections of methylene blue, alum carmine, neutral
red and other stains will produce intracellular granule-staining in
various organs of experimental animals. In the study of low forms
of animal life staining solutions may be injected, or the animal or
its parts may be examined in staining fluids. Human material can be
examined by this method immediately after being removed from the body
by operation or within 1-2 hours after death. (For details of these
methods see article on _“Färbungen, intravitale_,” _Encyklopädie der
mikroskopischen Technik_.)
11. =Injection.= Injections for the demonstration of blood-vessels,
lymphatics, ducts of glands, etc., are rarely used in pathologic
work. The organs to be injected must be fresh, warm from the
body, if possible. The vessels should be washed out by a freshly
filtered 8 per cent sodium nitrate or sodium sulphate solution,
followed by physiologic salt-solution. A cannula is introduced into
a main vessel, tightly secured, and then connected with a syringe
or a gravity injection-apparatus giving a constant pressure. The
injection-mass is then injected under a low pressure. In the case of
injections into lymphatic vessels the cannula should be introduced
into the largest lymphatics at the periphery of the blood-vessels
where larger lymph-vessels are more easily found. Injections are
made with either cold or warm solutions; the latter are preferable
but require that the organ to be injected be warmed by immersing
it in water at a temperature of 40°C. The injection fluid must
be of the same temperature. After the warm injection is given
the organ is put into ice-cold 10 per cent formol solution until
fixed and then after-hardened, imbedded and stained as desired.
After the use of cold injections the tissues are fixed in 10 per
cent formol, alcohol, or any other desired solution, and treated
according to the end sought. Nuclear and diffuse stains contrasting
with the color of the injection mass should be used. The process
of injection requires great care; the pressure must be carefully
regulated to prevent extravasations, and the injection-fluid must
be free from air-bubbles. The injection is continued until the
organ appears diffusely stained. Blood-vessels are fixed in their
natural blood-injection by such agents as formol and chromic acid,
so that stains acting upon the red blood cells cause the veins and
capillaries to appear as if they had been injected.
The following injection masses are advised:—
1. =Cold Injection Mass (Beale’s Glycerin Carmine):—=
Dissolve 0.3 grm. of carmine in a small quantity of water
containing 5 drops of ammonia; add 15 cc. of glycerin and shake;
add drop by drop 15 cc. of glycerin containing 8-10 drops of
glacial acetic acid. Then add further glycerin 15 cc., alcohol 8
cc., and water 24 cc.
2. =Warm Injection Mass (Thiersch’s Berlin-blue Gelatin):—=
(_a_) Dissolve 1 part of gelatin in 2 parts of water by allowing it
to soak 24 hours, and then warming. Filter through flannel.
(_b_) Saturated water solution of ferrous sulphate.
(_c_) Saturated water solution of red ferricyanide of potassium.
(_d_) Saturated water solution of oxalic acid.
Make a solution (_1_) by adding 30 cc. of _a_ to 12 cc. of _b_, and
a solution (_2_) by adding 24 cc. of _c_ to 60 cc. of _a_; both
mixtures to be made at a temperature of 30°C. At same temperature
add 24 cc. of _d_ to solution _2_, and then solution _1_, stirring
constantly so that Berlin blue is precipitated. Heat on a
water-bath to 90°C.; filter through flannel.
3. =Fischer’s Milk-Method.= The vessels are flushed with 8 per cent
sodium nitrate or sulphate solution, and then injected with milk.
When sufficiently injected the tissue is hardened for 24 hours in a
solution of water, 1,000 cc., formalin (40 per cent formaldehyde)
75 cc., and glacial acetic acid 15 cc. Freeze, cut and stain with
Sudan III or Scharlach R.; the course of the vessels is outlined by
the fat-globules.
4. =Silbermann’s= method of injecting indigo-carmine, eosin or
phlosin-red into the circulating blood has been used for the
demonstration of capillary thrombi, the latter remaining free from
the pigment.
12. =Warm Stage.= For the study of vital phenomena in the living
cell _Ross’s electrical warmer_ is recommended. It can be slipped on
and off the slide without changing the focus, and is managed without
any difficulty. It keeps the centre of the slide at a temperature of
37°C. Reagents can be applied as desired. Deetjen’s agar may be used
as a medium for the preservation of living cells. (See Methods of
Blood-Examination.) Various forms of warm and moist chambers used in
experimental embryological work can also be utilized in experimental
pathology.
13. =Tissue-cultivation.= The embryologic methods of growing tissues
in lymph and blood-plasma as developed by Harrison, Burrows and
Carrel have been applied in pathologic work to the experimental
study of repair and regeneration, grafting, transplantation and
tumor-transplantation. (For methods see _Harrison_, Journal of Exper.
Zoology, 1910; _Burrows_, Jour. of Amer. Med. Assoc., 1910; _Carrel_,
Jour. of Amer. Med. Assoc., 1910.)
II. REAGENTS USED IN THE EXAMINATION OF FRESH TISSUES.
In the examination of fresh tissue it is often desirable to use
certain reagents for the purpose of making chemical tests or to bring
out some structures more prominently than others. To introduce these
reagents beneath the cover-glass in such a way as to get the desired
effect without disturbing given fields requires some practice with a
very simple technical method. The preparation is first examined in
salt-solution, and the cover-glass adjusted so that it has a slight
rim of fluid about its edge, but not enough to make it float. The
reagent to be applied is dropped with a glass-dropper at one side
of the cover-glass, while at the other the salt-solution is removed
slowly by a piece of absorbent paper. The changes produced in the
tissue-elements during the progress of the reagent can be observed
under low or high powers. Care must be taken to change the fluids so
slowly that isolated cells will not be washed away.
1. =Physiologic or Indifferent Fluids.= Serous exudates,
blood-serum, hydrocele fluid, etc.; artificial serum made by a
mixture of 9 parts physiologic salt-solution with 1 part white of
egg; or physiologic salt-solution (0.9 per cent for warm-blooded
animals, 0.6 per cent for cold-blooded).
2. =Maceration Fluids.= 33 per cent alcohol (24 hours); chromic
acid 1:5000 (24 hours); potassium bichromate 0.1-0.2 per cent
solution (2-4 days for nervous tissue); 0.1 per cent osmic acid
(12-24 hours); 33 per cent potassium hydroxide (¼-1 hour, for
muscle, tissue must be examined in the solution, as the cells
dissolve when water is added); Arnold’s iodine solution (10 parts
of a 10 per cent potassium iodide solution to which are added 5-10
drops of a solution containing 5 grms. of iodine and 10 grms. of
potassium iodide in 100 cc. of water. Macerate one or more days.
If solution becomes discolored add more of the second solution);
very dilute formol solutions (1 cc. to 500 cc. physiologic
salt-solution); Müller’s fluid (2-3 days, good for nervous tissue).
3. =Glycerin.= Used without diluting as a _clearing agent_,
particularly when pigment is present; and as a _mounting medium_
for stained preparations that cannot be put into alcohol.
4. =Potassium Acetate.= Saturated water solution for clearing
and mounting fresh preparations. Does not clear as strongly as
glycerin, hence is better adapted for the examination of fresh
tissues.
5. =Acetic Acid.= 1-2-5 per cent solutions are usually employed.
Clears the protoplasm and causes the nucleus to shrink slightly
and to stand out more distinctly. It differentiates fatty and
albuminous granules, dissolving the latter; and is useful in the
demonstration of elastic tissue fibres, sharply outlining these
against the connective-tissue which swells and becomes clear.
6. =Acetic Acid Fuchsin.= A few drops of fuchsin are added to a 2
per cent solution of acetic acid. With this solution the nuclei are
not only brought out more sharply but they are stained red.
7. =Lugol’s Solution.= Dilute Lugol’s solution to a pale yellow
color. It brings out the contours of cell and nucleus, and has a
specific reaction with glycogen and amyloid, giving both a brown
color. Since glycogen is dissolved out in water-solutions tests
for glycogen in fresh tissues should be made with iodine-glycerin
or iodine-gum (Lugol’s one part, gum arabic 100 parts). Smears or
cover-glass preparations may be placed in covered dishes containing
a few crystals of iodine.
8. =Potassium and Sodium Hydroxides.= In solutions of 1-3 per cent
all tissue-structures swell and dissolve or become unrecognizable
except elastic fibres, fat, pigment, amyloid, bacteria, yeasts
and moulds. Used especially for examination of skin-scrapings or
pus for presence of blastomyces and various forms of parasitic
moulds (barber’s itch, ringworm, tinea, etc.). Solutions of 33 per
cent clear the tissues but do not destroy the cells. Useful for
maceration. When diluted the cells are destroyed.
9. =Mineral Acids.= HCl or H_{2}SO_{4} (3-5 per cent). Used to
dissolve areas of calcification. Calcium dissolves with liberation
of CO_{2}; the phosphates dissolve without gas-formation, but with
H_{2}SO_{4} form crystals of calcium sulphate. Sulphuric acid is
also used as a test for cholesterin (red or violet coloration), and
with iodine as a test for amyloid.
10. =Osmic Acid.= In a 1 per cent solution this is used to test for
the presence of fat (oleates), the fat-droplets become black or
brown.
11. =Sudan III or Scharlach R.= Alcoholic solutions of these dyes
are used for the demonstration of fat in fresh tissues. They stain
fat orange to scarlet. (For method see Staining of Fat.)
12. =Alcohol and Ether.= Used to dissolve fat-granules and to
differentiate between these and albuminous granules.
13. =Stains.= Fuchsin, methylene blue, methylene green in 1 per
cent solutions in physiologic salt-solution and acetic-acid-fuchsin
are the best stains used for the examination of fresh-tissues
beneath the cover-glass. They are drawn under the cover-glass
according to the method given above.
For the rapid staining of sections of fresh tissue cut by freezing
a section is floated from the salt-solution onto a slide, which is
then carefully lifted from the salt-solution and the excess of the
latter removed. Several drops of methylene blue, carbol-thionin
or carbol-kresyl-echt-violett are run upon the section with the
glass-dropper, and allowed to remain for 15-30 seconds. The stain
is then washed off with salt-solution, a cover-slip put on and the
section examined in the salt-solution. Thionin has been especially
recommended (Wood, Strouse and others) for the rapid staining of
frozen sections; but I prefer to use carbol-kresyl-echt-violett
(kresyl-echt-violett 1 grm., 5 per cent aqueous solution of phenol
80 cc., 95 per cent alcohol 20 cc.). This gives a very good
differentiation in the section examined in water, and the picture
is clearer than with thionin; the specific staining-reactions with
mucin, amyloid, mast-cells, etc., are also more marked than with
the latter stain. Such sections are not as clear as dehydrated
and cleared sections and their possibilities of diagnosis are
correspondingly limited, even in the hands of an expert with
sections of this kind.
To make permanent balsam-mounts of the frozen unfixed sections
the latter must be fixed in formol or alcohol, or by heat (hot
water). The sections may be placed in 4 per cent formol for
several minutes, then into 80 per cent alcohol, then stained,
washed, dehydrated in absolute alcohol, cleared in carbol-xylol
and finally mounted in balsam. Hæmatoxylin and eosin may be used
for the staining. Fresh sections may also be fixed in hot water,
stained in hæmatoxylin, dehydrated in alcohol and cleared in
xylol. To save time the sections may be fixed on the slide for a
few minutes in alcohol, care being taken to prevent the sections
from rolling up by dropping the alcohol onto the middle of the
section after it has been carefully flattened out on the slide.
The section may then be stained with hæmatoxylin, borax-carmine
or other stains, dehydrated, cleared and mounted in balsam. After
fixing with alcohol on the slide the section may be attached to the
slide by blotting it with absorbent paper, then covering section
with absolute alcohol, draining this off after a few seconds and
then running over the section a thin solution of celloidin, which
is allowed to drain off, leaving a very thin film over the section
and slide. The latter is then immersed in water for a few seconds,
and the celloidin-film on setting holds the section to the slide,
provided the celloidin has been of proper consistence. The section
can now be stained, washed, dehydrated, cleared in origanum and
mounted in balsam. When mounted the thin film of celloidin is
invisible (Wright’s method).
I have originated a much better method which is in use in my
laboratory, and can be applied to the staining of frozen sections
of fresh tissues in large numbers for class use. The sections are
floated from the salt-solution on to a warm solution of New Orleans
baking molasses diluted ten times, or a dilute sugar-dextrin
solution, and thence are floated on to a clean glass plate and
arranged in rows. The plate is drained, and then without drying is
immersed in absolute alcohol for 15-30 seconds; it is then flooded
with a thin celloidin, drained, the celloidin film allowed to set,
and the plate then put into warm water, where the celloidin sheet
floats off, carrying the sections, which can now be cut out and
treated as single celloidin sections, or the whole sheet can be
carried through the staining, dehydrating and clearing solutions to
be cut up into single sections before mounting.
CHAPTER XIX.
THE PRESERVATION OF MACROSCOPIC PREPARATIONS.
For preserving gross objects for museum specimens alcohol or formol
may be employed. The former bleaches the tissues so that ultimately
they are almost destitute of color. Formol in a 5 per cent solution
gives better color-effects than alcohol, as the blood-containing
parts remain darker. The fluid also remains clear and the tissues
are firm. When alcohol is used the fluid must be frequently changed,
as it becomes turbid and yellowish, and the tissues finally become
soft and lose their form. The best methods of preserving the natural
color are found in the various modifications of the =Kaiserling
method=. The organs or tissues are placed first in a formol solution
until they are just hardened, the formol changing the oxyhæmoglobin
into acid hæmatin. They are then transferred to alcohol to bring
back the natural color, which is accomplished by the change of the
acid hæmatin to an alkali hæmatin, which has a color very closely
resembling that of oxyhæmoglobin, so that the natural color is
approximately reproduced. The method is carried out as follows:—
Sol. I.—Formalin 200 cc.
Water 1,000 cc.
Potassium nitrate 15 grms.
Potassium acetate 30 grms.
The tissues are left in this solution, in the dark, for one to
several days, being watched carefully to see that they are not
over-hardened.
Sol. II.—80 per cent alcohol for 1-6 hours and then 95 per cent
until the color is fully restored (2-24 hours). Watch carefully and
remove as soon as best color effect is reached, and preserve in—
Sol. III.—Glycerin 400 cc.
Water 2,000 cc.
Potassium acetate 200 grms.
The specimens must be kept in air-tight jars, and crystals of
thymol added to prevent growth of moulds. This is sometimes very
difficult, and it becomes necessary to change the discolored fluid
for clear. I have found the rectangular museum jars best adapted
for the preservation of Kaiserling specimens. I use a wooden top
which fits over a thick piece of felt cut just the size of the
jar, which in turn fits over a piece of dental rubber cut to fit
the jar. The jar is placed upon a wooden bottom which has upright
steel rods at the corners, that pass through holes in the wooden
top, and have a screw-thread so that they can be fitted with
screws, which when screwed down hold the wooden top, felt and
rubber sheeting tightly in place, making the jar air-tight, but
giving a top easily removable. Very beautiful specimens can be
secured by the Kaiserling method, and they can be kept for several
years, but sooner or later the color-effect is lost. Light, heat
and exposure to the air cause a loss of color.
Some workers prefer the following in place of Sol. I:—
Hot water 2,000 cc.
Sodium sulphate 40 grms.
Magnesium sulphate 40 grms.
Sodium chloride 20 grms.
When salts are dissolved and solution cool add 200 cc. of formalin.
Melnikow-Raswedenkow Method:—
Sol. I.—Water 100 parts
Formol 10 parts
Sodium acetate 3 parts
Potassium chlorate 0.5 part
Leave in this 1-2-3-4-5 days, according to size of specimen. Large
organs must have solution injected into vessels.
Sol. II.—95 per cent alcohol, until color is restored.
Sol. III.—Preserve in: Water 100 parts
Glycerin 60 parts
Potassium acetate 30 parts
Pick’s Method:—
Sol. I.—Water 1,000 cc.
Formol 50 cc.
Carlsbad salts 50 grms.
Then transfer to 80 and 95 per cent alcohols, as for Kaiserling,
and preserve in water 9,000 cc., glycerin 5,400 cc., sodium acetate
2,700 grms.
=Westenhoeffer’s method= of preserving uric-acid: formol vapor 4-24
hours, then 80-90 per cent alcohol containing mercuric oxide, and
preserve in glycerin to which some mercuric oxide covered by cotton
or absorbent paper has been added.
=Claudius’s Method.= The specimen is placed on a grating in a
closed vessel containing a concentrated solution of ammonium
sulphate, an abundance of the crystals being left on the bottom of
the vessel. Carbonic acid or illuminating gas is passed through the
ammonium sulphate solution for 48-72 hours, and the specimens are
then preserved in the same solution. My experience with this method
has not been satisfactory.
=Gelatin method= of mounting Kaiserling preparations: Soak washed
Gold Label gelatin in distilled water for 12-24 hours. Take equal
parts of water-logged gelatin and glycerin and dissolve by heating
in a double boiler, stirring, for 15-20 minutes. Cool to 40°C.
Then clarify with white of egg (well-whipped whites of three eggs
to half a gallon of jelly: stir well; steam for ½ hour) and filter
through cotton-wool. Add to jelly a few drops of a weak aqueous
solution of crystal violet to remove yellow color (_Bruère and
Kaufmann_). To prevent growth of bacteria a small percentage of
formol or crystal of phenol may be added. Kaiserling specimens are
placed in glass dishes in the melted jelly, and covered with it.
When set the dishes containing the mounted specimens may be covered
with glass-plates and fastened to these by balsam or cement. I use
a deep Petri dish, filling it about two-thirds full with the jelly;
over this I pour melted paraffin of a very low melting point so as
not to melt the jelly. When the layer of soft paraffin is hard, a
thick layer of paraffin of a 52° melting-point is poured over it,
and the dish filled even. When the hard paraffin is set, it is
varnished with shellac. Liquefaction of the gelatin by bacteria or
enzymes constitutes the great drawback to this method.
CHAPTER XX.
THE FIXATION AND HARDENING OF TISSUES.
=GENERAL CONSIDERATIONS.= For the examination of material that may
be injured by freezing, or when very thin sections are required for
complicated staining procedures, it becomes necessary to prepare the
tissue by _fixation_ and _hardening_, so that it can be _imbedded_
in some medium permitting the _cutting_ on the microtome of as thin
sections as may be desired. Fixation is that process by which the
appearances of the tissue are preserved as they were when it was
taken for examination; hence in order to obtain pictures resembling
as closely as possible those of the living tissue the material
should be _fixed_ immediately upon its removal from the living body
by operation, or as soon as possible after death when obtained by
autopsy. _Fixing agents_ act by coagulating the cell albumins, in
this way “setting” or “fixing” the constituents of the cell so that
further change is stopped. Fixation, therefore, _hardens_ the cell,
and all fixatives are also hardening agents. A practical distinction
between fixing and hardening is made, however, resting upon the fact
that not all fixatives harden the tissue so completely that the
proper consistence for the cutting of thin sections is attained. To
achieve this the tissue must be dehydrated. Alcohol and acetone are
the only reagents fixing and hardening perfectly at the same time, as
they remove the water from the tissue; for all other fixing agents
an _after-hardening_ in alcohol is necessary. In the case of such
reagents the division of the process into a _primary fixation_ and
a _second hardening_ stage has been the cause of the divergence in
meaning of the two terms.
The best fixing agents are those that kill the cells at once, but
cause a slow coagulation with little or no shrinking. They must
penetrate and diffuse through the tissues rapidly so that the deepest
cells are quickly reached. Acid media, especially those containing
small percentages of acetic acid, are therefore better than alkaline
solutions. The tissue-elements, particularly the nuclei, must
be preserved as perfectly as possible so that they will not be
affected by further procedures of microscopic technique. The chemic
properties of physiologic and pathologic substances must likewise be
preserved. The preservation of karyokinetic figures is a criterion
of good fixation. In pathologic work it is also desirable that the
fixing agent should preserve the red blood cells, and permit of the
staining of bacteria in sections. Since fixing media are more or less
selective in their action, it follows that there is no one fixative
that gives equally good results in all cases. Especial fixing
reagents must be used for the demonstration of certain substances
(fat, etc.), or for the use of certain staining methods. Some
stains cannot be used at all after certain fixing agents have been
employed. For general use that fixing agent having the widest range
of usefulness should be employed; and for this reason fixing media
composed of several fixing agents are often employed in preference to
the use of a single one.
GENERAL RULES FOR FIXING AND HARDENING.
The tissue should be put into the fixing fluid as soon as possible
after its removal from the body. It must not be allowed to dry. There
should be an abundance of the fixing solution, 25-50 times the volume
of the object to be fixed. The tissue should never be put into a
dry vessel and the fixing fluid poured upon it; the vessel should
first be filled with the fixing solution and the tissue then dropped
into the latter. A slight agitation of the fluid will prevent the
sticking of the tissue to the bottom or sides of the vessel. The
size of the pieces must be adapted to the penetrating power of the
fixing reagent, but as a rule the pieces should not be more than 2-3
cms. in thickness, and for some reagents 0.5 cm. is as thick as they
can safely be. The reagents used should be changed when they become
cloudy or discolored. The used solution may be filtered and used
again, but some reagents can be used but once. Alcohol may be saved
for redistillation. In the case of some reagents (mercuric chloride,
chromic acid, osmic acid, etc.), the time limits of the fixation
should not be exceeded, as over-fixation will ruin the staining-power
of the tissues. Alcohol is practically the only solution in which
tissues may be left indefinitely, but even with it there are certain
limitations. As a general rule the time required for fixation may
be shortened by keeping the reagents at incubator-temperature. As
the different fixing reagents vary so greatly as to their especial
advantages and disadvantages these will be considered separately.
Only the best and most commonly used methods are here included.
1. =ACETONE.= A water-free acetone is employed by placing pure
dried white copper sulphate in the bottom of the bottle or vessel
in which the fixation-process is carried on. Several layers
of filter-paper are put over the copper sulphate to keep the
tissues from touching it, and the acetone is then poured into the
vessel. As soon as the copper-sulphate becomes blue it must be
again fused. It is only by this method of constant dehydration
that acetone can be employed to any advantage as a fixing agent;
if fused copper sulphate is not employed the amount of acetone
necessary to fix well is so great that the method becomes too
troublesome and expensive to be recommended. But with the simple
copper-sulphate method of constant dehydration acetone becomes the
cheapest, most rapid and one of the best fixing reagents. The use
of alcohol is avoided, and the period of infiltration in xylol
and in paraffin shortened. For very quick work the entire process
of fixation, hardening and dehydration may be achieved by the use
of acetone alone, small pieces of tissue being fixed ½-2 hours
in acetone and then transferred directly to soft paraffin. For
ordinary work pieces of tissue 0.5 cm. thick are put into acetone
over fused copper sulphate for 20-60 minutes. A judgment of the
degree of fixation can be obtained by pressing the tissue lightly
between the fingers; if it is of uniform consistence, and does
not give as if the inner portions were softer than the surface,
the fixation is complete. From the acetone the tissues may be
brought directly into xylol for 5-10 minutes, until they obtain a
cloudy transparency, thence into paraffin for 15-30 minutes and
then blocked. The whole process of fixation, imbedding, cutting
and staining can be carried out in 30 minutes. Acetone may be
combined with formol, alcohol or any of the other fixing agents,
but when it is desired to use any one of these for some especial
purpose it is better to fix first with the desired reagent and
then to use acetone for the dehydration-process alone, instead of
alcohol. We have found that formol-fixation followed by acetone
dehydration gives excellent results for general pathologic work.
=Formol-acetone= (acetone 100 cc., formol 10 cc.) may be found to
have advantages. The quick fixation in water-free acetone causes
less contraction than fixation with absolute alcohol; fixation with
graded acetone-solutions causes practically none.
=Advantages.= It is the cheapest and quickest method. It penetrates
well and causes little contraction of the tissues, shortens the
time in xylol and paraffin and makes more easy the cutting of dense
fibrous structures. Cell-division figures are as well preserved as
by alcohol fixation, and the staining of bacteria in the tissues
can be carried out as well after acetone-fixation as after alcohol.
It preserves lecithin, hence can be used for the fixation of
nerve-tissues. When combined with formol (formol-acetone) the red
blood-cells are well-preserved and take a brilliant eosin stain.
=Disadvantages.= The disadvantages are practically the same as with
alcohol fixation but not so marked. Fat is dissolved, cell-division
figures are not so well-preserved as with mercuric chloride and
Flemming’s solution, and the blood-cells not so well preserved as
with formol and mercuric chloride.
2. =ALCOHOL.= Absolute, 95-96 per cent alcohol, or graded alcohols
(70, 95 per cent. and absolute) may be used for fixation. For
this purpose the stronger alcohols are preferable, as the weaker
solutions do not fix quickly enough. On the whole the use of 95-96
per cent is to be advised. The pieces of tissue must not be thick.
Plenty of alcohol should be used and it should be changed several
times during the process of fixation, which for larger pieces
requires several days. For the last change absolute should be used.
Very small bits such as uterine curettings can be fixed in one
hour, by using three changes of absolute alcohol. Since alcohol
both fixes and hardens it has been generally used, but it is a
relatively poor fixative. For after-hardening it is indispensable.
Absolute alcohol may be made from 96 per cent by the use of fused
copper sulphate. To test the strength of alcohol mix a few drops
with pure water-free xylol; if no sediment appears when viewed
against a dark background the alcohol is absolute or practically
so. Many of the disadvantages of alcohol fixation can be obviated
by the use of =formol-alcohol= (95 per cent alcohol 100 cc.,
formalin 10 cc.).
=Advantages.= Cheapness, quickness, and ease of method. Can be used
for quick diagnostic work. Penetrates well, and can be used for
large pieces of tissue. Preserves glycogen, is especially good for
the staining of bacteria in sections, and the majority of stains
work well with alcohol-fixation.
=Disadvantages.= Causes much shrinking and loss of finer details;
preserves cell-division figures not at all or poorly; destroys
the red blood cells; dissolves fat and other chemic products:
does not permit of the use of certain specific staining methods
(nerve-tissues); causes excessive hardness of fibrous and elastic
tissues and makes cutting difficult.
3. =CHROMIC ACID AND SALTS.= Chromic acid is rarely used alone in
pathologic work, but is a constituent of Flemming’s solution (see
below). Its salts are employed in the form of:—
A. =Müller’s Fluid= (Potassium bichromate 25.0 grms., sodium
sulphate 10.0 grms., water 1,000.0 cc.) This was formerly the
favorite fixing solution, but is now used chiefly for the eye
and nervous tissues, either alone, or after formol fixation,
or mixed with formol. Large pieces may be used, even an entire
brain, but the process requires months or even a year for the
best results. Even small pieces take several weeks. The process
may be hastened in the incubator. The solution should be changed
whenever it becomes cloudy. When fixation is complete the fixed and
hardened tissue may be cut directly on the freezing-microtome or
after-hardened and dehydrated in alcohol or acetone when it is to
be imbedded. The dehydration in alcohol should be carried out in
the dark and without previous washing of the tissue in the case of
nervous tissue. Greenish or brownish chrome precipitates appear if
the dehydration takes place in the light; but for ordinary material
this precipitate can be avoided by washing the fixed material in
running water for 24 hours before transferring to alcohol. Moulds
grow luxuriantly in Müller’s fluid, but may be inhibited by the use
of pieces of camphor, thymol, etc.
=Advantages.= Cheap, penetrates well, causes little shrinking,
permits special nerve-stains, preserves red blood-cells, gives
beautiful results with ordinary stains, preserves fat.
=Disadvantages.= Slowness; does not preserve division-figures;
does not permit of staining for bacteria; does not give good
results with many special staining methods (fibrin, elastic tissue,
reticulum, etc.).
B. =Erlitzky’s Fluid= (Potassium bichromate 25.0 grms., copper
sulphate 5.0 grms., water 1,000.0 cc.). Used for fixation of
nervous tissue. The formation of pigment precipitates may lead to
misinterpretation: the artefacts may be removed by hot water or
dilute acetic acid.
C. =Orth’s Fluid= (Müller’s fluid 100 cc., formol 10 cc. Make
fresh before using, as the mixture precipitates on standing.).
Fix for 3-12 hours in the incubator, or for 24-48 hours at room
temperature. Wash in running water for 12-24 hours; cut on the
freezing microtome, or after-harden in acetone or alcohol and imbed.
=Advantages.= Combines the good features of Müller’s and formol
fixations, and obviates some of the disadvantages. It is a good
general fixing solution.
3. =FORMOL OR FORMALIN= (40 per cent solution of formaldehyde
gas). Used in a _ten per cent solution_ (water or physiologic salt
solution nine parts, formalin one part), often incorrectly called 4
per cent formol or formalin, the mistake arising from the confusion
with 4 per cent formaldehyde gas. For nerve-tissues it is better to
dilute the formol with physiologic salt-solution than with water.
Fix for 3-4-12 hours according to size of tissue. As it penetrates
well large pieces can be used. Wash in water before after-hardening
in alcohol, if tissue is to be employed for general work, otherwise
it can be transferred directly to the alcohol without washing.
Tissues can be kept in formol for some weeks, but after that time
the staining-power is slowly affected, and the finer structures
suffer.
=Advantages.= Probably the best fixing reagent for general
pathologic work. It is cheap; easily made and kept in solutions
of proper strength for fixing; hardens while it fixes; does not
require after-washing; penetrates well; causes little shrinking;
permits freezing directly from the fixing solution; preserves fat;
permits the use of after-hardening with bichromate solutions for
especial nerve-stains: preserves the red blood-cells: gives good
results with nearly all stains, and differentiates bile-pigment
from hæmatoidin. It is the best fixing reagent when tissues are to
be sent some distance, as over-fixation occurs only after several
weeks or even months.
=Disadvantages.= Affects many people unpleasantly, causing coryza,
eczema of the hands and arms, and affections of the finger-nails,
so that workers having acquired this idiosyncrasy cannot expose
themselves to formol vapor; it dissolves glycogen and uric acid;
does not fix cell-division figures as well as mercuric chloride
or Flemming’s solution; causes the precipitation of diffuse
hæmoglobin in the form of brown or black pigment-granules that
may be taken for melanin, malaria pigment or hæmosiderin; causes
a pseudo-ochronosis of cartilages; and, unless thoroughly washed
from the tissues before after-hardening in alcohol, it makes
carmine-staining difficult or unsatisfactory and affects also the
specific staining-reactions for amyloid and mucin; it is not as
good as alcohol or mercuric-chloride when the sections are to be
stained for bacteria. In spite of these disadvantages it can be
recommended as the best general fixing reagent.
For =Orth’s fluid=, =formol-acetone= and =formol-alcohol= see above.
4. =FREEZING AND DRYING.= The fresh tissue is frozen and dehydrated
in a vacuum over sulphuric acid, at a temperature of 20-30°C.; when
completely dried it is imbedded directly in paraffin. This method
has been especially recommended by Altmann on the ground that the
tissues are simply deprived of water without any change in volume.
5. =HEAT.= Physiologic salt-solution is heated to 80°C. Thin pieces
of tissue are placed in the hot water for two minutes, and then
after-hardened in alcohol. For larger pieces of œdematous tissues,
cysts, etc., that cannot be cut into thin pieces, the salt-solution
should be brought to 100°C and the tissues boiled for several
minutes. This method is advised particularly for the coagulation of
albumin in cysts, œdematous tissues, for the study of renal casts,
etc.
6. =MERCURIC CHLORIDE.= This is used most commonly in the form
of =a concentrated water solution= (mercuric chloride 7.5 grms.,
sodium chloride 0.5 grm., glacial acetic acid 5 cc., water 100.0
cc.), or as =Zenker’s solution= (mercuric chloride 5.0 grms.,
sodium sulphate 1.0 grm., potassium bichromate 2.5 grms., water 100
cc.; dissolve by heating, add 5 cc. glacial acetic acid just before
using. The use of a 5 per cent formol solution instead of acetic
acid is recommended). The pieces of tissue should not be thicker
than 5 mm. Fix 6-24 hours, then wash 24 hours in running water,
and after-harden in alcohol. Should the sections show mercuric
precipitates they should be treated with Lugol’s solution for 30-60
minutes, then washed in a dilute solution of lithium carbonate and
thoroughly washed out in water and alcohol. Much better stains can
be obtained by this treatment of the sections with Lugol’s. The
use of iodine in the alcohol during the process of after-hardening
is not advisable because of the action of the iodine upon the
albuminates of mercury. A 5 per cent solution of =sublamine= in
distilled water has recently been recommended by _Klingmüller_
and _Veiel_. Fix 1-3 hours, wash and after-harden in alcohol.
Precipitates are not formed, and good staining-results are obtained.
=Advantages.= The mercuric chloride solutions preserve well the red
blood-cells, mitotic figures and finer details of cell-structure,
and permit the staining of bacteria and animal parasites in
the sections. Certain especial staining methods (Mallory’s
reticulum-stain, etc.) can be used only after mercuric chloride
fixation, while many others (Heidenhain’s iron-hæmatoxylin,
Biondi-Heidenhain triple stain, etc.) give best results after
this fixation. For ordinary work the saturated mercuric chloride
solution is preferable to Zenker’s, as the latter does not give
good results with the commonly-used hæmatoxylin stains.
=Disadvantages.= More troublesome and expensive; require thorough
washing and subsequent removal of precipitates, and affect
(Zenker’s particularly) certain stains.
7. =OSMIC ACID.= Osmic acid is used alone in a 1 per cent solution,
or in such combinations as =Flemming’s solution= (chromic acid 1
per cent. sol. 15 cc., 1 per cent osmic acid 4 cc., glacial acetic
acid 1 cc.), =Hermann’s solution= (same as Flemming’s, with 15
cc. of a 1 per cent platinic chloride substituted for the chromic
acid), =Altmann’s solution= (5 per cent potassium bichromate
solution 50 cc., 2 per cent osmic acid solution 50 cc.), =Marchi’s
solution= (Müller’s fluid 2 parts, 1 per cent osmic acid solution
1 part), and that of =Pianese= (1 per cent sodium-chloroplatinate
15 cc., 2 per cent osmic acid 5 cc., ¼ per cent chromic acid
5 cc., formic acid 1 drop). The pieces of tissue must be very
thin, as osmic acid penetrates very slightly. Fix 6-24 hours in
the dark, and wash thoroughly in running water, and after-harden
in graded alcohols. These solutions have but limited use in
pathology, and are used chiefly for the study of fat (oleates) and
mitotic figures. Flemming’s and Hermann’s solutions are the best
for the study of mitotic figures, the latter bringing out plasma
details more clearly. Marchi’s fluid is used especially for the
study of nerve-degeneration, and Altmann’s for the demonstration
of Altmann’s granules. The method of Pianese is used for the
demonstration of cell-inclusions. The osmic-acid mixtures are all
expensive, penetrate poorly, cause precipitates, and affect greatly
the staining-power of the tissues, so that it becomes necessary
to use certain stains (safranin, carbol fuchsin, aniline gentian
violet, etc.) as counterstains.
8. =PICRIC ACID.= A saturated water solution of picric acid is
usually employed. Fix 12-24 hours, and wash in alcohol, not water,
and after-harden in graded alcohols. Preserves mitotic figures,
fine details of cell-structure, and is very good for bone and
calcified tissues, as it decalcifies and fixes at the same time.
Numerous modifications and combinations of the above methods
have been proposed such as _Flemming’s chrom-acetic solution_,
_Rawitz’s chromic-picric-nitric fluid_, _Rabl’s chrom-formic
mixture_, _Burckhardt’s chrom-osmic-nitric solution_, _Merkel’s
fluid (chromic acid-platinic chloride)_, _Carnoy’s mixture (glacial
acetic acid 1_, _absolute alcohol 6_, _chloroform 3_), and many
others. They have a limited use in pathologic work.
In the judgment of a section as to its fixation the following points
may be of service: in alcohol fixation the red blood cells are
haemolyzed, and there is much shrinking; with formol fixation the red
cells stain copper-red with eosin; in mercuric chloride fixations the
red cells stain rose-red with eosin, and pigment precipitates are
present; in bichromate fixation the red cells preserve their natural
color, and fat cells show a brownish color; osmic-acid fixation is
shown by the black color of the oleates, and the failure of the
tissue to stain by ordinary stains.
CHAPTER XXI.
DECALCIFICATION.
Bone and tissue containing deposits of lime must be decalcified
before they can be sectioned on the microtome. The decalcification
should be carried out after fixation and before the after-hardening
in alcohol. Some reagents may combine decalcification with fixation,
but this is satisfactory only when the amount of lime-salts is
relatively small. Fresh tissues should not be put into any of the
stronger acid decalcifying fluids, as they alter unfixed cells so
that the staining-power is lost and the fine histologic details
destroyed. The fixed tissue cut into small pieces is put into the
decalcifying reagent, which is used in large amount and must be
frequently changed. It is left in the decalcifying fluid until the
calcium salts are removed, as shown by tests with needle or scalpel.
The tissue must not be left in the fluid after decalcification is
attained, as the staining-power is affected by all decalcifying
reagents; it is therefore necessary to make frequent tests in
order to judge of the progress of the decalcifying process. After
decalcification the tissue should be washed in running water for 24
hours, and then after-hardened in alcohol. Alkaline solutions may
be used to remove the acid before washing. Sections of decalcified
tissue always stain slowly, and it is advisable to remove any acid
remaining in the tissues by soaking the sections in a saturated water
solution of lithium carbonate before staining. Numerous formulæ for
decalcifying fluids have been recommended; a few of the best methods
only are given here.
1. =Combined Fixation and Decalcification.= Picric acid or Müller’s
fluid may be used for this purpose when the amount of lime-salts
contained in the tissues is very small. The process is slow.
2. =Trichloracetic Acid.= Fix tissues in 10% formalin and decalcify
in trichloracetic acid 90 cc., formol (40 per cent formaldehyde) 10
cc. Change frequently. Decalcification is rapid, the tissue is but
little changed and the staining-power not affected.
3. =Concentrated Sulphurous Acid.= Fix in 10 per cent formol;
decalcify in concentrated sulphurous acid for 24 hours or longer if
necessary. Wash thoroughly in alkaline water. This is a very good and
rapid method; the staining-power is but little affected.
4. =Haug’s Solution.= (Pure nitric acid 3-9 cc., absolute alcohol 70
cc., sodium chloride 0.25 grm., water 30 cc.). For tissues fixed in
mercuric chloride.
5. =Phloroglucin.= (Phloroglucin 1 grm., pure nitric acid 10 cc.,
distilled water 50 cc.). The solution must be carefully dissolved
over the flame in a hood. Decalcification is rapid, and the tissue is
protected from the acid by the phloroglucin.
6. =Ebner’s Fluid.= (Hydrochloric acid 5 cc., sodium sulphate 5
grms., alcohol 500 cc., water 1,000 cc.).
7. =Schaffer’s Method.= Imbed the fixed and hardened tissue in
celloidin, harden the celloidin preparation in 85 per cent alcohol,
then place celloidin block in a 3-5 per cent water solution of pure
nitric acid and agitate in Thoma’s water wheel, for 12 hours, or
longer according to the size of the piece. Transfer block to a 5
per cent solution of lithium carbonate or sodium sulphate for 12-24
hours, changing solution several times, wash in running water for 48
hours, dehydrate in graded alcohols up to 85 per cent, and cut.
CHAPTER XXII.
IMBEDDING.
The most perfect methods of fixation and hardening do not permit
the cutting of fine sections on a microtome without the freezing of
the tissue, or its =infiltration= and =imbedding= in some substance
which surrounds it with a protective coating, and preserves and
holds together its structural elements in their relative positions.
For the cutting of very thin sections, or for the preparation of
serial sections, it is absolutely necessary to employ the process of
imbedding. At the present time =paraffin= and =celloidin= are the two
substances in general use for this purpose. While each one of these
possesses certain advantages over the other, and we find consequently
one laboratory worker preferring celloidin and another paraffin for
general work, a long and varied experience makes me believe that for
a teaching laboratory and for diagnostic work when much material is
examined, the paraffin method answers all purposes much better than
the celloidin, and that the latter need be employed only in very
exceptional cases. Since paraffin sections can be transferred into
celloidin by the molasses- or dextrin-fixative method, thus enabling
the use of staining-methods that require celloidin sections, very few
advantages are left in the favor of celloidin as an imbedding agent.
The paraffin method requires a more expensive outfit to start with in
the form of a paraffin oven and thermo-regulator, but otherwise the
two methods cost about the same. The paraffin method requires more
careful attention than the celloidin. As a rule thinner sections can
be obtained in paraffin than in celloidin, and for the preparation
of serial sections the paraffin method is the only method. Paraffin
blocks can be labeled and filed away, and kept indefinitely without
any loss of staining-power. With careful attention paid to the
different steps of the imbedding process practically everything that
can be cut in celloidin can be cut in paraffin. For very large pieces
a slow imbedding in celloidin is, however, preferred by most workers.
Hard and brittle tissues are as a rule more easily cut in celloidin.
For the staining of bacteria in sections paraffin imbedding is
necessary. Both methods should be learned and practiced with equal
facility; a working knowledge of both is essential in pathologic
investigation and diagnosis.
1. =CELLOIDIN IMBEDDING.= The granular form of Schering’s celloidin
is the best preparation to use, although good results can be obtained
by using a cheaper well-washed gun-cotton. In purchasing the latter
care should be taken to secure a sample that dissolves easily in
alcohol and ether, and does not give off yellow fumes when exposed
to the light. Schering’s granular celloidin keeps well, and forms
on solution a firm, tough, transparent imbedding mass, so that thin
sections are obtainable without difficulty. When kept long in stock
celloidin becomes hard and dissolves more slowly. For use three
solutions are made, thick (10 per cent), thin (2 per cent), and
medium (5 per cent). The celloidin granules or shavings are put into
a wide-mouthed bottle having a tight stopper, and are covered with
absolute alcohol and well shaken, and left for 24 hours. An equal
quantity of pure ether is then added, the mixture is well stirred
and allowed to stand for another 24 hours, when it is again stirred
and evenly mixed, and is then ready for use. When gun-cotton is used
it is torn into fine shreds and added to a mixture of equal parts of
absolute alcohol and pure ether and shaken until sufficient has been
added to give the solution the desired strength.
Slow Celloidin Method.
1. Absolute alcohol 24 hours.
2. Equal parts absolute alcohol and ether 24 hours.
3. Thin celloidin for 1-3 days.
4. Medium celloidin for 1-3 days.
5. Thick celloidin for 1-3 days.
6. Block.
The tissue is blocked by removing it from the thick celloidin on a
section-lifter and placing it on a block of vulcanized fiber or wood
with enough of the thick celloidin about it to form a good matrix.
The preparation is then allowed to evaporate in the air until the
surface of the celloidin becomes firm (does not stick to the finger).
The block is then placed in 80 per cent alcohol or pure chloroform
until hard enough for cutting (1-24 hours). Cork should not be used
for blocking, nor should wood unless the tannin has been removed by
long treatment with alcohol-ether. The celloidin will adhere more
firmly to the fiber block if the latter is dry, and if there is a
sufficient layer of celloidin between the tissue and the block. The
imbedded tissues may be preserved in 80 per cent alcohol for a long
time, but gradually lose their staining power. They may also be kept
dry by coating them with melted paraffin. The blocks when preserved
in alcohol may be marked with an indelible pencil.
For imbedding large pieces of tissue in celloidin a glass dish may be
filled with thick celloidin and the infiltrated tissue placed in it
with cutting surface down. The celloidin is then allowed to evaporate
slowly under a glass cover, and fresh celloidin may be added as
shrinkage occurs. When well-hardened the celloidin is cut out of the
dish and the block trimmed to the proper proportions, and attached
directly to the object-holder of the microtome or to a block of wood
by a few drops of thick celloidin, allowing it to dry for a minute or
so and then immersing in 80 per cent alcohol. The block may be cut on
the freezing microtome by soaking the hardened celloidin in water to
remove the alcohol (when block sinks), then coating it with saturated
gum arabic solution, and freezing.
Rapid Celloidin Method.
I have used the following method in my laboratory for a number of
years as a regular procedure in practical diagnostic work, and the
results have been uniformly good.
1. Fresh tissue cut thin, or uterine curettings, in absolute
alcohol 1½ hours, three changes of fresh absolute during this time.
2. Alcohol-ether 1 hour.
3. Thin celloidin at incubator temperature 12 hours (over night).
4. Medium celloidin 1 hour.
5. Block from medium celloidin, evaporating celloidin by blowing,
and building up matrix by adding successive layers of celloidin.
6. 80 per cent alcohol 1-3 hours.
7. Cut.
The quick celloidin methods recommended in the literature (Kaufmann,
Stepanow, Scholz and others) require more time, and do not give
better results. Material received from operative clinics late in the
afternoon can be sectioned and stained usually by ten o’clock the
next morning.
2. =PARAFFIN IMBEDDING.= A paraffin of a melting-point sufficiently
high enough to withstand summer heat is advisable; a 52°C. paraffin
answers for this latitude. The use of softer paraffins is not
necessary. The paraffin-oven should be regulated at a constant
temperature of 54-55°C. Over-heating of the tissue while in the
oven must be carefully safeguarded, so that in the management of a
paraffin-oven the care of the thermo-regulator is the most important
thing.
Slow Paraffin Method.
1. Thorough dehydration in absolute alcohol 12-24 hours.
2. Aniline oil to remove alcohol, until tissue becomes transparent
or sinks.
3. 1st. Xylol, ½ hour, to remove aniline oil.
4. 2nd. Xylol, 1-2 hours, until translucent.
5. 1st. Paraffin, 52°C. in oven, 1 hour, to remove xylol.
6. 2nd. Paraffin, 52°C. in oven, 1-12 hours.
7. Block.
The use of xylol-paraffin is not necessary. For blocking staining
dishes, watch-glasses, glass salt-cellars, paper-boxes, metal
frames., etc., may be used. A thin smear of tincture of green soap
or glycerin is rubbed over the inside of the imbedding box, and
it is nearly filled with fresh melted paraffin. With warm forceps
the tissue is taken out of the bottle of second paraffin in the
oven, and arranged in the melted paraffin in the imbedding dish
in the proper position for cutting. Care must be taken that the
melted paraffin is not hot enough to “burn” the tissue, else its
staining-power may be affected. The surface of the paraffin is
then cooled by blowing upon it, and as soon as a film appears upon
the surface, the dish is carefully immersed in cold water, so that
the paraffin may set quickly. When cool the paraffin-block should
slip out of the dish and float to the surface. It is then trimmed
to the desired shape, leaving a good matrix of paraffin around the
tissue. The paraffin-block is then fastened to a wooden block or
to the object-holder of the microtome by means of melted paraffin
(a hot knife is drawn along the under-surface of the block and the
latter immediately pressed upon the wooden block or object-holder).
Chloroform, cedar oil, benzene, carbon bisulphide, etc., may be used
instead of xylol, and each one possesses certain advantages for
certain purposes. Benzene is advisable for osmic-acid preparations.
Rapid Paraffin Method.
The above method can be greatly shortened for uterine curettings,
thin bits of tissue, etc., if the various steps are closely watched,
and if the entire process is carried on in the oven. The whole
process may be carried out in 1-3 hours, a very great advantage over
the quick celloidin method. A simpler and cheaper method is that
recommended by _Heller_, _Henke_ and _Brunk_, as follows:—
Acetone Method.
1. Small bits of fresh tissue, or tissue fixed in formol, in
water-free acetone over copper sulphate for ½-1½ hours.
2. Transfer tissue directly to fluid paraffin in the oven for ½-1½
hours; the acetone evaporates, and the tissue is infiltrated with
paraffin; or put into xylol 5-10 minutes, then in paraffin 15-20
minutes.
3. Block.
By this method the entire process of fixing, hardening, imbedding,
cutting and staining can be carried out in half an hour, and by it
the freezing-microtome can be dispensed with in a large part of quick
diagnostic work.
Pyridin Method.
1. Fix in formol.
2. Dehydrate and clear in pyridin.
3. Paraffin.
This method requires a longer time than the acetone method, and is
not so good.
Combinations of celloidin and paraffin may be employed by imbedding
first in celloidin, transferring block to origanum oil, then xylol
and finally paraffin. Formol-agar has been recommended by _Bolton_
and _Harris_ for simultaneous fixation and imbedding. It offers no
especial advantages. _Wright_ uses formol-gelatin for imbedding
tissues for sectioning on the freezing microtome. The bits of tissue
are placed in warm 20 per cent pure gelatin; this is allowed to set,
and the block placed in 10 per cent formol for 24 hours; it is then
frozen and cut. Soap-gelatin, glycerin-gelatin, gum-glycerin, etc.,
are now rarely used for imbedding.
NOTE:—When a number of pieces of tissue are blocked at the same
time, either in celloidin or paraffin, they may be tagged with paper
labels fastened to the tissue with a drop of gum tragacanth or gum
arabic. As the gum is not soluble in any of the infiltrating and
imbedding media the labels remain attached until the block is ready
for cutting.
CHAPTER XXIII.
SECTION-CUTTING.
Fixed and hardened tissues may be sectioned on a microtome without
freezing or imbedding, or they may be cut on a freezing-microtome;
or, having been imbedded in either celloidin or paraffin, they may be
sectioned on any form of microtome suited to the purpose desired. The
choice of the microtome depends, therefore, upon the character of the
work. For ordinary purposes the small sliding microtomes furnished
with a crank are preferable, as they are more easily kept in order
and can be used for rapid cutting. For serial paraffin sections
the Minot automatic rotary microtome is advised. Especial types
of microtomes can be obtained for the cutting of large sections,
particularly for brain-sections. The Bardeen freezing-microtome
I have already recommended as the most practical instrument for
freezing work. In all cases it is absolutely necessary that the
instruments be in good order and that they work with precision. The
microtome knife must be carefully honed and stropped. A heavy, rigid,
biconcave knife-blade should be used, and a good hone and strop
are absolute necessities. In honing, the knife is drawn from heel
to toe with cutting edge forward; while, in stropping, the motion
is reversed, the back of the knife being forward, and the motion
from toe to heel. The knife during the honing should be fitted in a
knife-holder, and the back of the blade protected from the hone by a
knife-back. The knife-blade must be kept free from grease and dirt,
and nothing must be permitted to touch its edge. When many sections
are cut at one sitting frequent stropping is necessary. When the
cutting is finished the knife should be removed from the microtome,
carefully cleaned and dried, and placed in its box. The slide and
other bearings of the microtome should be well-oiled with the best
microtome oil, and kept free from dust.
1. =Cutting of Fixed Tissues Without Imbedding or Freezing.= It
occasionally becomes necessary in pathologic work to cut tissues
directly upon the microtome without freezing or imbedding. This can
be done satisfactorily in the case of very firm substances, such
as amyloid liver and spleen, etc. The sectioning is done in the
wet, using 80 per cent alcohol, as in celloidin cutting. A large
celloidin knife should be used, and the blade should be flooded in
alcohol. A large piece of the tissue, or, better, several pieces,
are clamped in the object holder with only a small layer of tissue
above the holder. The knife-blade is placed at the least possible
angle to the pieces of tissue. To obtain sections of 15-20 microns in
thickness the object-holder is raised each time about 25-30 microns.
The sections will vary in thickness, some thin, others very thick.
2. =Sectioning of Frozen Fixed Tissues.= Tissues fixed in formol,
Müller’s fluid or Orth’s fluid may be frozen directly on the
freezing-microtome without previous washing. For other fixations
previous washing is necessary. Tissues in alcohol must have the
alcohol thoroughly washed out before freezing. Celloidin blocks may
also be cut by freezing after the removal of alcohol. The fixed
tissues cut to the proper size are placed on the object-holder of the
freezing-microtome in a drop of saturated gum-arabic and the freezing
carried out as directed above. (See Page 214.) As the freezing
causes little or no damage to fixed tissue, the frozen sections may
be transferred directly to alcohol, or floated out on the dilute
molasses or sugar-dextrin solution, and stained separately or in
sheets, according to the directions given in the next chapter.
3. =Sectioning of Celloidin Blocks.= The celloidin blocks are
fastened in the object-holder of the microtome, and the knife-blade
(a longer and broader one than for paraffin-sectioning is advisable)
set nearly parallel with the longitudinal axis of the microtome,
so that the cutting-edge, striking the block at a very slight
angle, will be utilized for the greater part of its length in
cutting entirely across the surface of the block. Both object and
knife-blade must be kept constantly wet with 80 per cent alcohol;
a broad camel’s-hair brush or a drip-bottle can be used for this
purpose. The sections, as they are cut, are transferred from the
knife to 80 per cent alcohol by sweeping the finger or brush from
above toward the edge of the blade. Care must be taken not to strike
the brush against the edge of the blade, and this can be avoided by
using the brush always with a downward stroke. As the sections are
cut they are guided onto the knife-blade by means of a fine-pointed
camel’s-hair brush, if the sections show any tendency to curl. They
should be smoothed out at once on the knife, and then transferred to
the dish of 80 per cent alcohol. Sections 7-10µ thick are easily cut
in celloidin, if the process of imbedding has been carefully carried
out.
To prepare _serial-sections_ from celloidin blocks the sections as
they are cut must be arranged in their order on the knife-blade and
thence transferred in this order to a slide or glass-plate, to which
they are either fastened so that they cannot become loose during
the staining process, or they are fastened together by a film of
celloidin and stained in one piece. To fasten the celloidin sections
to the slide _Mallory_ and _Wright_ suggest the cutting of the block
in 95 per cent alcohol, and the arrangement of the sections in their
order on the knife, whence they are drawn on to a dry, clean and
numbered slide. Ether vapor from a half-full bottle of ether is
then poured over the slide, slowly, to flatten out the sections and
fasten down the frilled edges. The slides are then transferred to
80 per cent alcohol to harden the celloidin, and they are kept in
this solution until ready for staining. Celloidin sections may also
be fastened to the slide by albumin glycerin (equal parts white of
egg and glycerin with a crystal of phenol or thymol). The celloidin
sections, as they are cut, may be transferred to a dry gelatinized
slide or plate (16 grms. gelatin in 300 cc. warm water), and are
then covered with a thin celloidin; the slide is then placed in
water at 50°C., the gelatin dissolves and the celloidin film floats
off; the latter is then stained as one section. Serial sections of
celloidin-blocks may also be arranged upon slides or plates covered
with a coating of the _Schmorl-Obregia_ sugar-dextrin solution or
diluted New Orleans black molasses; the slide or plate is flooded
with absolute alcohol; drained; a thin celloidin is then poured over
it; it is immersed in warm water and the celloidin-sheet containing
the sections is liberated, and is then stained as a whole or cut
into strips as desired. The molasses method is the cheapest and
simplest method, and much to be preferred to the Weigert closet-paper
method, by which the sections are arranged upon a strip of moist
closet-paper which is either held upon the slide or kept upon a
piece of blotting-paper wet with 80 per cent alcohol. A slide or
glass-plate is covered with a thin layer of celloidin and the strip
of paper containing the sections is laid upon the celloidin surface,
the sections down, so that these stick to the slide as the paper is
carefully peeled off. The preparation is dried with filter-paper and
a thin layer of celloidin poured over the slide and sections. The
celloidin is then hardened in 80 per cent alcohol and the sections
stained on the slide; or the celloidin film is removed by immersion
in warm water, and then treated as one section. Celloidin films may
be marked with a brush dipped in a water solution of methylene-blue.
This should be done as soon as they are made. _Langhans’ method_
is advised for the remounting of serial sections from tissues
stained in bulk and imbedded in celloidin. The sections are cut in
origanum-alcohol (1 part absolute alcohol to 3 parts origanum oil)
and placed on a slide in a layer of origanum oil, blotted and mounted
in balsam. Should sections become milky in the oil, renew the latter
until they are cleared.
4. =Sectioning of Paraffin Blocks.= The paraffin block trimmed to
the proper proportions, leaving a rim of paraffin about 2-3 mm. wide
all about the tissue, is fastened to the wooden block by melting the
under side of the paraffin by drawing a hot knife across it, and then
immediately pressing the block upon the wood. The wooden block is
then clamped into the object-holder. If desired the paraffin block
can be fixed directly to the metal plate of the object-holder in the
same manner. The block is raised until the level of the edge of the
knife is reached. The knife is placed transversely or at a slight
angle, and the cutting done with a relatively small portion of the
edge. The paraffin block may be breathed upon to warm slightly the
upper layer when a hard paraffin is used, and the knife is then drawn
carefully through the block, guiding the section onto the knife-blade
by means of a fine-pointed camel’s-hair brush held in the left hand.
It is preferable, I think, to have a wider rim of paraffin at one
corner of the block and to place the block with that corner at a
slight angle to the knife, so that the edge of the blade will first
engage the block at that point. The tip of the camel’s-hair brush
moistened in water catches the corner of the section as the knife
begins to cut, and pushes it over onto the blade, thus holding the
section flat and preventing curling. When entirely cut through, the
section is removed from the knife by the brush, still holding it at
the corner first touched, or if necessary it is again moistened and
applied to the upper side of the section, lifting the latter off
the knife. The block is trimmed down to the proper level by cutting
thick sections first, then sections of the desired thickness when
the level of the entire block is reached. The sections are then
transferred, with their shiny side down, just as they come off the
knife, to a slide, sheet of paper, warm water, warm molasses or
sugar-dextrin solution, or 70 per cent alcohol, and treated further
according to the directions given in the next chapter. The presence
of ridges on the cut surface of the block is an indication that the
knife needs stropping. The knife must cut the paraffin, not scrape
it. Crumbling of the paraffin is the result of imperfect infiltration
during the imbedding process; water, alcohol, aniline oil or xylol
may be left in the tissue, the paraffin may contain oil, or the
cooling process may have been delayed. Curling of the sections can
be prevented by using a sharp knife and slightly warming the surface
of the block by breathing upon the block, use of a warm knife or
spatula, heat-focus, etc. For ordinary work paraffin sections 5-7µ
thick are easily obtained; for especial work 1-2µ sections may be
obtained by especial care in imbedding, using graded paraffins and
finally imbedding in a 56°C. paraffin. The pieces of tissue should
be small, and the sections may be cut with a knife wet with water or
alcohol. _Serial paraffin sections_ are easily obtained; in fact,
paraffin imbedding is by far the best method for serial cutting. To
cut ribbons of sections upon the ordinary slide microtome the block
must be clamped into the object-holder so that the edge of the block
facing the knife, as well as the opposite side of the block, is
parallel with the knife-edge. The knife should be placed at right
angles to the microtome. If the paraffin has the right consistency
the edges of the sections as they are cut will adhere, and a ribbon
of serial sections will be pushed up over the knife. This ribbon
can be cut into pieces of the desired length and mounted according
to the directions given in the next chapter. The Minot automatic
rotary microtome is especially well adapted for the cutting of ribbon
sections. Failure of sections to adhere to each other is usually
the result of too hard consistence of the paraffin, and this can be
remedied by warming slightly, according to the method given above. If
the paraffin is too soft the sections fold together. The block may be
cooled in ice-water, or the sections may be cut with a cooled knife.
Paraffin knives so constructed that they may be cooled by a stream
of ice-water are supplied by dealers in microtomes. The conditions
essential for successful paraffin cutting are perfect infiltration
and imbedding, sharp clean knives and a certain degree of skill in
manipulation that can only be secured by practice.
CHAPTER XXIV.
THE PREPARATION OF MOUNTED SECTIONS.
Sections of fresh material, unimbedded or imbedded tissues must be
treated by a series of processes before they are finally permanently
mounted and ready for use. These processes in general are:
=preparation for staining=, =staining=, =differentiation=, =washing=,
=dehydration=, =clearing= and =mounting=. The general procedure will
be modified to some extent by the character of the tissue, manner
of preparation (fixed or unfixed, imbedded or unimbedded, unstained
or stained), character of stain (affected by alcohol, xylol, etc.),
and the mounting agent (glycerin, balsam, damar, colophonium). Two
or more of these steps may be combined in one; the same agent may
differentiate, dehydrate and clear. Several stains may be combined in
one solution, or it may be necessary to use them in succession. The
very greatest variation is possible in pathologic technique; in fact,
practically every laboratory worker modifies methods according to the
light of his individual experience. The really important thing is
to be master of the method, and not allow the method to control the
situation. One of the greatest attractions about laboratory work is
the infinite possibility of variation and improvement of methods and
the invention of new ones.
I. PREPARATION FOR STAINING.
a. =Frozen Sections of Fresh Tissue.= Frozen sections of fresh
tissues, as well as those obtained by the single or double razor,
may be stained by floating the section on a slide, staining it
directly (carbol-kresyl-echt-violett or carbol-thionin), examining
in the stain or washing, dehydrating, clearing and mounting; or the
section may be fixed to the slide with molasses or sugar-dextrin
solution, covered with a celloidin-film, and treated according to
the methods followed for paraffin or celloidin sections. Sections
of fresh tissue may be fixed in formol or alcohol, and then treated
by the same methods as celloidin sections. (See also Page 220.)
b. =Sections of Unimbedded Tissues.= These may be handled for
staining in the same way as paraffin, celloidin or fresh-tissue
sections, either when sectioned directly or after freezing. The
sections may be stained directly, on the slide, cover-slip, or in
the staining solution, or they may be transferred into celloidin
sheets by the same methods employed in the preparation of paraffin
sections.
c. =Celloidin Sections.= These may be transferred from water or
alcohol directly to the stain. It is not necessary to remove the
celloidin. If not stained soon after cutting they should be
preserved in 95 per cent alcohol. Celloidin sections may be stained
on the slide by simply blotting the section firmly on the slide,
without permitting it to become dry, and manipulating it carefully
through the various solutions; or the section may be fixed to the
slide by the use of 95 per cent alcohol, ether-vapor and fixation
in 80 per cent alcohol; or the section may be fixed to the slide
by the methods given above under the cutting of serial sections
of celloidin blocks. The most common method of preparation of
celloidin sections for staining is to transfer the sections from
alcohol into water to straighten them out, and then to transfer on
the spatula into the stain. For the treatment of serial celloidin
sections see above.
d. =Paraffin Sections.= Paraffin sections may be stained directly
_without removing the paraffin_. This is especially advisable in
the staining of tubercle-bacilli and in other cases where the use
of alcohol is to be avoided. For many stains this method cannot be
used. The sections as they are cut are floated directly into the
warm stain, on which they flatten out, and are then transferred to
the other solutions on the section-lifter, finally dried on the
slide, in the incubator or over the flame, cleared in xylol and
mounted in balsam. Paraffin sections may also be stained without
removing the paraffin by being transferred directly from the
knife on to 80 per cent alcohol, stained, washed, dehydrated in
absolute alcohol or by drying, cleared and mounted. The section is
transferred from one solution to another on the slide or spatula.
The paraffin is removed during the clearing in xylol in both of
these methods. The treatment with xylol must be on the slide, else
the section may go to pieces. The staining of the section in the
paraffin usually takes more time than staining after the paraffin
has been removed, but the process can be hastened by heating the
stain.
_Slide and Cover-slip Preparations._ Paraffin sections may be
affixed to a slide smeared with a thin film of albumin-glycerin
(equal parts of filtered beaten white of egg and glycerin, with
crystal of phenol or thymol, or 1 grm. of sodium salicylate to
100 grms. of the mixture as a preservative). A drop of fixative
is placed upon a clean slide, and is rubbed over the slide in a
fine film with the back of the finger. The dry paraffin section
with glossy side down is placed upon the smeared slide, flattened
with a brush and then pressed firmly against the slide with the
ball of the thumb. The albumin-fixative is then coagulated in
the incubator or over the flame; the paraffin is melted over the
flame without over-heating the section and the slide covered at
once with xylol to remove the paraffin. It is then put into 95
per cent alcohol, thence into the stain; and after staining, the
section is washed, dehydrated, cleared and mounted. _Cover-glass_
preparations of paraffin sections are made by floating the
sections with glossy side downward on warm water (just below the
melting-point of the paraffin) until they straighten out and are
perfectly flat. They are then floated on to cover-slips covered
with a thin film of albumin-glycerin, the albumin having previously
been coagulated by passing the smeared cover-slips through the
flame quickly so that they do not scorch or burn. The cover-slips
with the adherent sections are then placed in the incubator for 12
hours. The paraffin is then removed by xylol, the xylol is washed
out in 95 per cent alcohol, and the cover-slips are then carried
through the processes of staining, washing, dehydration, clearing
and mounting. The cover-slips must be handled with forceps and
the section side should always be uppermost. Slides covered with
a film of albumin-glycerin may be used instead of cover-slips.
The albumin-glycerin film may be omitted, and the sections with
glossy side down floated in warm water on to clean covers or
slides; the water is drained off and the slides or covers are put
in the incubator for 12 hours. Sections adhere fairly well by this
method (_capillary attraction method_). Bubbles are removed by
careful heating. Serial ribbons of the size desired can be floated
and mounted on slides by the albumin-glycerin or the capillary
attraction method.
By far the best method of preparing paraffin sections for staining
is the _molasses plate method_, a modification, originating in
my laboratory, of the _Schmorl-Obregia_ sugar-dextrin method.
When many sections are to be stained at once it is the most
convenient method and gives uniform results. In the preparation
of sections for class-work it has no equal. It can be used also
for giving out unstained sections. When many sections must be
stained in diagnostic work the method saves much time and labor.
Fifty sections can be stained as easily as one. It combines all
the advantages of the celloidin and paraffin methods, as does the
_Schmorl-Obregia_ sugar-dextrin method, but is much cheaper than
the latter.
_Schmorl_ advised the use of a sugar-dextrin solution (cane sugar
solution [1:1] 300 cc., 80 per cent alcohol 200 cc., yellow
dextrin solution [1:1] 100 cc.) to be run over a perfectly clean
glass plate or slide until the entire surface is covered with an
even layer. The paraffin sections as they are cut are arranged in
order on the wet plate, and when the plate is full, it is heated
sufficiently to flatten and smooth the sections. The plate is then
placed in an incubator for 3-12 hours to harden and dry. When dry
it is immersed in xylol to take out the paraffin, then treated
with absolute alcohol for 10-15 minutes, the alcohol drained off,
and the plate covered with a thin layer of celloidin (celloidin or
photoxylin 10, absolute alcohol 100, ether 100). As soon as the
celloidin sets (1-2 minutes) the plate is immersed in warm water
and the celloidin film containing the sections is detached. It
can now be carried through the staining, washing, dehydrating and
clearing solutions as one section, and in the clearing solution cut
into strips or single sections, as desired, for mounting. _Huber
and Snow_ improved the method greatly by floating the paraffin
sections directly on to warm dilute sugar-dextrin (a 10 per cent
solution of Schmorl’s stock-solution will suffice), and plating
the sections directly from the latter. This method of using the
dilute solution is less expensive, much cleaner, and saves time
in drying in the incubator. The results are in every way better
than with the Schmorl solution in full strength. The formation of
bubbles and crystals is almost wholly prevented, and less dust is
caught on the plate. In my laboratory we have modified the method
still further by using a 10 per cent solution of New Orleans black
(or baking) molasses instead of the more expensive sugar-dextrin
solution. As the molasses costs but 20 cents a gallon, a gallon of
the dilute solution costing 2 cents can be used indefinitely if
fermentation be prevented by a crystal of phenol or thymol. The
paraffin sections are floated on to this dilute molasses solution
warmed sufficiently to smooth out the sections; 4 × 5 glass plates
(old negatives) thoroughly cleaned and kept in alcohol are immersed
in the warm molasses solution and the sections arranged on them as
desired, lifting out of the solution that part of the plate covered
with sections as they are drawn upon it. As soon as the plate is
covered it is drained, and is then flooded with absolute alcohol.
After 1-2 minutes the alcohol is drained off and the plate flooded
with thin celloidin, which is allowed to set for a minute or so,
and the plate then immersed in warm water in which the celloidin
film containing the paraffin sections is detached. This film is
then handled by catching it at the two corners of one end with
the fingers, or better still by a pair of forceps held in each
hand. The film is put first into xylol to remove the paraffin,
then into 95 per cent alcohol, then into water and thence into the
staining solution. After staining the film is washed, dehydrated
and cleared, and in the clearing solution is cut into strips or
single sections by means of the wheel-shaped paper-cutter used by
paper-hangers. The pieces are then mounted. A dilute sugar-dextrin
solution can be used instead of the molasses-solution, but the
latter is much cheaper and does just as well. Aside from this
advantage our method of transferring the paraffin-sections into the
celloidin film without first removing the paraffin saves a great
deal of time, as it is not necessary to wait for the plates to dry
in the incubator. The same method can be applied to the staining
of single paraffin sections on the slide. The conversion of the
paraffin section into a celloidin preparation without any loss of
time for drying is so quickly and easily carried out that I advise
it above all others. The same method may also be applied to the
staining of fresh and fixed tissues cut on the freezing microtome
or sectioned without imbedding. The success of the plate-method
will depend largely upon the state of the glass-plates when put
into the molasses solution. They must be perfectly clean or the
celloidin sheet will not separate well. It is best to keep them
in alcohol until they are needed. The celloidin must be of the
right consistency, the layer must be thin, and cover the entire
plate uniformly. It must not be allowed to harden too much before
immersion in water or it will be tough and will shrink. Handling
of the celloidin-sheets with the bare hands is not advisable
because of the large number of epithelial cells adhering to the
celloidin. The sheets are easily changed from one solution to
another by catching them with forceps; the use of a glass-plate
to transfer them is not necessary. When it is desired to preserve
sections for future staining the celloidin sheet containing the
paraffin-sections can be kept in 80 per cent alcohol indefinitely.
NOTE:—If the glass-plate is numbered with a blue wax-pencil
after the paraffin sections are floated on, the marking will be
transferred to the celloidin sheet, and the latter will retain the
marking through all solutions.
II. STAINING AND DIFFERENTIATION.
Staining is necessary to bring out clearly the constituent elements
of the tissues and their relations with each other, and for the
demonstration of histologic structures or chemical substances that
would otherwise be nearly or wholly invisible. The technique of
staining depends upon the fact that stains or dyes possess certain
affinities for the tissue-elements or for certain simple or complex
substances present in the tissues (_microchemic reactions_). These
affinities vary greatly with the dye. Some dyes have an affinity
only for single constituents of the tissue (_elective_ or _specific
stains_); others have an especial affinity for the nucleus (_nuclear
stains_), others stain all the tissue-constituents diffusely
(_diffuse_ or _protoplasmic stains_). There are but few pure elective
stains for single tissue-elements; the majority of stains will stain
more than one of the tissue elements, but may show an especial
affinity for certain ones. As a result of these variations in the
affinities of dyes for the constituents of the tissues it becomes
possible to manipulate the dyes or to combine them in such a way
that a specific differentiation of many tissue-elements is possible
through the use of _different methods of staining_. These methods are
based in part upon the use of different mordants, the employment of
several stains in combination or in succession, the mixture of stains
to form a new staining compound, the phenomenon of _metachromasia_,
the differentiation of certain tissue-elements by the removal of
the stain from the structures for which it possesses a weaker
affinity, and by the employment of different microchemic reactions.
The two most commonly employed methods are the _progressive_, in
which the stain is allowed to act until the affinities of certain
tissue-elements have been satisfied when the staining process
is interrupted; and the _regressive_, in which the tissue is
over-stained, and the dye withdrawn from the tissue-elements for
which it possesses the weakest affinities leaving the other elements
stained. This latter process is usually called “_differentiation_,”
and the chief substances used for such differentiating are dilute
acid, acid alcohol, acid stains, aniline oil, aniline-xylol and
alcohol. Some workers use the regressive method for such simple
stains as hæmatoxylin, overstaining, and then differentiating with
acid alcohol before counterstaining with eosin. The results obtained
in this way are much less satisfactory than is possible with the
progressive method.
Tissues may be stained in the body during life (_intravital
staining_), or immediately after removal from the body (_supravital
or survival staining_), either before or after sectioning. (See Page
217.) Fixed tissues may be stained in bulk or in sections.
Staining Tissues in Bulk.
This method is not often used in pathologic work. The fixed and
hardened tissue is cut into small pieces, placed in the staining
solution for several days, washed thoroughly, dehydrated in
alcohol, imbedded, cut, and mounted without further staining.
Alcoholic solutions penetrate best; hæmatoxylin, hæmalum, carmine
and alcoholic solutions of the aniline stains may be used.
_Metallic impregnation_ (gold or silver salts) of fresh or fixed
tissues is but little used in pathology. (See Staining of Nervous
System, and Spirochætes.)
Staining of Sections.
=Celloidin sections= are lifted from water or alcohol into the
stain by the needle or section-lifter. The use of the latter
is advised, as by it the section can be floated flat on to the
staining solution. When many celloidin sections are to be stained
at once they can be stained in small tea-strainers and transferred
in these from one solution to another. =Paraffin sections= may be
floated directly on to the stain without removing the paraffin; or
they may be stained on the slide or cover-glass after removing the
paraffin, the stain being dropped on to the section, or the slide
or cover-slip is immersed in the stain. Special staining-dishes
for the staining of paraffin sections on slides and covers can be
obtained. Paraffin sections transferred to celloidin sheets by the
plate method can be put into the staining-solution while on the
glass-plate, or the films can be detached and transferred from one
solution to another by means of forceps. This is the easier way,
and it is not necessary to touch the films with the fingers.
General Rules for Staining.
1. The stain should be filtered just before being used, in order
to remove precipitates, moulds, etc. Unless they have been diluted
most of them can be used over and over again, hence after using
they should be filtered back into the stock bottle.
2. A liberal amount of stain should be used. Slides and cover-slips
are given enough stain to cover completely the section, when
the staining is done on the slide, or they may be immersed in
staining-dishes. Plates and celloidin sheets should be stained in
large trays. Sections and celloidin films should be flat without
folds or wrinkles, and they should not touch one another when
several are stained at the same time. Transference of the section
from water or dilute alcoholic solutions to dilute or stronger
alcohol respectively for a moment and then back again will usually
straighten out curled or wrinkled celloidin sections.
3. Stain until the section is properly stained. Control this by
removing it from the staining-solution and examining it in water on
a glass-slide without a cover-slip, using the low-power. Sections
will always appear more deeply-stained when cleared than when
examined in water, hence due allowance should be made. Celloidin
sheets can be examined on glass-plates. The time-limits given in
staining methods are only approximate; no absolute rules can be
laid down as to the length of time necessary to obtain a good
stain. The methods of fixation and hardening, age of the tissue,
age of the stain, etc., affect the staining power. Some stains
lose their staining-power after a time; others require a period of
ripening before they yield the best results. As a rule staining
may be intensified or hastened by staining in the incubator or at
a higher temperature, by concentrating the stain, or by the use of
such substances as aniline oil. When differentiation is necessary
the process should also be controlled by frequent examination of
the section, as above for staining. Usually the section can be
examined in the differentiating fluid.
III. WASHING.
Thorough washing after staining is necessary after nearly all
stains. The washing should usually be done by soaking the sections
in several changes of distilled water, although tap-water, alcohol,
alum-water, and other solutions may be used to intensify the
staining effects. When this is done a final washing in distilled
water or alcohol is usually necessary. Differentiating fluids
should always be thoroughly removed from the section before
mounting. Sections should not be allowed to lie in wash-water that
is colored by the stain; as soon as the wash-water becomes colored
it should be replaced by fresh. When sections are left lying in
the wash-water for some time the vessel containing them should be
covered to prevent the settling of dust on the sections, as it is
practically impossible to remove from the latter dust or other
precipitates that may become attached to them. Some stains give
better results after long washing; others are easily washed out if
the sections are left standing in the wash-water. The time-limits
of washing will depend upon the character of the stain employed.
IV. DEHYDRATION.
Dehydration of the sections is usually produced by passing them
through two alcohols, 80 per cent and absolute, or 80 per cent and
95 per cent. For certain clearing reagents (xylol) it is necessary
to use absolute alcohol. When carbol-xylol is used as a clearing
reagent absolute alcohol is not necessary, and 95 per cent can
be used instead for the second dehydrating solution. Usually a
minute in each alcohol is sufficient for the dehydration of single
sections. If dehydration with alcohol is not desirable because of
its action on the stain it is possible to dehydrate and clear in
xylol by repeatedly blotting the section with absorbent paper,
covering the section several times with xylol and then blotting.
The section should never be allowed to become perfectly dry.
Dehydration with alcohol may also be avoided by staining paraffin
sections without removing the paraffin, drying in the incubator
or over the flame, removing the paraffin in xylol, and mounting.
Imperfect dehydration is shown by the presence of white spots or a
milky cloud in the section when it is put into the clearing fluid.
V. CLEARING.
After dehydration, sections must be cleared in some solvent of
balsam before they can be mounted in the latter medium. When 95
per cent alcohol has been used for the final dehydration the
sections may be completely dehydrated and cleared at the same
time by the use of carbol-xylol (xylol, 3 parts; melted crystals
of carbolic acid, 1 part; add melted carbolic acid to the xylol
to prevent formation of crystals). The sections (on the slide
or cover-slips, in celloidin sections or films) are transferred
from the alcohol, draining or blotting off excess of the latter,
into the carbol-xylol, and left until perfectly clear. This can
be most easily determined by viewing the sections against a dark
background. Carbol-xylol cannot be used for sections treated with
aniline stains. These are dehydrated in 95 per cent alcohol, and
the final dehydration and clearing accomplished by repeatedly
placing xylol upon the slide and blotting it out until the sections
are transparent. Turpentine, chloroform, benzine, toluol, the
oils of bergamot, cloves, thyme, lavender, origanum cretici, and
cedarwood, aniline oil, and various mixtures of these oils are
also used as clearing agents. The majority of these will clear
from 95 per cent alcohol, but not so readily as carbol-xylol; they
have individual disadvantages of taking out the eosin, affecting
aniline colors, dissolving celloidin, making sections brittle,
slow action, clinging odor, etc. Chloroform and benzine may be
used for clearing osmic-acid preparations; oil of turpentine is
also good for clearing sections stained with kresyl-echt-violett,
and Wright’s blood-slain. With but few exceptions carbol-xylol
and xylol meet all requirements better than any other clearing
reagents. There is but one disadvantage in the case of
carbol-xylol; some of the phenols in the market cause a fading of
eosin and hæmatoxylin stains. _DeWitt_ has shown that this fault
can be corrected by redistillation, stopping the distillation as
soon as the temperature begins to rise above the constant boiling
point of the phenol; or the carbol-xylol that fades the stains can
also be corrected by supersaturating it with a mixture of sodium
bicarbonate one part, and sodium-potassium tartrate two parts.
Sections kept in xylol or carbol-xylol should be protected from
dust and evaporation; it is not a good plan to keep them in these
solutions for more than 24 hours.
VI. MOUNTING.
Permanent mounts are made in glycerin, potassium acetate, lævulose,
glycerin gelatin, balsam, damar or colophonium. For celloidin and
paraffin sections a solution of =Canada balsam= in xylol is most
commonly used for mounting. Celloidin sections (celloidin films are
best cut into strips and single sections when in the carbol-xylol;
the wheel-shaped paper-cutter used by paper-hangers is the best
instrument for this purpose) are lifted onto the slide from the
clearing-fluid; folds or wrinkles in the celloidin are straightened
or removed by cutting the celloidin at right angles to the section
in order to relieve the tension, and the section is then blotted
firmly against the slide by means of a pad of absorbent paper.
The greatest care should be taken to prevent wrinkling, folding
or turning over of the edge of the section. As soon as the pad
is removed a drop of balsam is placed upon the section and the
cover-glass put over it. There should be just enough balsam used to
fill the space between cover-slip and slide, so that air-bubbles
are not formed. The balsam must not be so thin that the cover-glass
will float about on the liquid, or so thick that it does not spread
well. In the latter case warming the slide may cause it to spread
more readily, but care must be taken not to injure the stain by
over-heating. Paraffin sections on the slide are similarly blotted
and covered with balsam and cover-glass; those on cover-slips are
blotted between folds of absorbent paper and immediately placed
with section-side downward upon a drop of balsam that has been put
upon the slide.
=Xylol-damar= may be used in place of Canada balsam; it is cheaper
and colorless, but it tends to become cloudy. =Colophonium= is
the cheapest of the three and has but little color; it is highly
recommended by many workers. In a xylol-solution it may be used for
aniline stains; a chloroform solution is advisable for the mounting
of osmic-acid preparations; while a solution in turpentine and
shellac is recommended for Weigert’s neuroglia method, Wright’s
blood-stain, and other special staining methods.
=Glycerin=, =potassium acetate=, =laevulose= and =glycerin-gelatin=
are used for the preservation of amyloid-, mucin- and fat-stains,
as well as for other preparations that do not permit the use of
alcohol and xylol. =Glycerin-gelatin= is probably the best medium
for this purpose. It is made according to _Kaiser_ by soaking 7
grms. of gelatin for 2 hours in 42 cc. distilled water, then adding
50 grms. glycerin and 1 grm. carbolic acid; the mixture is warmed
10-15 minutes, stirring constantly, and filtered while hot. It is
also made by taking water, 200 cc.; gelatin, 20 grms.; powdered
white shellac, 2 grms.; Farrant’s solution (gum arabic, glycerin,
solutio acidi arsenicosi conc., _aa._ 30.0 grms.), dissolve by
warming, and filter while warm. To mount in this medium the section
is placed on the slide, and blotted with absorbent paper. A drop
of warm glycerin-gelatin is then placed on it and the cover-slip
affixed. The drop spreads evenly beneath the cover-glass and
becomes solid as it cools. Mounts in glycerin, glycerin-gelatin,
potassium acetate and laevulose must be cemented around the borders
of the cover-slip with asphalt, wax, paraffin, gold size, etc.,
using a brush or glass rod for this purpose.
VII. SUMMARY OF METHODS OF PREPARATION OF CELLOIDIN SECTIONS.
1. Fixation of the tissues in alcohol, formol, etc.
2. Wash 24 hours, when necessary.
3. After-harden in 80 and 95 per cent alcohols for 1 to several days.
4. Complete dehydration in absolute alcohol for 24 hours.
5. Equal parts of pure ether and absolute alcohol, 24 hours.
6. Thin celloidin, 1-3 days.
7. Thick celloidin, 1-3 days.
8. Block. Harden block in 80 per cent alcohol.
9. Cut; keep sections in 80 per cent alcohol.
10. Stain; differentiate.
11. Wash thoroughly.
12. Dehydrate in 80 and 95 per cent alcohols.
13. Final dehydration and clearing in carbol-xylol.
14. Place on slide and blot with absorbent paper.
15. Mount in xylol-balsam or xylol-colophonium.
VIII. SUMMARY OF METHODS OF PREPARATION OF PARAFFIN SECTIONS.
1. Fixation of the tissue in alcohol, formol, etc.
2. Wash 24 hours, when necessary.
3. After-harden in 80 and 95 per cent alcohol for 1 to several days.
4. Complete the dehydration in absolute alcohol for 24 hours.
5. Aniline-oil until tissue becomes transparent.
6. 1st. Xylol, ½ hour, to remove aniline oil.
7. 2nd. Xylol, 1-2 hours, until translucent.
8. 1st. Paraffin (52°C.), ½ hour in oven, to remove xylol.
9. 2nd. Paraffin (52°C.), 1-12 hours in oven, until infiltrated.
10. Imbed and block; cool quickly.
11. Cut sections; mount on slides or covers.
12. Remove paraffin in xylol.
13. Remove xylol in absolute alcohol.
14. 80 per cent alcohol, for a few minutes.
15. Stain; differentiate; wash.
16. Dehydrate in 80 and 95 per cent alcohols.
17. Clear in carbol-xylol.
18. Mount in Canada-balsam or colophonium.
IX. ARTEFACTS IN MOUNTED SECTIONS.
A mounted section, after passing through the various stages indicated
above, must of necessity present some appearances that are the result
of the technical methods employed. The number and degree of such
artefacts will depend upon the character of the methods employed
and the care exercised in their performance. The trained observer
ignores the presence of artefacts as having nothing at all to do
with the significance of the section itself; but to the beginner in
microscopic work they often appear to be the most important thing
in the preparation, and are given a pathologic interpretation. How
frequently do we see students, undergraduates and postgraduates, take
up a section and pick out a fold, wrinkle, tear, staining-defect,
precipitate, dirt, etc., as pathologic features! It is necessary,
therefore, for the student to acquaint himself with the nature of
artefacts so that he may ignore them and not give them an incorrect
interpretation. The most important artefacts are as follows:—
1. =Artefacts due to fixation= (mercuric, chromic, osmic, etc.,
precipitates; alterations in blood-pigment due to formol; loosening
of cells from basement membrane due to contraction [kidney tubules,
etc.: loosened endothelium in blood-vessels, particularly confusing
to students]; destruction of red blood cells, as in alcohol
fixation; poor staining due to over-fixation).
2. =Artefacts due to hardening= (contraction, desquamation of
cells, etc.).
3. =Artefacts due to imbedding and cutting= (tears, holes,
ragged edges, irregular thickness, knife-streaks, compression of
soft structures, dislocation and tearing-out of firm tissues or
material, wrinkles, folds, etc.).
4. =Artefacts due to poor staining= (uneven, spotted or streaked
staining, overstaining, understaining, poor differentiation,
precipitates, insufficient washing, fading, poor contrasts,
defective staining due to presence of paraffin, dextrin, etc., in
section).
5. =Artefacts due to poor mounting= (imperfect dehydration and
clearing, cloudiness, milkiness or opacity of section; folds;
wrinkles; turned-over edges; tears in section caused by striking it
with balsam-dropper or needle; air-bubbles; lack of balsam).
6. =Dirt and foreign-material= (opaque and translucent dirt, above
or below section; coloring-matter in balsam; ink; pigment from
pencil; cotton-, silk, wool-, linen-, vegetable and paper-fibres,
hairs, desquamated squamous epithelium, portions of insects, etc.)
CHAPTER XXV.
STAINS AND STAINING METHODS.—NUCLEAR AND PROTOPLASMIC STAINS.
=THEORIES OF STAINING.= The exact nature of the process of staining
has not yet been determined. Various theories have been advanced,
explaining the affinity of the tissues for certain dyes, on the
ground of a chemical, mechanical or chemicophysical action. The
chemical theory assumes the formation of an insoluble compound
through the chemical combination of tissue and stain; the physical
theory is based upon the assumption that the process is purely
physical or mechanical, while the chemicophysical theory holds that
it is neither purely physical nor purely chemical. The process is
not controlled by the molecular weights alone of the substances
concerned, but does depend upon the conditions controlling the
formation of solutions in general. Therefore, the theory most widely
accepted at the present time is the solution-theory, which assumes
that the staining-process is a solution of the dye in the tissue,
and that the stained tissue-element is a fixed solution of the stain
in its substance. This solution of the dye in the tissues may be a
direct action between the two (direct or substantive stains); or it
may be brought about only by the interaction of a third substance
(indirect or adjective stains). The third substance is called a
_mordant_, and the combination of the dye with the mordant is known
as a _lake_. The mordant may be added to the stain or to the tissue,
either before or at the time of staining. Many of the fixing-fluids
are mordants, particularly those containing chromic acid or its
salts. Alum, iron, and many of the metals are the most commonly used
mordants. In a general way acid mordants are used for basic colors,
and basic mordants with acid colors.
The =stains= most commonly used in pathologic work are:—
1. =Natural Dyes=:—Hæmatoxylin and Carmine.
2. =Aniline Dyes=:—_a, Acid._—Eosin, erythrosin, acid fuchsin,
orange G, picric acid, sudan III, and scarlet R (Fett-ponceau).
_b, Basic._—Methylene blue, methylene violet, thionin, toluidin
blue, kresyl-echt-violett, methyl violet, gentian violet, crystal
violet, basic fuchsin, dahlia, aniline blue, methyl green, iodine
green, safranin, Bismarck brown, and vesuvin.