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HISTOLOGICAL TECHNIQUE

BY
B. F. KINGSBURY, Ph.D., M.D.

Professor of Histology and Embryology

1915

CARPENTER & COMPANY
ITHACA, N.Y.

NOTE.

The methods in use in the Department of Histology and Embryol-
ogy and applied in the instruction, have been in the past twice
gathered together and put in printed form for the convenience of
students and staff: "Histological Methods," by Professor S. H.
Gage; the technique portion of "Vertebrate Histology" by Gage
and Kingsbury. In the following presentation the latter of these two
has been largely drawn on and certain loose sheets of technical direc-
tions by S. H. Gage incorporated. In addition the writer is particu-
larly indebted to The Microscope, by S. H. Gage; The Microtomists
Vademecum, by A. B. Lee, and Die Enzyklopaedie der mikrosko-
pischen Technik, together with a large number of original articles,
some of which are included among the list of References. When
referred to in the text, the reference is indicated by a bracketed
number. Acknowledgment is also due to Instructors in the Depart-
ment, J. A. Badertscher and H. M. Kingery. Their names in
parenthesis indicate methods particularly due to them.

B. F. KINGSBURY.

CARPENTER & Co.

1915

CONTENTS.

Page

INTRODUCTION 5

FIXATION 7

Fixers, List of . 11

ISOLATION 16

SECTIONING & IMBEDDING 19

Schema for Imbedding 21

The Paraffin Method 22

The Celloidin Method '. 26

The Freezing Method 33

STAINING 34

Stains, List of 38

Preparation for Staining 46

Schema for Staining 51

MOUNTING, (Sealing, Labeling) 52

Slides and Covers, Knife 57

SPECIAL METHODS 59

The Cell 59

Connective Tissue 61

Muscle 66

The Nervous System 68

The Blood 79

Fine Injection 82

Silver Nitrate Impregnations 83

Histo-chemical Methods 84

REFERENCES 90

INDEX 93

INTRODUCTION.

Very few structures of the animal organism can be adequately
examined microscopically without being first subjected to a prepara-
tory treatment involving in many cases the employment of compli-
cated methods. Save in the case of the body fluids and certain
membranes, animal tissues are bulky, more or less opaque, and
therefore unsuited for examination under the microscope which
requires surface or thin layers of substance. Examination is made
possible in such cases in one of two ways, — the elements composing
the structure may be separated from each other, or thin slices may be
prepared.

The above, however, presents but the grosser aspect of the neces-
sity of preparation of animal tissues for examination with the micro-
scope. The histological analysis of bodily structure makes further
demands on the refinement of methods. Treatment with chemicals
and stains (Fixation and Staining) has for its purpose not only the
preservation and delineation of structure, but its identification by
means of more or less definite chemical (physical) reactions. The
goal from this side of histological technique is an analysis from the
chemico-physical as well as the morphological aspect and the inter-
pretation of morphology in terms of physiology. Increase in our
knowledge of the finer structure of the body in the past has been, as
advance in the future will be, accompanied by and dependent on the
application of a more exact technique along these lines; while for
those who aim to do practical work in histology and pathology a
mastery of the more important methods is indispensable.

Furthermore, in working with chemically altered structure
there is always the danger of losing sight of the conditions existent
in the living protoplasm. It is well, therefore, in addition to study
structure in the living or fresh state, as little altered from the natural
as may be. There is also very desirable the acquisition of skill in
the application of simple methods which require neither expensive
apparatus nor expenditure of time, — methods which while they
may not advance knowledge, serve often to meet the needs of a
preliminary examination or rapid clinical diagnosis.

Of the multitudinous methods employed in microscopic work
only those are here given which meet the requirements for a general
working knowledge in histology. In special investigations, it is

necessary to make a study of the particular technical needs of the
problem, and for this it is well to consult the larger works on technique
of which may be mentioned the Encyclopedia of Microscopical
Technique [6]; The Microtomist's Vade-mecum by A. B. Lee [30];
Physiological Histology, by Gustav Mann [36]; Mallory & Wright,
Pathological Technique [35]. The books of Gage, Hardesty and v.
Kahlden-Gierke will also be found valuable for consultation.

General Histological Technique involves then :

A. Examining fresh, by either B. or C. Advantageous or necessary

when haste is required, or in examining the tissue alive.

B. Isolation or Dissociation. Separating out the elements composing

a tissue by (a) teasing or (b) treatment with reagents and
teasing.

C. Cutting thin sections of the tissue or organ.

For C. are generally necessary: .
1. Fixing the tissue (§ § 1 — 34). Hardening.
%. Sectioning by one of the following methods :

(a) Free-hand, without an imbedding mass, or

(b) With an imbedding mass, as

(1) By the Paraffin method (§ 50) , or

(2) By the Celloidin method (§ 61) , or

(3) By the Freezing method (§ 73) .

D. Staining; — to outline and differentiate the structure, or pick

out definite chemical substances.

E. Mounting; — for examination under the microscope and per-

manent preservation.
In addition, — numerous

F. Special Methods and methods for the

G. Histo-chemical Analysis of structure must be frequently applied.

FIXATION.

§ 1. Fixation is one of the fundamental processes in the ex-
amination of plant and animal tissues. A fixer may be defined as
a fluid (or gas) into which the living or at least very fresh tissue is
placed in order to preserve the structure of its elements as nearly as
possible as in life. Living tissue when allowed to die and remain
undisturbed, gradually loses the structural features it had in life
and undergoes disintegration and decay. Fixation depends upon
physico-chemical processes wherein the chemical constituents of the
tissue are thrown down in situ by being rendered insoluble in some
form or represented by substitution products; the whole being at-
tended by as little distortion as possible. It should be appreciated
that the chemical constituents of cell protoplasm and of the tissues
are numerous and diverse in their chemical and physical properties,
so that a universal or ideal fixer not only does not exist but is logically
inconceivable. The bulk of protoplasm and the tissues is protein
and the basis of fixation in general is the precipitation or coagulation
of these chemical substances. It should be remembered however
that fats (lipoids*) are a constant though variable component of
cytoplasm; that carbohydrates (glucose, glycogen, etc.) are usually
present in small amounts, and that the products of cell activity such
as secretion-granules, zymogen, etc., may be quite distinctive in
their physico-chemical properties. Within the cell a certain "antag-
onism" exists between nucleus and cytoplasm, the former oxidative,
the latter reducing, requiring often somewhat different fixation
conditions.

For the best results, the fixer should be chosen with a view to
the preservation of some particular part or constituent, though a
number of general fixers are very serviceable for routine work.
Rational fixation will depend upon a detailed knowledge of the
chemical and physical properties of the constituents that it is de-
sired to preserve and their interaction with the chemicals of the
fixer. In many respects, rational fixation still awaits further knowl-
edge of the physics and chemistry of fixation.

§ 2. The chemicals of most service as fixers are: (1) osmic
acid (osmium tetroxid), (2) platinic chlorid, (3) picric acid (tri-

*Lipoid, while not a good chemical term is one that is quite useful in histology,
to include fats, fatty acids, phosphatids, cholesterol, etc., substances that have
the same solvents and which are found associated in protoplasm.

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system (§ § 199, 201, 204) are; (IQ) Muller's fluid, (20) Erlicki's fluid
(21) Potassium dichromate (in aqueous solution).

In the combination of chemicals in fixing solutions such as those
mentioned above, they should be chosen to supplement each other's
actions as far a> po>>ible and correct or counteract their defects. The
combinations must be chosen always with a view to the result desired
and frequently the components and their relative amounts determined
empirically, — by experiment .

§ 7. The following general rules should be regarded in the
fixation of tissues niid organs:

(1). The volume of the fixing fluid used should be large, ex-
ceeding the volume of the tissue at least thirty times. The less
energetic the action of the fixer the greater the amount of fluid to be
employed. When the fluid becomes turbid it should be changed to
fresh at once.

(2). Fix only as small pieces of tissue as possible, or as is prac-
ticable from the results desired. The block of tissue should not
be more than 1 cm. in one dimension, and if possible let it be much
shorter, — only 2 to ,5 mm. In some cases (Flemming's and Hermann's
fluids) much smaller masses are needed (1 to 2 mm. thick). This is
desirable for the rapid and complete penetration of the fixer. Of
course, in the case of entire organs it may not be possible to comply
with the conditions. Fixation by injection may then be resorted to.
Physiological salt solution (§ 35) is first injected through artery of the
organ or part in order to remove the blood, and this is followed by
the injection of the fixing fluid.

In addition to these two general principles, there are four points
to be carefully considered, upon which the excellence in the results
attained depends; they are (a) the fixer chosen, (b) the time of
fixation, (c) the trashing out of the fixer, (d) the hardening in alcohol
and the subsequent treatment.

(a) The choice of a fluid into which the tissue is placed should
be made dependent on (1) a consideration of the particular feature
whose preservation is desired and the degree of excellence of fixation
that is desired or necessary,— whether detail of cell structure or the
structure of the tissue in terms of cells or structural elements be
sought ; ("2 ) the penetrating power and the size of the piece that it is

necessary to have; and (3) the stain that is desired subsequently
which is largely determined by the fixation.*

(b) The time a fixer is allowed to act should be considered in
connection with the character of the fluid and the tissue. Usually
the exact limitation of time is a matter of secondary importance
and the tissue may remain in the fixer indefinitely. In some cases,
however, its disregard affects the results seriously and as a general
rule there is a minimum and a maximum time and between them
an optimum time that should be adhered to.

(c) After the tissue has been in the fixing fluid a proper length
of time, it is necessary that it be washed thoroughly to remove
the fixer from it. Usually this should be done by means of water
or alcohol or both. In general, fixers containing salts insoluble in
alcohol or but slightly soluble, as osmic acid, chromic acid, potassium
dichromate, etc., should be thoroughly washed in water. Fixers
containing picric acid or alcohol should always be removed by
alcohol; mercuric chlorid may be washed out by either water or
alcohol.

Inadequate washing out of the fixer may either seriously affect
the cutting quality of the tissue (if it is to be subsequently im-
bedded), the ease with which it can be stained, or there may be
formed precipitates in the tissue giving illusory effects, distortions,
or at least a dirty appearance to the preparation. Time in properly
washing out a fixer is always well spent, as it is a matter for serious
attention.

§ 8. Resume. In brief, "then: In fixing, take relatively large
amounts of fluid and small pieces of tissue, choose the fixer well with
a view to the tissue and the results desired, permit the fixing to
proceed for a sufficient length of time, and wash out thoroughly.

HARDENING AND STORING.

§ 9. Each fixer has also more or less of a hardening action upon the tissue.
Some fluids spoken of above as fixers were primarily used as hardeners, such as
Muller's fluid or Erlicki's fluid, while with others, e. g., picric acid in aqueous
solution, the hardening action is a minimum. The hardening action of the fixer

*Numerous papers have been written on the nature of fixation and the action
and relative value of the various chemicals used for that purpose. The contribu-
tions of Berg [4], Fischer [7], Mann [36], and v. Telly esniczky [6] may be
particularly mentioned.

is generally supplemented by the subsequent use of alcohols of increasing strengths
(50% to absolute, — 99%), as well as in preparation for the paraffin and celloidin
methods of imbedding. In fact, with modern methods of imbedding excessive
hardening of the tissue is not necessary and indeed often should be avoided as
affecting the cutting quality of the tissue. Tissue after fixation has been com-
pleted may be stored in 82 or 95% alcohol, or (better) imbedded at once (§ 47).

Alcohols. 50%, 67%, and 82% alcohols form a series of increasing strengths
sufficient for most purposes. They may be prepared from 95% alcohol by taking
—(a) for 50% alcohol; 95% alcohol 1 part, water 1 part; (b) for 67% alcohol;
95% alcohol 2 parts, water 1 part; (c) for 82% alcohol, 95% alcohol 5 parts,
water 1 part. Dilutions of other strengths may easily be prepared as desired
from 95% alcohol. 95% (94%) alcohol and absolute alcohol are necessary in
imbedding by the paraffin and celloidin methods (§ 49-).

§ 10. Stock Solutions. It is advantageous to have on hand strong solutions
of the chemicals employed as fixers and stains. Where feasible, 10% solutions
are most c6nvenient. The following are the more important: In aqueous
solution; — 10% potassium dichromate, 10% copper dichromate, 10% chromic
acid, 10% platinic chlorid, 40% formaldehyde (formalin), 4% sodium sulphate,
4% copper sulphate, 2% osmic acid, saturated solution of mercuric chlorid,
saturated solution of picric acid, 95% alcohol, absolute alcohol, etc., as well as
the strong acids, stock staining solutions, etc.

FIXERS
i

§ 11. Mercuric chlorid. One may employ (a) a saturated solution in
water or (b) a saturated solution in normal salt solution, with 1 to 5% glacial
acetic acid. Water will dissolve about 5%, normal salt solution about 12%
of the mercuric chlorid. This is a good fixer, especially when the piece is small.
It fixes as soon as it penetrates and is apt to make tissue brittle if it is left too
long. Staining after it is brilliant. The larger percentage of acetic acid is,
perhaps, to be preferred for most histological objects.

Fix the fresh tissue l/2 to 24 hours according to the size of the piece. Remove
to 67% alcohol for 1 or 2 days, 82% alcohol several days, changing often. The
82% alcohol should contain enough tincture of iodin to give it a yellow color,
and fresh tincture added or the alcohol changed when the yellow color of the iodin
in the alcohol is lost. As long as the alcohol is decolorized, washing should be
continued, since it is important that the mercuric chlorid be all removed from the
tissue; otherwise precipitates will form in the preparation after it is mounted
or before, and spoil the result. Wash out in alcohol thoroughly and carefully.
Almost any stain may be employed after a mercuric chlorid fixation.

§ 12. Zenker's fluid. Formula: Potassium dichromate, 2.5 grms. ; sodium
sulphate, 1 gram; mercuric chlorid, 5 grms.; water, 100 c.c.; and add before
using, glacial acetic acid, 5 c. c. A stock solution, without the acetic acid should
be kept on hand. This is a well balanced fixer; the potassium dichromate seems
to check the brittleness that the mercuric chlorid would cause and improves the
fixation of the cytoplasm while the mercuric chlorid and acetic afford a good
nuclear fixation. It is distinctly better than mercuric chlorid; staining after it,
however, is apt to be a little more difficult and not as brilliant as with mercuric

chlorid fixation. Its penetration is surprisingly good. The sodium sulphate is
probably unnecessary. Use of physiological (0.6%) salt solution instead of water
to prevent the formation of precipitates or the addition of a small amount of salt
to the fluid when made up for use, is recommended by some.

Fix in Zenker's fluid 12 to 48 hours, wash well in water, running or frequently
changed, 12 to 48 hours, to remove the dichromate; transfer to 67% alcohol
for 1 or 2 days, 82% alcohol for several days, keeping in the dark while in the
alcohol. To the 82% alcohol add a drop or so of tincture of iodin, adding fresh
iodin or changing the alcohol when the color is lost. This should be continued as
long as the iodinized alcohol is decolorized in order to avoid a precipitate of the
mercuric chlorid in the tissue. Avoid, however, adding an excess of iodin, since
it will affect the staining quality of the tissue. Stain as you wish.

§ 13. Kelly's fluid. (Zenker-formol). Formula: Zenker's fluid, with
formalin substituted for the acetic acid. Potassium dichromate, 2.5 grams;
sodium sulphate, 1 gram; mercuric chlorid, 5 grams; formalin, 5 or 10 c. c., to be
added before using. Valuable for the fixation of cytoplasm and cytoplasmic
granules, where the acetic acid is to be avoided. Make up from the Zenker's
stock by adding the formalin.

Fix 6 to 24 hours or longer, wash in running water 6 to 12 hours, alcohols.
If it is desired to give a longer mordantage in dichromate, transfer to Muller's
fluid or other simple dichromate solution.

§ 14. Dichromate-acetic. (Tellyesnicky's fluid) Potassium dichromate,
3 grms. ; glacial acetic, 5 c. c.; water, 100 c. c. This fluid gives good preservation
of nuclei and of the cytoplasm, the acetic acid checking the bad effects of the
dichromate. A simpler fixer than Zenker's fluid.

Fix 1 to 14 days according to the size of piece and the object. Wash in
running water 12 or 24 hours, and pass up through 25, 50, 67, and 82% alcohols,
12 to 24 hours in each.

§ 15. Formol-Dichromate (Orth's fluid). Formula: Potassium dichro-
mate, 2.5 grms. ; sodium sulphate, 1 grm.; water, 100 c. c. (i. e., Muller's fluid);
formalin, 10 c. c. A good fixer of cytoplasm, but an indifferent one for nuclear
detail. It has been especially used in the case of the nervous system (§ 199).
Other proportions of formalin and dichromate may be taken, the sodium sulphate
being omitted. It should be freshly made up as it soon deteriorates.

Fix 3 to 24 hours or longer, wash in running water 12 to 24 hours; 50, 67,
82% alcohols, 1 day in each. If a long fixation is given, the fluid should be
frequently changed.

§ 16. Copper dichromate-sublimate-acetic. Formula: 10% copper dichro-
mate, 1 part; 4% copper sulphate, 1 part; sat. solution mercuric chlorid, 2
parts; glacial acetic acid, rV to 5%, according to need. Similar to Zenker's
fluid in its general properties and should be used in the same way. Excellent as a
preserver of cytoplasmic granules of lipoid nature (§ 177-9). A further mordan-
tage in 2.5% copper dichromate is often advantageous (3 to 4 days).

By the substitution of formalin (5 to 10%) for the acetic acid, a fixer compar-
able to Kelly's fluid (§ 13) is obtained.

§ 17. Picro-aceto-formol. (Picro-formol). Formula: Picric acid, satu-
rated aqueous solution, 75 c. c.; formalin, 25 c. c.; glacial acetic acid, 4 c. c.

A delicate fixer useful in some cell work and with small objects. The formalin
may be omitted (Picro-acetic).

Fix 3 to 24 hours, transfer to 67% alcohol, 1 day, 82% alcohol several days
changing the fluid frequently as it becomes yellow. Leave in the alcohol until
the picric acid has been well washed out.

§ 18. Hermann's fluid. Formula: 1% aq. sol. platinic chlorid, 15 parts;
2% aq. sol. osmic acid, 4 parts; glacial acetic acid, 1 part; or you may take
10%%aq. sol. platinic chlorid, 3 parts; 1% aq. sol. osmic acid, 16 parts; glacial
acetic acid, 2 parts; water, 19 parts. This is generally recognized as one of the
finest fixers known, and it is also the most expensive. The form and structure
of cells are well preserved. It should only be employed, however, with very
small pieces of tissue, and is to be used especially when cell structure is to be
studied. Fat and the myelin of nerve fibers are stained black.

Fix in this 1 to 24 hours (or longer — days or weeks are used by some), wash
well in water (running or frequently changed) 2 to 24 hours, and then place
in 67% and 82% alcohols, 12 to 24 hours in each. In using this fluid, the smaller
the pieces taken the better the fixation will be, and in order that it be possible
to obtain a good stain afterwards tissue should not be over-fixed and the fixer
should be thoroughly washed out. If there is a blackening of the tissue, or a
precipitate in it, both may be removed by treatment of the sections on the slide
with a 10 or 20% solution of hydrogen dioxid in 67% alcohol, or with perhydrol
(Merck). Employ after Hermann's fluid, as stains, Heidenhain's iron hema-
toxylin, Delafield's hematoxylin, safranin (as a red stain), or gentian violet (as a
blue stain), or neutral stains.

§ 19. Flemming's fluid (Chrome-aceto-osmic) . Formula: 1% aq. sol.
chromic acid, 15 parts; 2% aq. sol. osmic acid, 4 parts; glacial acetic acid,
1 part; or, 10% aq. sol. chromic acid, 3 parts; 1% osmic acid, 16 parts; glacial
acetic acid 2 parts; water, 19 parts. This is a fine fixer and in most cases gives
as good results as Hermann's, and is not as expensive. It browns tissue less,
and while it blackens fat, does not blacken the myelin of myelinic nerve fibers
as does Hermann's. It should be employed in general in the same cases and in
the same way as Hermann's fluid, and is especially useful in the preservation of
free fats and lipoids (§ 227) . The presence of chromic acid gives it a distinct value
as a cytoplasmic fixer.

Fix tissue 1 to 24 hours (or longer); wash well in running water 2 to 24
hours; place in 67% and 82% alcohols, 12 to 24 hours in each. Bleaching of
the sections may be necessary, as with Hermann's fluid. Take only very small
pieces of tissue. Employ the same stains as with Hermann's fluid.

The reduction of the acetic acid to 3 or 4 drops (about TJT%) is advisable when
cytoplasmic granules of lipoid composition (§ 178) are to be preserved. The fluid
may then be spoken of as Benda's fluid.

§ 20. Mercuro-nitric (Gilson's fluid, modified). Formula: Nitric acid (46°,
sp. gr. 1.456, 80%), 15 c. c.; Glacial acetic acid, 4 c. c.; Mercuric chlorid, 20 grms.;
95% alcohol, 60 c. c.; distilled water, 920 c. c. A good fixer, especially useful
where rapid penetration is a factor, and as a fixer of cells rich in yolk (amphibian
ova, etc.).

Fix 12 to 24 hours, transfer to 50, 67, and 82% alcohols. lodin should be
used to ensure the removal of all the mercuric chlorid (see § 11).

§ 21. Picro-nitric. Formula: Distilled water, 95 c. c.; nitric acid (strong),
5 c. c.; picric acid, to saturation. Only to be recommended for eggs rich in yolk
and where a high power of penetration is required.

Fix 4 to 12 hours, transfer to 67% alcohol, 1 day 82% alcohol several days
changed frequently. It is necessary to wash out thoroughly which is accomplished
however, with difficulty.

§ 22. Perenyi's fluid. Formula: 10% aq. sol. nitric acid, 4 parts; 95%
alcohol, 3 parts; K% aq. sol. chromic acid, 3 parts. An embryological fixer
of much value. It is also serviceable for general work.

Fix tissue for 4 to 5 hours, place in 67% alcohol for 1 day, 82% alcohol several
days.

§ 23. Alcohol. 95%, 67 or 70% alcohol. The employment

of most of the fixers so far mentioned requires the expenditure of considerable
time, rendering them inapplicable or unsuitable in many instances. 95% alcohol
itself answers admirably for most histologic objects, fixing well, hardening and
likewise dehydrating (§ 49) preparatory to imbedding in paraffin or collodion,
affording thus a considerable economy of time. It is also most serviceable in
pathological tissues where the presence of bacteria is suspected. In some instances
67% alcohol answers as well or better, while in other cases absolute (99%) alcohol
should be employed.

Fix in 95% alcohol for 1 to 3 days, changing two or three times, after 3 or 4
hours and after 24 hours. The tissue will probably be found to be dehydrated
and ready for the next step of the imbedding process (§ 51 or 62). Stain as you
wish.

§ 24. Alcohol-acetic. The addition of 5% glacial acetic acid to 95%
alcohol or absolute alcohol increases the penetrating power and improves the
cutting quality of objects containing much connective tissue. The following
formula possesses high penetration and is sometimes useful:

§ 25. Alcohol-acetic (Carnoy's fluid, 3-1). Formula: Glacial acetic acid

1 part, 95% or absolute alcohol, 3 parts. Transfer after fixing to 95% or absolute
alcohol, changing each day until the acetic acid is well washed out.

§ 26. Chloroform-alcohol-acetic. (Carnoy's 6-3-1 ). Formula: 95% or
absolute alcohol, 6 parts; chloroform, 3 parts; glacial acetic acid, 1 part. Fix

2 to 24 hours or longer, transfer to 95% alcohol, changed two or three times.

§ 27. Carnoy's Sublimate mixture. The above (§ 26) with mercuric chlorid
added to saturation. A very penetrating and excellent fixer, particularly useful
for nuclear preservation and for embryos.

Fix 1 to 24 hours, transfer to 95% alcohol for 1 day, then to 82% alcohol for
several days, changing daily and adding iodine solution (§11) until the excess
mercuric chlorid is removed.

§ 28. Alcohol-acetic-formol. Combinations such as the following are
useful for general purposes. V. Luko's formula: Alcohol, 67%, or 82%, 100 c. c.
glacial acetic acid, 5 c. c.; formalin, 5-10 c. c.

Fix for 12 to 24 hours or longer; remove to 67% or 82% alcohol for 1 day
or more.

§ 29. Formaldehyde. Solutions of this chemical have been found to be
good preservatives and hardeners and fairly good fixers. It penetrates rapidly,

and preserves the natural transparency and pigmentation of the tissue, making it
valuable for gross anatomy and museum purposes. As a fixer, an aqueous solu-
tion of 2 to 4% strength may be employed, or it may be used, which is preferable,
in conjunction with other chemicals, as picric acid, in picro-formalin, or chromic
acid and acetic acid.

Formalin is a 36 to 40% solution of formaldehyde (gas) in water. A small
amount of formic acid is also present. A 10% solution of formalin, that is a 4%
solution of the formaldehyde is a satisfactory strength for most histological
purposes.

Fix 12 to 24 hours, remove to 67% alcohol for a day, 82% alcohol one to
several days. Stain as you wish.

§ 30. Osmic acid. A very useful as well as expensive reagent and somewhat
difficult to use. It is generally employed as a fixer in conjunction with other
reagents, as in the mixtures (§ § 18 and 19). When used alone as a fixer weak
solutions are generally best — A to 1%. It penetrates slowly and it "over-fixes"
cells very easily, obscuring detail and giving the parts a homogeneous, glassy
appearance. Over-fixed cells cannot be stained, or with great difficulty. More
or less blackening of the protoplasm also occurs. It may be used chiefly to demon-
strate fat, which is blackened by it, and the zymogen of pepsin and trypsin, which
it preserves and browns slightly.

Fix small (about 2 mm. thick or less) pieces of tissue in 1% osmic acid for 6 to
12 hours, wash well in water (running or changed frequently) for 12 to 24 hours,
and place in 67% and 82% alcohols. It is somewhat difficult to prevent pure
osmic acid of this strength from over-fixing the tissue, and cell detail is generally
lost, though the form of cells is well preserved.

§ 31. Nitric acid. A 10% solution of nitric acid is serviceable in fixing the
blastoderm of the chick.

§ 32. Muller's fluid. Formula: Potassium dichromate, 2.5 grams; sodium
sulphate, 1 gram; water, 100 c. c. Make up from stock solutions. This fluid is
more of a hardener than a fixer; it should be avoided (as likewise Erlicki's fluid
and potassium dichromate) when the preservation of nuclear structure is desired.
Staining after its use is sometimes difficult. It is, however, occasionally useful
for general work, although such formulas as Zenker's fluid or Kelly's fluid are
generally to be preferred.

Place the object in an abundance of the fluid and harden for from 1 to 8
weeks, changing the fluid at first each day. In general, 10 to 14 days will be
sufficient. Wash in running water for 24 to 48 hours or longer, remove to 67%
alcohol for 1 to 2 days, 82% alcohol several days. Keep in the dark while in the
alcohols, and change to fresh when the fluid is colored yellow. Tissue hardened
in Muller's fluid cuts well, and it is useful in preparing sections of large organs, or
organs with much connective tissue. Its chief usefulness is, however, in the study
of the nervous system (§ 199-).

§ 33. Erlicki's fluid. Formula: Potassium dichromate, 2.5 grams; copper
sulphate, 1 gram; water, 100 c. c. Make up from stock solutions. This is quite
similar to Muller's fluid in its action and results, save that its action is more rapid
and stronger. Therefore, it had better be employed with smaller objects, and
allowed to act only 2 to 14 days. Otherwise, employ like Muller's fluid.

§ 34. Potassium dichromate. 2%, 3%, and 5% aqueous solutions. This
is quite similar to Miiller's fluid in its action, and may be employed in the same
cases. It is generally used for the central nervous system.

Harden in an abundance of the solution for 2 to 8 weeks, beginning with
the 2% solution for 2 to 6 days, 3% solution 1 to 3 weeks, 5% solution 1 to 3
weeks. Wash out in running water 24 to 48 hours. Place in 67% and 82%
alcohols several days, keeping in the dark meanwhile, changing when the alcohol
is colored.

ISOLATION.

§ 35. One of the simplest ways of examining the structure of
a tissue is the separation from one another of the structural elements
composing it, thus permitting its analysis. Likewise, for a correct
conception of the forms of the cells and fibers of the various tissues
of the body, one must see these elements isolated and thus be able
to inspect them from all sides. It frequently occurs also that isola-
tion is not quite complete and one can see in the clearest manner
the relations of the cells or fibers to one another.

In the employment of this method the tissue may be taken
fresh and isolation accomplished by teasing with needles or similar
instruments; or it may be treated with media which will serve to
render teasing partially or entirely unnecessary. In such cases
simply shaking or gently tapping the preparation will often suffice.
In many instances it is desired to examine the tissue while the
elements are still alive, as, for example, in the study of ciliated
cells, and recourse must be had to some "normal," "indifferent"
medium. Best of all is the medium with which they are bathed dur-
ing life — in the case of tissue from the animal body, blood serum,
the aqueous humor of the eye, liquor amnioticus, or, as an artificial
substitute often more convenient if reagents are to be used subse-
quently, physiological or normal salt solution, being a 6/10 to 9/10%
solution of common salt (sodium chlorid) in distilled water. 6 /10%
is suitable for use with Amphibian tissue; 75/100% is normal for
reptiles and 9/10% best for mammals. Other normal physiological
solutions may in some cases be preferable, such as Ringer's solution,
Locke's solution, etc.

§ 36. The use of chemical solutions to facilitate isolation of the
elements is called Maceration. The chemical agents or solutions for
isolating are, in general, the same as those used for fixing and hard-
ening. But the solutions are only about one-tenth as strong as for

fixing and the action is very much weaker and requires from one or
two hours to as many days. In the weak solution the cell cement or
connective tissue is softened so that cells and fibers may be separated
from one another, and at the same time the cells are preserved. In
other words, a weak fixing action is retained while the hardening
action is reduced on dilution. The time required for the action of
the dissociator varies inversely as the vehemence of the fixer and
the density of the tissue, 2 to 3 hours to several days. In fixing and
hardening, on the other hand, the cell cement, like the other parts of
the tissue, is made firmer. It is better also to dilute the fixing agents
with normal salt solution than merely with water [15]. Those
chemicals that are "cytoplasmic fixers" such as potassium dichro-
mate, Muller's fluid, formaldehyde, osmic acid, appear to be especi-
ally useful in the dissociation of epithelia.

For the isolation of muscle, alkalis or mineral acids, which soften
or dissolve the connective tissue are to be employed. Horn, nail and
hair require strong mineral acid or (usually) weak alkali.

§ 37. Of the many maceration fluids or dissociators the follow-
ing may serve most of the needs of histology: (1) Muller's fluid dis-
sociator; (2) formaldehyde dissociator; (3) osmic acid (1/10%);
(4) sulphuric acid (strong); (5) nitric acid dissociator; (6) caustic
potash dissociator.

§ 38. Muller's fluid dissociator. Formula: Muller's fluid, 1
part; physiological salt solution, 9 parts (i. e., potassium dichromate,
2.5 grams; sodium sulphate, 1 gram; sodium chlorid, 9 grams;
water, 1,000 c. c.)% [15].

This is a good dissociator for epithelia, including glands. Dilution
decreases the hardening action of Muller's fluid as is shown especially
on the cell-cement, — hence its dissociating action. Considerable
latitude in time is allowed in the use of this dissociator; 12 hours
being often sufficient, although a stay of several days in the dis-
sociator usually does no harm.

§ 39. Directions for use. In the employment of this fluid for
the isolation of epithelial cells, proceed as follows:

Place the tissue covered with the epithelium which it is desired
to isolate in the dissociator in a shell vial or dish, where it may re-
main from 2 to 3 hours to 2 to 3 days; for the epithelium of the tra-
chea, intestines, etc., the action is sufficient in 2 to 3 hours, although
good preparations may be obtained after two days or more. For the

stratified epithelia, like those of the skin, mouth, etc., it may require
1 to 3 days for the most satisfactory preparations. After the tissue
has remained in the dissociator a sufficient time, scrape the epithelial
surface gently with a scalpel and place the scrapings on a slide in a
drop of dissociator; cover and examine. If one proceeds after two
hours or so, probably most of the cells will cling together, and in
the various clumps will appear cells on end showing the tops or
bases, and other clumps will show the cells in profile.

Tap the cover gently with a needle-holder or other light object
in order to separate the cells from each other more completely.
Many fully isolated cells as well as cells in groups will be seen. Ex-
amine carefully.

§ 40. Staining. Scrape gently the epithelial surface in a fresh
spot and place the scrapings on the slide in a drop of eosin (§ 114)
or congo red (§ 118). Mix well so that the stain can penetrate. If
for temporary examination, cover immediately and examine as be-
fore. For methods of making permanent preparations of dissociated
cells and mounting in glycerin, see § § 151—.

§ 41. Formaldehyde Dissociator. Formula: 40% formaldehyde
(formalin), 2 c. c.; physiological salt solution, 1,000 c. c. (i. e.,
.08% sol. of formaldehyde in normal salt solution). This is a good
general dissociator and as such may be employed instead of Miiller's
fluid dissociator. It is especially serviceable in the isolation of the
nerve cells of the brain and spinal cord [13], and for its use with
that material, see § § 151—. It is however excellent for the dissocia-
tion of epithelia.

§ 42. Osmic acid. A 1-10% solution of osmic acid is a valua-
ble dissociator, especially serviceable in the isolation of nerve-fibers,
my clinic and amyelinic, and when fat is present, since fat and the
myelin of myelinic nerve-fibers are blackened by it. Twelve to twen-
ty-four hours generally affords sufficient time for it to act. (§ 193,

§ 43. Sulphuric Acid. This is used in the concentrated form
as a dissociator of the epithelial cells of hair, horn, and nail. If
heated, a few minutes suffice; employed cold, a day or two may be
required.

§ 44. Nitric acid dissociator. [8] Formula: Strong nitric
acid, 20 c. c.; water, 80 c. c. See § 189a. This fluid is employed in
the isolation of muscle fibers, both striated and plain.

§ 45. Caustic potash dissociator. [8] Formula: Caustic
potash, potassium, hydroxid (in sticks), 35 to 40 grains; distilled
water, 65 or 60 c. c. This solution will be used for the isolation of
cardiac muscle 'cells', although it may be used for striated or plain
muscular tissue, or as a general dissociator. It may also be employed
for isolating the cells of hair, horn or nail, either full strength or
diluted.

Ten to fifteen minutes or longer will be enough for the isolation
of heart muscle (§ 189 6) ; 2 to 3 days may be required for the ade-
quate maceration of cornified epithelial cells.

SECTIONING.

§ 46. In addition to the examination of tissue by the separa-
tion of the component elements — isolation — it may be examined
microscopically after cutting very thin slices or sections of it. This
may be done free-hand or by means of a special machine, a micro-
tome, and with or without an imbedding and supporting mass.

For the finer work an imbedding mass and a microtome must
be used. Free-hand sectioning without an imbedding mass, and
even without previous fixing is, however, necessary or advisable
when economy of time is a desideratum, as in clinical examinations
of tissue, when one wishes to study the part alive or fresh (i. e.,
not treated with reagents), or if the reagents necessary for fixing
and imbedding destroy or alter the structural features to be investi-
gated.

The ability to recognize tissues and organs unaffected by re-
agents and without the employment of methods involving the ex-
penditure of time and effort is very desirable, especially in patho-
logical work, when haste often forbids the employment of the finer
methods, were facilities for their use available, as in some cases
they are not. Great skill in the use of simple tools may be gained
and counts for much. It should be remembered also that the greater
one's knowledge of a structure the less the need to resort to special
methods of preparation for its recognition.

IMBEDDING METHODS.

§ 47. When the consideration of time is not so important and
finer results are more to be desired, the sections should be prepared

according to some method in which an imbedding mass is used. The
interstices of the tissue are completely filled with some substance
that wrill give support and greater consistency and homogeneity to
the tissue, and thereby enable the cutting of much thinner and more
perfect sections.

There are three methods that are generally employed, (a) the
Paraffin method, (b) the Celloidin (Collodion) method, and (c) the
Freezing method; the imbedding masses to fill the spaces being re-
spectively paraffin, collodion and a congelation mass,— ice. The last
is the simplest; it requires less expenditure of time, fewer reagents,
and its results are in some ways the crudest. As in cutting free-hand
sections without imbedding, the freezing microtome should be em-
ployed when haste is necessary and finer detail unimportant, as in
clinical work. The two remaining methods may be employed in most
cases and give good results. A choice between them mUst be deter-
mined by the special requirements of the case and a consideration of
the differences of the two methods as set forth in tabular form below :

CELLOIDIN. PARAFFIN.

No heat required. Heat required.

Sections relatively thick ; 10 to 25 mi- Sections relatively thin ; 1 to 10 microns

crons or more. or more.

Imbedding mass usually not removed. Imbedding mass removed.

Sections usually cut wet (with alcohol or Sections cut dry.

oil).

Knife oblique. Knife usually set at right angles.

Cutting stroke slow. Cutting stroke usually rapid.

Form of the organ better preserved. Usually more or less distortion.

Imbedding requires more time. Imbedding requires less time.

Celloidin stains with basic dyes.

In general, better for larger specimens. Better for smaller objects.

§ 48. Despite the differences, the two methods may in most
cases be used interchangeably. The advantages of the paraffin
method are (1) the facility of its use and the ease with which thin
sections may be obtained and its adaptation to serial sectioning.
Celloidin is particularly useful when heal is injurious or the paraffin
solvents (clearers) dissolve out substances which it is desired to
preserve. Its main defect is the readiness with which the celloidin
stains with basic stains, particularly such as Iron Hematoxy lin and
the coal-tar dyes (safranin, gentian violet, methylene blue, etc.)
The celloidin may, however, be dissolved out. See § 136.

§ 49. The following table will indicate the steps in the employ-
ment of the two methods:

Paraffin Method

21
Living tissue

(

Fixing

(§ § i-)

Washing

i
Alcohols (§ 9)

(50%, 67%, 82%)<-

Dehydration
(95%-99% alcohol)

-Staining in toto
(§79)

Celloidin Method

Toluene, 1 part
Abs. Alcohol, 1 part.
(Ihr. to 1 day)

1
Pure toluene,

(1 hr. to 1 day)

Toluene, 1 part,
Paraffin, 1 part.
(2 hrs. to 2 days)

Pure Paraffin

(2 -to 24 hrs.)

Clearing

Ether-alcohol
days)

. . Infiltration

Imbedding paraffin,

mass cooled. Imbedding.

Paraffin sections
cut.

Sectioning

Thin celloidin

(2% solution)

(1-14 days)

Thick celloidin
(6% solution)
(1-14 days)

1
Thick celloidin

(12% solution)
(1-14 days)

! -

Celloidin mass
hardened in
chloroform

Celloidin block
clarified or
placed in 82%
alcohol

Celloidin sections
. .cut

THE PARAFFIN METHOD.

§ 50. As seen by the above scheme, the aim is to fill all the
interstices of the tissue with paraffin of the right degree of hardness
to have it cut well. Paraffin is not soluble in water or alcohol, but
is soluble in a number of fluids which in turn are miscible with alco-
hol. Hence the following steps are necessary: (1) the tissue must
be first water-rid, thoroughly dehydrated with strong alcohol; (2)
freed from the alcohol, cleared by a fluid that mixes with melted
paraffin which (3) takes the place of the clearer in the tissue, infil-
trates it, filling the spaces; (4) finally, the tissue is imbedded in
paraffin of the right degree of hardness, the mass cooled, and it is
ready (5) to cut, or section.

§ 51. Dehydration. After the various steps pertaining to the
fixing and hardening (§ § 7, 9) of the tissue have been properly
pursued it may be stored in alcohol of 82% to 95% strength depend-
ing on the tissue and its purpose. The dehydration necessary
in imbedding may be accomplished by immersion in alcohol of 95%
strength. For most work it is perhaps better to employ stronger
(absolute) alcohol. If xylene or toluene are used for clearing absolute
alcohol must be used.

Immerse small pieces 2 to 3 mm. in diameter for at least 6 to 8
hours in 95% alcohol changed once or twice. A longer time, even
days, usually does no harm and is preferable to ensure complete
dehydration. For larger pieces of tissue or entire organs a corres-
pondingly longer period of dehydration should be employed, a several
days' stay, with the alcohol changed daily, being often advisable. In
any case, dehydrate thoroughly, changing the alcohol 1 to 3 times, the
last change usually being to absolute alcohol. Let the tissue dehy-
drate for a longer rather than a shorter period of time.

§ 52. Clearing. The alcohol must next be replaced by some
solvent of paraffin which is miscible with alcohol, — a step spoken of
as clearing. Toluene* is one of the most serviceable clearers, although
for special purposes other media such as xylene, cedarwood oil,
bergamot oil or chloroform may be preferred. Toluene (and xylene)
mix well only with absolute alcohol, hence the dehydration needs to
be thorough, and the clearing and infiltrating is best accomplished by
a number of steps. After the absolute alcohol, the tissue is placed (1)

*Benzene, toluene, xylene, etc.; the terms adopted by American chemists.
In Europe the same substances are designated xylol, benzol, etc.

in equal parts of toluene and absolute alcohol for 1 hour to 1 day; (2)
this is replaced by pure toluene for an equal period when the tissue
will be clear and translucent, — except, of course, such as is dark in
color.

Xylene may be used in place of toluene in nearly all cases. Steps 1 and 3 may
often be omitted with very small objects. In some cases it is well to clear with
cedarwood oil first (§ 54) and then transfer to toluene (or xylene), toluene paraffin,
etc. Familiarity with the tissue and the reagents will permit considerable depart-
ture from a fixed line of procedure.

§ 53. Infiltration. After the tissue is completely cleared in
the toluene, remove it to a dish of (3) melted infiltration paraffin
1 part, toluene 1 part, and set it in a warm place (about 38° C.), so
that the paraffin may remain melted and the toluene slowly evaporate.
After several hours or even days with very large pieces place the tissue
in (4) pure paraffin in the paraffin oven for 2 to 24 hours, depending on
the size of the piece. Quite large pieces may be left longer ; with them
one or two changes to fresh infiltration paraffin may be necessary.
The melted paraffin replaces the toluene, filling in the interstices of the
tissue. Paraffin melting at about 52-54° C. is used. It is best not to
expose to a higher temperature than is necessary, or for a long period
of time as the heat tends to shrink and toughen, especially if the dehy-
dration (and consequently the clearing) have been incomplete; this
is particularly true of organs rich in connective tissue. The paraffin
oven wTill be maintained at a temperature of 54-5° C.

§ 54. Other Clearers. Cedarwood oil is a good clearer. It will clear from
95% alcohol if the dehydration has been carefully done. The tissue should be left
in the cedarwood oil until it sinks and the alcohol currents have ceased to rise from
it. The steps are given below. Thickened cedarwood oil such as is used for
immersion objectives has been found to give excellent results with small objects
and is to be recommended for some cell work. Chloroform gives excellent results
but it penetrates (clears) slowly, so that it should be used only with small pieces
of tissue. Chloroform or thickened cedarwood oil is to be employed with objects
in which it is desired to preserve fat which has been blackened by osmic acid
(§ 227).

CHLOROFORM. CEDARWOOD OIL.

Alcohol (97-99%) . 95% or absolute alcohol.

Chloroform, until it sinks below the sur-
face.

Chloroform and paraffin, equal parts,
for 4-48 hours, at incubator tempera-
ture (38 C.) . cedarwood oil.

Pure paraffin, in the paraffin oven. Pure paraffin, in paraffin oven.

§ 55. Imbedding. It is best to use fresh paraffin for imbed-
ding and sometimes with a melting point higher than that of the in-
filtration paraffin, — 52 to 54° C. paraffin, answers well in a room of 19°
to 20° C., and will be generally used. If the cutting is to be done in
a room of lower temperature, a softer grade of paraffin may be used
for imbedding; if at a higher temperature, a harder paraffin should be
chosen, as when summer work is necessary.

As a general rule, hard tissues require a harder imbedding par-
affin which is also better when very thin sections are desired. Large
sections which usually must also be relatively thicker need a softer
paraffin. It is better to work with a paraffin harder than the room
temperature itself would call for and then regulate the cutting tem-
perature by placing a source of heat such as an electric light nearer
or farther away from the microtome knife.

Make a small paper box, fill it with the melted imbedding paraffin;
float the box on a dish of cold water; transfer to it the tissue from the
paraffin oven, arrange it carefully in the box in the way you wish it
for cutting, and let the mass cool.

§ 56. In imbedding in paraffin observe the following rules:
(1) Take no more paraffin (no larger box) than is needed to form a
mass of convenient size around the specimen. The aim is to have
as homogeneous a mass as possible; paraffin tends to crystallize if
it cools slowly, hence the smaller the mass the more rapidly may it
be cooled. (2) Let the imbedding paraffin when poured into the
box be several degrees above its melting point, and the tissue like-
wise should have an equal temperature. Should the imbedding
paraffin (or the tissue) be too cool it will not set well around the
specimen, and a film of air may be enclosed. On the other hand,
take care that the paraffin is not hot enough to "cook" the tissue,
thereby shrinking it and rendering it hard and tough or ruining it
altogether. (3) Cool the paraffin by floating the box on cold water.
A homogeneous, translucent, paraffin mass can only be secured if it
is quickly cooled. When a film has formed on the surface strong
enough to resist rupture, immerse the block, or drop 95% alcohol
upon the surface. Ice is an advantage in summer imbedding if cold
water is not available. When ice or cold water are not available,
good results have been secured by floating the box on a shallow dish
of (used) ether-alcohol (§ 63). A homogeneous paraffin is only
secured if the paraffin is allowed to shrink in cooling; it is therefore
well to make the boxes as shallow as possible, — that is, much broader

and longer than high. Watch glasses, watch crystals, small tin pans,
etc., may be used as imbedding receptacles.

§ 57. Crystallization of the imbedding mass. Paraffin that has crystallized
is crumbly and will not give good sections. When crystallization has occurred,
it is best to reimbed. Its occurrence is usually due to too slow a cooling of the
imbedding mass, or cooling under conditions that prevent the paraffin from shrink-
ing. It is sometimes due to the presence of impurities in the paraffin, such as
water (?), excess of clearer, etc.

§ 58. Cutting the sections. The essentials for good paraffin
sectioning are (1) well-imbedded tissue, (2) a sharp microtome knife
(or section razor), (§ 173), (3) a room of the proper temperature, and
(4) the paraffin block properly trimmed and arranged in the micro-
tome. Furthermore, tissues fixed and hardened in different ways cut
very differently. Tissue fixed in Hermann's, Flemming's, Mailer's,
Zenker's fluid or Carnoy's fluid, etc., cuts well; alcohol and mercuric
chlorid tissue is more apt to be tough or hard, etc. The different
organs and tissues have of course very different adaptabilities to the
method.

After the imbedding mass is well cooled, remove the paper box
and trim the part containing the tissue in a pyramidal form, two of
the sides at least being as nearly parallel as possible. Clamp the
block of paraffin in the holder of the microtome so that the tissue
will be at the proper level for cutting, being careful to have the par-
allel sides also parallel to the edge of the knife. If a ribbon micro-
tome is used, heat the holder and melt the end of the block upon it.
Cool and place the holder in its place in the microtome, again hav-
ing the parallel sides and the edge of the knife parallel. Use a very
sharp, dry section knife for cutting the sections. Clamp it in the
microtome slightly inclined to the cutting surface of the tissue. If
the temperature of the room is right for the paraffin used, the sec-
tions will remain flat, and if the directions given above for trimming
and arranging the block be observed, they will adhere and thus form
a ribbon. If the room is too cold or the paraffin too hard, the sec-
tions will roll; if it is too warm, the sections will crush or be imper-
fect. If a microtome in which the knife is not fixed, is used, make
the sections with a rapid straight cut as in planing. Do not try to
section with a drawing cut as used in celloidin sectioning. 10^ will
be found the most convenient thickness for the sections, though in
special cases they should be thinner or even thicker. Handle the
sections by means of a camel's hair brush, a needle, or sometimes on a
scalpel handle, when cutting ribbons, etc.

§ 59. Difficulties in sectioning, (a) Rolling of the sections indicates too
hard a paraffin, tissue or both, or too low a cutting temperature; bring the
source of heat nearer or await better conditions, (b) Crushing together of the
sections or wrinkling. Too soft a paraffin, too high a temperature, imperfect
infiltration: alter conditions or reimbed, or reinfiltrate, etc. (c) The sections do
not ribbon. Due usually either to (a) or to a failure to have the two sides of the
block trimmed parallel and set parallel to knife edge, (d) The sections crack
parallel to knife edge: tissue very brittle, (e) The sections are electric. The
electricity is mainly due to friction in the cutting. The hardness of the tissue, —
in some cases due to presence of metallic salts, — or imperfection in the knife edge
seem in many cases to be responsible; although in many instances the cause
appears obscure. A tube of radium or a strong induction coil operating in the
immediate neighborhood is said to obviate the difficulty.

Remember to have the paraffin block trimmed with two sides
parallel and the knife edge parallel to these. Also, do not attempt
to cut if the temperature of the room is too high, — above 23° C.

§ 60. Resume of the method. To obtain as good results as
possible with a certain organ fixed and hardened in a certain way,
the steps must be carefully and exactly followed. Let the dehydra-
tion be complete, clearing thorough, infiltration sufficient; imbed,
carefully observing the three cautions mentioned; and in cutting,
remember to have a sharp knife, a cool room, and the imbedding
block properly trimmed. Success also depends largely on the pre-
vious treatment in the fixer and on the care with which the fixer is
washed out.

Properly employed, the paraffin method is widely serviceable,
being only useless where the tissue is very large, very hard, hard-
ened or injured by heat, or w^here the exact form of a large organ is
important.

THE CELLOIDIN METHOD.

§ 61. A comparison with the paraffin method has already been
given (§ § 47, 48); there may be emphasized here three points: (1)
writh paraffin heat is required, with celloidin no heat; (2) paraffin
must be removed from the sections subsequently, celloidin need not be
and usually is not dissolved out; (3) by the paraffin method may
be obtained small sections (5 cm. square or less), and thin, by the
celloidin, larger sections, but thicker. With paraffin heat (melting
and cooling) is necessary, and the mass is sometimes spoken of as a
fusion imbedding mass; celloidin is a solution, and the mass is left
in the tissues by evaporation, or its equivalent.

In the celloidin method the imbedding mass with which the
spaces of the tissue are to be filled is collodion, a solution of celloidin
or proxylin* (soluble cotton) in ether and alcohol, hence the steps,
which are comparable with those of the paraffin method (see § 47),
are (1) Dehydration, removal of the water; (2) Saturation with
either-alcohol, the solvent of the celloidin; (3) Infiltration with cel-
loidin solutions, a thin and a thick; (4) Imbedding in a thick cel-
loidin mass, which is hardened and (5) sections cut.

§ 62. Dehydration. Let it be complete, as in the prepara-
tion for paraffin imbedding (§ 51). Immerse the tissue in 95%
alcohol for 12 to 24 hours or longer, changing 1 to 3 times. Consult
also § 51 upon the dehydration of tissue.

§ 63. Saturation with ether-alcohol (equal parts of pure ether
and absolute alcohol). Remove the tissue from the strong alcohol
and place it in a stoppered vial of ether-alcohol for 12 to 24 hours. In
addition to preparing the tissue for the collodion solutions, it com-
pletes the dehydration, should it be imperfect. In special cases, or
if the dehydration is very thorough and the specimen small, this step
may be omitted. A satisfactory infiltration is, however, more cer-
tain if ether-alcohol be used.

§ 64. Infiltration, (a) with thin celloidin. Pour off the
ether-alcohol and add the thin (2%) solution of celloidin in ether-
alcohol. This, being a solution in ether-alcohol with which the tissue
is saturated, readily permeates it. It is best to allow at least a day
for this to take place, although if there is time a stay of several days
is better, there being little or no danger of deterioration while in the
solution. With large (1 c. c. +) objects an infiltration of a week or
even a month is advisable.

Infiltration, (b) with thick celloidin. Pour off the thin col-
lodion solution and add thick (5 or 6%) solution (in ether-alcohol).
In this there is gradual concentration of the solution in the tissue.
Allow small specimens to remain a day, or, better, several days;

*Celloidin is a specially prepared and purified form of pyroxylin. It is-
about twice as expensive as pyroxylin or soluble cotton but with it better solutions
can be prepared. The pyroxylin on the market seldom affords stronger solutions
than 8%; with celloidin a 12% solution is easily prepared. The trimmings from
the celloidin blocks (after the alcohol or chloroform hardening) may be dried out
and redissolved and thus used over and over again. Pyroxylin may be used
equally well.

larger objects should be given a proportionately longer time, a week
to a month, or even longer.

If the object to be imbedded, such as many embryological speci-
mens, is one with large interior cavities with thin walls the transfer
from the thin solution to the thick solution may be attended by a
collapse of the walls and a consequent shriveling and distortion of
the specimen. Avoid this by allowing the thin solution to thicken
very gradually by evaporation in a dry atmosphere, as under a bell-
jar with calcium chlorid present until the solution has attained the
right consistency. To accomplish this it is only necessary to have
the cork of the vial containing the specimen perforated by a small
hole. A small piece of paper may be inserted with the cork, or with
porous corks no special effort need be made. Unless the thick solu-
tion has itself thickened by evaporation, with large specimens it is
advisable to follow the 6% bath with a stay in a thicker solution, as
10 or 12%, for a day or so.

§ 65. Imbedding. Pour off the 6% or 12% solution and add for
a short time at least a 12% solution of celloidin (in ether-alcohol).
The tissue is now ready for imbedding in 12%, which may be ac-
complished in either of two ways : (a) on a holder or (b) in a paper
box. Only those specimens need be imbedded in a box that, from
their shape, or for purposes of careful orientation or serial sectioning,
require a larger imbedding mass around them.

(a) On a holder (wooden-block). Choose a block of a conveni-
ent size; put a drop or two of celloidin upon one end and insert a
pin vertically to the surface near the edge. Transfer the tissue from
the vial of thick celloidin to the block and lean it against the pin.
The shape of many tissues will obviate the need of a pin. Pour the
thick celloidin onto the tissue, drop by drop, moving the block in
such a way that the thick viscid mass may be made to surround and
envelop the tissue. Continue to add drops of celloidin at intervals
until the tissue is well surrounded, and then as soon as a slight film
hardens on the surface invert the holder bearing the tissue in a
shell-vial of large diameter or glass box containing enough chloroform
to cover the specimen. Cork or cover so that the chloroform will
not evaporate. If the piece of tissue is of awkward size and shape,
oiled paper may be wound around the end of the wooden holder and
tightly tied, the projecting hollow cylinder being long enough to
receive the object. The tissue may be put into the cylinder as before,
the celloidin slowly poured in drop by drop until the specimen is

completely covered. When a film has formed, place in chloroform
as before.

(b) In a paper box. When a box is required for imbedding
proceed as follows: The inside of the paper box should be slightly
oily to prevent the celloidin from sticking to it. Rub upon the
paper that is to be folded to form the box a little vaseline, and then
with a cloth or lens paper remove as much as possible. Fold the
paper into a box of convenient size and shape. Remove the object
from the thick celloidin and place it in the box, arranging it in the
manner wished with a view to sectioning it later. Pour over it
slowly, drop by drop or a little at a time, a 12% solution of celloidin
until the specimen is well covered and the box sufficiently filled.
It is better to have a deep layer over the specimen. The 12% solu-
tion does not afford the best mass for cutting, so that, with large
objects, it is better to allow the mass in the box to thicken by evap-
orating it slowly under a bell-jar (aquarium jar) until it has attained
such a consistency that it is no longer fluid.

§ 66. Hardening. When the celloidin mass is thick enough
so that it only dents when touched with the finger nail it is ready for
hardening. This may be done by pouring chloroform into the jar
in which the imbedded material is placed, covering from the air.
The chloroform vapor hardens the mass. W7hen it is well set it may
be transferred to a jar of the chloroform wrhich takes out the ether-
alcohol and hardens the celloidin mass, for which a few^ hours is
sufficient. Allow the chloroform to act for 6 to 24 hours. The
imbedding mass remains quite transparent when no water is present.
The hardening action of the chloroform may be quickened and
intensified by carefully heating the chloroform until bubbles of ether
begin to come from the specimen. Do not let the chloroform evapor-
ate.

§ 67. Alcohol hardening. When the celloidin mass is hard,
whether clear or not, it may either be transferred to alcohol of about
82% strength in which it is stored until cut, or it may be placed in
Clarifier (castor oil, 1 part; xylene, 3 or 4 parts). Alcohol of higher
percentage softens the mass; lower grades such as 67% usually in-
crease the hardness of the celloidin and in some cases are to be
recommended.

The choice between alcohol and clarifier involves no decision of importance
in technique. The method of clarification has the advantage that the orientation
of the specimen in the microtome preparatory to cutting can be more perfectly

done. If the tissue has been stained in toto (§ § 79, 142) the sections may be
mounted directly from the clarifier as soon as cut. Any mercuric chlorid precipi-
tate (§11) that may be present can be dissolved out by means of a solution of iodin
in the castor-xylene. The castor oil, however, renders the microtome, knife, hands
of the operator, etc., sticky and the method is not so cleanly as the alcohol method.
On the other hand, alcohol tends to rust microtome and knife. Clarification is
preferred by the writer for serial work in celloidin.

§ 68. Clarification. Celloidin blocks transferred from the
chloroform hardener to an oil mixture such as castor-xylene (§ 67)
will become quite transparent (clarified) and hardly discernible, so
that the tissue is readily seen. Sometimes, however, the celloidin
remains white and opaque, due to the presence of moisture, and
considerable time is required for its clarification. In such cases the
process may be hastened by placing the tissue in the clarifier in a
warm place, and changing the clarifier several times. If the block
still remains opaque, remove to 95% alcohol for a day for dehydra-
tion, pass through chloroform, and into clarifier. In this way the
mass may usually be cleared perfectly. Change the clarifier to fresh
after the first day or so. The sectioning may be done after a few
hours' immersion, although a several days' clarification is preferable.*

§ 69. Cutting the Sections. There is no marked difference
in the sectioning of celloidin blocks preserved in alcohol and those
that have been clarified. In the following paragraphs 67% alcohol
should in the reading be substituted for clarifier if alcohol was used
in the hardening.

If a paper box was used, after the celloidin is ready for cutting,
remove the paper, trim the block as is desired (see below), put some
thick celloidin upon the wooden block or other holder and press the
base of the celloidin block firmly against it; within two minutes it
will be firmly cemented and one may proceed at once to clamp the
holder in the microtome and commence cutting.

For celloidin sectioning, a long drawing cut is necessary in order
to obtain thin, perfect sections. The knife should, therefore, be
set at an obliquity of 15 to 20° or less, so that half or more of the

*The imbedded object may remain in the castor-xylene clarifier indefinitely
without harm. The celloidin grows somewhat tougher by a prolonged stay in it.
After cutting all the sections desired at one time, the imbedded tissue is returned
to the clarifier for future sectioning. It should be remembered, however, that
pure castor oil is a solvent of celloidin, hence it is necessary to have the container
tightly stoppered, otherwise the volatile xylene will evaporate leaving the castor
oil behind.

blade is used in cutting the section. Recall that in the paraffin
method the knife is usually to be set at right angles to the direction of
the cut, and the stroke is a rapid straight one. Trim away the sur-
rounding celloidin mass leaving enough, however, to serve as a support
to the tissue and prevent its bending under the impact of the knife ; if
the celloidin mass is too tapering, bending will occur and thin sections
cannot be cut. To avoid this the celloidin block is best trimmed in
the form of a four sided truncated pyramid with as broad a base as
possible.

Clamp in the jaws of the microtome, placing it so that the mass
of celloidin is opposite the side to which the pressure of the knife is
applied in cutting. It is advantageous also to have the object placed
with its long diameter parallel with the edge of the knife.

When knife and tissue are properly arranged wet the tissue
well with clarifier or alcohol, — as the case may be, — and flood the
knife with the same. Make the sections with a slow, steady, motion
of the knife. With a small object (3x5 mm.) and a good sharp
knife, sections 5^ to 6^ can be cut without difficulty. In addition to
a sharp knife, however, there are necessary well-infiltrated tissue
and a hard, firm mass. If serial sections are not desired, it may be
more expeditious to cut dry and with a rapid stroke.

§ 70. Transferring the sections. If the sections are quite
thick they may be transferred from the knife to a slide or a dish by
means of forceps or a brush; if they are thin, however, it is better
to handle them by means of an absorbent tissue paper, as follows:
Flood the sections well with clarifier and then by means of a pipette
remove the clarifier from the knife and place over the sections the
end of a piece of the tissue paper, pressing it down upon the sec-
tions if necessa^. Carefully pull the paper off the edge of the
knife; the sections will adhere to the paper. Place the paper, sec-
tions down, on a slide, taking care that the sections are in the de-
sired position. With the finger carefully press the sections (through
the transfer paper) to the slide, and then lift the paper, with a roll-
ing motion, from the slide; the sections will adhere to the slide.
Should they stick to the paper instead, lower the paper again and
again firmly press the section to the slide. For further procedure
see § § 135, 136. If it is not desired to mount the sections upon a
slide immediately, or if they are to be kept in bulk, as for class
work, the transfer paper may be shaken gently in a dish of clarifier

or 05% alcohol and the section (or sections) will float free and sink
to the bottom.

§ 71. Serial Sectioning. If it is desired to mount the sections
in series, proceed as follows: With a. camel's hair brush or needle
draw the first section, when cut, up toward the back of the knife and
make the next section. Place this section to the right of the first, and
so on, arranging them in serial order, section after section, and line
below line, until enough are cut to fill the area that the cover-glass will
cover. Flood the sections as before by letting the clarifier flow over
them, being careful, however, not to float them from their places.
Absorb the clarifier from the knife with a pipette, and place over
the sections a piece of the transfer paper twice the width of a slide;
press it down if necessary, and slowly draw it off the edge of the
knife. Should it then be seen that some of the sections are adhering
to the knife instead of the paper, it means that the clarifier had been
allowed to thicken* on them, cementing them to the knife, and the
preliminary flooding to insure their being free, was insufficient. In
that case it is best to flood the paper with clarifier, carefully lift it,
arrange the sections again, flood them with clarifier, place a clean
piece of transfer paper over them and try again. One soon becomes
accustomed to the behavior of the sections, and accidents are rare.
In cutting a series of many small sections, some time is consumed
and it is necessary to flood the sections on the knife frequently with
clarifier while cutting in order to prevent the clarifier thickening and
cementing them to the knife.

§ 72. Resume of the method. Success in the employment of
the celloidin or collodion method depends upon the thorough infil-
tration with the solutions, requiring days or even months, and the
employment of a thick imbedding mass giving when hardened a firm
unyielding support to the tissue. This may be gained by employing
a relatively long period of infiltration, and taking pains in imbed-
ding to have the imbedding mass well thickened.

Observing these two cautions, celloidin may be used in almost
all cases as an imbedding mass, except such as are affected by the
conditions of the methods already mentioned (§ § 47 and 61).

*If one is a long time cutting a series of sections, it sometimes occurs that
the xylene evaporates leaving the castor oil that is thick and viscid and also
a solvent of the collodion, so that the sections are not easily transferable but stick
rather firmly to the knife. In such a case, fresh clarifier or even a little xylene
to dissolve the castor oil must be used.

THE FREEZING METHOD.

§ 73. This method is expeditious and of use in the rapid ex-
amination of tissues, and therefore especially serviceable in the
pathological laboratory and in clinical diagnoses. It may also be
used in cutting tissues that are too hard to be cut satisfactorily by
means of either the collodion or paraffin methods, and in the exami-
nation of tissues for substances (e. g., fats) which the solutions
necessary for the paraffin and celloidin methods dissolve out. Both
fresh and fixed tissue may be cut by means of the freezing micro-
tome and with or without any special mass such as is used in paraffin
or celloidin imbedding. Some histologists quite prefer the freezing
method to the paraffin or celloidin methods for general use.

When no mass is employed the tissue is simply frozen and cut,
or, if it is fixed tissue, soaked well in water first and then frozen.
When extreme haste is not so essential it is better to first saturate
the tissue with some solution that does not crystallize on freezing,
but simply hardens, since the formation of the ice crystals is hurtful
to the tissue. Such are solutions of gum arable or sugar and anise-
seed oil, and they are spoken of as Congelation masses.

§ 74. Infiltration. Gum arabic or anise-seed oil may be used.
(a) Gum arabic. If the tissue has been fixed and is in alcohol re-
move the alcohol by soaking it for several hours to 1 day in water.
Remove to a thick solution of gum arabic in water, in which it may
remain for about 24 hours. It is then ready ta freeze and cut.

(b) Anise-seed oil. For this method the tissue should be first
dehydrated (§ 51).* W^hen dehydration is complete, transfer the
tissue to anise-seed oil, in which it may soak for 12 to 24 hours; it is
then ready to freeze and cut. It is particularly adapted for use with
tissue that has been stained in toto.

§ 75. Cutting. Place a drop of the solution of gum arabic
(or anise-seed oil) upon the object carrier of the freezing microtome
and turn on the carbonic acid (or ether) spray. When the mixture
begins to harden, place the object upon it in an abundance of the
solution and freeze it nearly solid. Covering with an inverted cup
hastens the freezing. An especially wedge-shaped knife is necessary
because of the hardness of the mass.

*Anise-seed oil will, however, clear from 90% or even 82% alcohol; this
is sometimes of advantage.

When the tissue is completely frozen, cut it with a straight move-
ment of the knife, as in the paraffin method, holding it firmly upon the
knife rest and making the strokes as rapidly as possible, at the same
time rapidly raising the tissue a few microns at a time by means of
the microtome screw. There are a number of automatic microtomes
specially designed for use \vith the freezing method.

The mass of sections is transferred to a dish of water in which
the gum arabic is dissolved away and the sections are ready for stain-
ing (§ § 137, 146). If anise-seed oil is used, the sections are to be
transferred to 95% alcohol which will dissolve out the oil; if the
tissue has been stained in toto the sections may be transferred to
anise-seed oil (or other clearer) and mounted in balsam directly.

§ 76. Rapid Method. Blocks of tissue 1 centimeter thick should
fix in 10% formalin 12 to 24 hours. If haste is a factor, take thinner
pieces and fix for 1 minute or more. Trim the block so that it is about
5 mm. thick; rinse in water for a few seconds, transfer to the freezing
microtome, freeze and section.

Float the sections when cut from the knife into water from which
they may be gotten upon the slides by means of a camel's hair
brush. Drain off the water and press the sections out smooth by
means of blotting paper, filter paper or other absorbent paper. Cau-
tiously drop over the sections 95% and absolute alcohol and follow
this immediately with thin (% or /4%) celloidin solution (§ 64)
which when it has partially evaporated out will serve to support the
section and- fasten it to the slide. It is now ready for staining.
(§ 144).

STAINING.

§ 77. Staining has for its first and primary purpose, the ren-
dering outlines and structures more distinct by giving them a color
contrast with their surroundings (color image). A second and
more important use is for the differentiation of particular struc-
tures or substances which by their selective staining facilitate the
histological analysis. Rational staining, like rational fixation, de-
pends upon the physics and chemistry of staining reactions; indeed,
in the demonstration of particular substances the fixation and stain-
ing should be determined by the mutual interdependence of their re-
actions, since they have the same purpose, — the preservation and
demonstration of the substance sought for.

In some cases the differential staining may be accomplished in
the fixing (fats, impregnations). Differential staining, histo-chemical
methods, micro-chemical methods (as applied to the animal organism)
thus belong in the same category and rest upon a physico-chemical
basis. As in the case of fixation, a great deal remains to be done
in the perfection of this side of histological technique.

§ 78. Classification of Stains. Stains may be grouped: (a)
according to their chemical composition as (1) organic; — hematoxylin
stains, carmine stains, anilin stains, (coal-tar dyes; benzene deriva-
tives), and (2) inorganic, (b) From another chemical aspect as (1)
basic or (2) acid, depending upon the chemical reaction of the staining
principle or group, (3) neutral (§ 85). (c) Histologically, stains are:
(1) nuclear (chromatin stains), (2) plasma or general stains, (3) special
stains, (4) impregnations. The first are usually basic, the second
acid, the fourth inorganic.

§ 79. In toto staining. When in toto or bulk staining is em-
ployed, the piece of tissue is stained entire and imbedded and sec-
tioned afterwards. In this case the tissue should be stained before
the process of embedding has begun, after the washing out of the
fixer has been completed (§ § 7, 9). But a single stain may be given
and the one chosen is generally a nuclear one, — hematoxylin, cochi-
neal, or carmine (§ § 91, 98 — ). A counter stain may be given sub-
sequently after sectioning, orange G after hematoxylin or cochineal,
Lyon's blue after carmine being most satisfactory. In toto staining
is particularly useful in serial sections of embryological material and
in morphological work in general, as it saves time and manipulation.

§ 80. Section Staining. The application of the staining proc-
esses after the tissue is imbedded and sectioned. This is more
serviceable, especially if highly differential results are desired.

§ 81. Progressive and Regressive Staining. According to the
mode of application, staining is either progressive or regressive.
Progressive staining consists in permitting the staining to proceed
to the right degree of intensity and then stopping it. In regressive
staining the tissue is over-stained and the excess of stain removed
by the application of a Differentiator. Uusally in the regressive
method there is employed a Mordant which is a chemical solution
serving to make the stain "take." It possesses a double affinity, —
to the tissue and to the stain, which is usually made to operate in the
interest of differential or selective staining. The mordant is not

always in the form of a separate solution (e. g., aluminium hema-
toxylin) and in many cases the mordantage is given in the fixing.
Mordantage, directly or indirectly employed, is also useful in staining
by the progressive method. Stains or dyes that do not require a
mordant are termed Substantive, those requiring a mordant to make
them "take" are Adjective stains.

Delicate results in exact differential staining by either the progres-
sive or regressive methods can best be secured only by section stain-
ing, although differentiation of the sections after in toto staining may
be resorted to.

§ 82. Differentiation. In the regressive method it is neces-
sary to remove the excess of stain by the application of a solution
that will usually differentiate it, or bring out the selective action of
the stain. A small amount of differentiation is usually necessary
in any case for the most delicate results. Alcohol, 95% or other
grade, may often be used as a differentiator. 95% acidified with
hydrochloric acid (1/10 to 1%) is sometimes used, or^a special dif-
ferentiator is required (acetone, alum solution, clove-oil, etc.).
It is usually necessary to control the differentiation by use of the
microscope.

§ 83. Impregnations. In addition to the typical methods of
coloring tissue by means of stains there is a group of methods in
which the coloring matter is deposited in the cell or tissue that it is
desired to differentiate, in the form of a precipitate. These are
known as impregnation methods and are of great value, especially
as applied to nervous tissues. A hard and fast line, however, can-
not be drawn between true staining methods and impregnations.
Silver nitrate and gold chlorid are the substances most generally
employed in the impregnation of tissue (§ § 221, 223).

§ 84. Choice of stains. Remember that the staining is greatly
affected by the previous treatment, brilliancy or selectivity in the
result being in many cases dependent on the fixer employed or im-
paired by the improper or incomplete washing out of the fixer.

In staining, therefore, consider three things, (a) what it is de-
sired to bring out, — what kinds of stains you need to employ, (b)
the mode of fixation that has been emploj^ed, and (c) the imbedding
method must also be considered in the choice, since if celloidin is
employed certain stains that color it deeply should be avoided.

(a) For general purposes it is customary to use a nuclear and
a cytoplasmic stain either in combination or successively. The
double staining most employed is hematoxylin and eosin. Triple
stains are generally not so satisfactory; hematoxylin and picro-
fuchsin is one commonly used. Neutral stains (§ 85) are essentially
triple stains. Quadruple stains are rarely serviceable; see, however,
§ 129. For stains to be employed for special purposes, see Special
Methods, § 174-.

(b) In the case of most of the fixers given § 11-, there are no
restrictions as to the stains that may be employed though some
are more satisfactory than others. With Flemming's fluid and
Hermann's fluid, however, it is well to employ only such stains
as iron hematoxylin (§94) and anilin stains, — safranin being suggested
as a red stain, gentian violet as a blue stain. Dichromate fixers whose
action has been prolonged usually require strong stains. Iron hema-
toxylin and similar stains may be used after any fixer. The Ehrlich-
Biondi-Haidenhain stain (§ 111) is most satisfactory after Mercuric
chlorid (§ 11).

(c) Celloidin is deeply stained by such stains as iron hema-
toxylin, safranin, methylene blue, gentian violet, and basic stains in
general. If it is necessary to use these, with celloidin material, the
celloidin should be removed from the sections before mounting
(§ 136).

§ 85. Neutral Stains. In the case of most of the anilin stains, if an aqueous
solution of a basic stain is added to an aqueous solution of an acid stain, there is
formed by combination a neutral stain which is usually but slightly soluble in
water and hence precipitates out. Neutral stains are however moderately soluble
in strong methyl or ethyl alcohol. These facts it is well to keep in mind in using
anilin stains. The reaction may be made use of for increasing the selectivity,
sharpness and color tones of the original stains. Neutral stains may be used in
one of three ways: (a) The neutral stain is often soluble in excess of the acid
or basic stain or in an aqueous solution of another acid or basic stain (§111). (b)
Dissolved in methyl (or ethyl) alcohol and diluted nearly to the precipitation point
the activity and selectivity of the component stains seem to become specially
marked (§ 128, 214). (c) The formation of the neutral stain may be carried out
in the staining process itself, — "on the slide," — by staining first strongly with an
aqueous solution of the acid stain, rinsing away the excess stain and staining with
the aqueous solution of the basic stain (§ 106) and differentiating (§ 82) with
alcohol, or alcohol and clove oil.

Of the large number of stains that combine to forrn neutral stains may be
mentioned, — the acid stain being given first; — eosin and methylene blue (§ 128);
orange G. and gentian violet (§ 131), erythrosin and toluidin blue, thiazin red and
toluidin blue, coerulein S. and safranin, acid violet and safranin. These may be
combined and used in any one of the three ways mentioned above.

§ 86. The time of staining. Although in general certain time
limits can be given to the period during which a stain should be
allowed to act, with most stains, especially those with which no dif-
ferentiation is needed, such as hematoxylin, and most carmines, the
correct intensity of color should be determined by examining the
preparation with the microscope. One soon becomes able to judge
of the right stain in this way better than if a given time were adhered
to.

In the use of stains requiring a subsequent differentiation, the
rule is 'to over-stain and watch the differentiation carefully with the
microscope, stopping it when sufficient. In this case it is the differ-
entiation and not the staining that should be carefully regulated.
In general, for the best results, it is advisable to use staining and
differentiating solutions in dilute form and prolong the time during
which they act.

The following formulas include the more generally useful stains
and those to be employed in the "Special Methods" given subse-
quently.

STAINS.

§ 87. Hematoxylin Stains. Hematoxylin is a colorless compound of acid
properties forming therefore salts with bases which oxidize readily forming
"hemateates." The oxidation product of hematoxylin is hematein which is
the real staining principle and may in some cases be used with real advantage
instead of hematoxylin. Hematoxylin (hematein) itself has little value as an
acid (plasma) stain; combined with metallic bases it becomes a valuable basic
(chromatin) stain. The metals usually employed as mordants for hematoxylin
are: aluminium, iron, copper, chromium, molybdenum, vanadium. Their
salts may be used either in the same solution as the hematoxylin (aluminium,
molybdenum, vanadium) or separately (iron, copper.) Solutions of the same
metals may in some instances (aluminium, iron, chromium,) be employed also as
differentiator. For preparing hematoxylin stains it is a great convenience to
have a 10% stock solution in 95% alcohol.

§ 88. Chloral Hematoxylin. [11] Formula: Potassium alum, 8 grams;
distilled water, 250 c. c.; hematoxylin 2/10th gram or 2 c. c. hematoxylin stock
solution. Boil 5 or 10 minutes in an agate dish. After cooling add 6 grams of
chloral hydrate. Place in a bottle and permit the hematoxylin to oxidize for a
week or two, or 1 to 2 c. c. hydrogen peroxid may be added. Its staining quality
improves up to an optimum and then begins to deteriorate. Old hematoxylin
generally contains a precipitate and should be filtered often or before using.

Stain sections 5 to 30 minutes according to the age of the solution, the charac-
ter of the tissue and the fixation employed. After staining wash well with dis-
tilled or tap water. Usually no differentiation is required unless a purer chroma-
tin stain is desired (§ 175). Counter stain as desired.

§ 89. Mayer's Haemalum. Formula: haematein, 1 gram; 90% (95%)
alcohol, 50 c. c.; potash alum, 50 grams; distilled water, 1,000 c. c. Dissolve
the haematein in the alcohol, the alum in the water; mix.

This is an excellent formula, giving a good stain immediately after it is made
up and retaining its selective staining quality for a year or longer. It is one of the
best hematoxylins for ordinary work.

For this formula, Mayer has now substituted the following: hematoxylin,
1 gram (10 cc. stock solution); distilled water, 1,000 c. c.; sodium iodate, 0.2
grams; potassium alum, 50 grams. The hematoxylin is first dissolved in the
water and then the other ingredients are added. Dissolve and filter. 50 grams
of chloral hydrate and 1 gram of citric or acetic acid may be added as a preserva-
tive. Stain as given in § 88. In using it is frequently advisable to dilute one or
more times with distilled \vater.

§ 90. Ehrlich's acid hematoxylin. Formula: Water, 100 c. c.; 95%
alcohol, 100 c. c. hematoxylin crystals, 2 grams (20 c. c. stock solution) ; dissolve in
the alcohol; glycerin, 100 c. c.; glacial acetic acid, 10 c. c.; alum in excess. Let
the mixture ripen in the light until a dark red. Sections stain in this hematoxylin
in a short time, generally 5 to 10 minutes. Wash with water after staining.

§ 91. Delafield's hematoxylin. Formula: Saturated aqueous solution
of ammonia alum, 200 c. c.; hematoxylin .stock solution 20 c. c. Allow the
mixture to stand in the light and air in an unstoppered bottle for 4 or 5 days;
filter and add glycerin, 50 c. c., and methyl alcohol, 50 c. c. Permit it to stand
for a week or so to ripen; filter and keep in a stoppered bottle. The staining
power increases for several months. In using, dilute 3 or 4 times or more with
distilled water. It is useful for in toto staining diluted 1 :9 with 20% alcohol or
distilled water.

Stain sections from water; 4 to 5 minutes will generally be sufficient. Wash
well with water after staining. In toto staining 1 to 14 days may be necessary
depending upon the size of the object.

This is a very strong hematoxylin stain and may be used to advantage with
tissues that stain with difficulty. It is likewise a more diffuse stain than either
chloral or Ehrlich's hematoxylin, staining cell-body as well as nucleus, — a feature
having its advantages. Old solutions (several months to a year) should be
filtered before using.

§ 92. Acid Hematoxylin (Delafield's). The dilution of Delafield's hema-
toxylin ten to twenty times with water, and making slightly acid with acetic
gives a useful stain particularly for differentiating nuclei (embryological work.)

§ 93. Muchematein (Mayer). Aqeuous formula: Rub up 0.2 grm. hema-
tein with a few drops of glycerin; add 0.1 grm. aluminium chlorid, 40 c. c. glycerin,
60 c. c. distilled water. Filter if necessary. Alcoholic formula: Hematein, 0.2
grms.; aluminium chlorid, 0.1 grm.; 70% (67%) alcohol, 100 c. c.; 1 or 2 drops
nitric acid. Formulae with a minimum amount of aluminium designed as specific
stain for mucus. The aqueous solution is more selective; the alcoholic formula
designed for the staining of mucus that swells excessively in water (§ 234).

§ 94. Iron Hematoxylin (Heidenhain). Formula: (a) Mordant- 2%
aqueous solution of ferric alum (iron-ammonium-persulphate), (b) Stain;
aqueous solution of hematoxylin (10% alcoholic stock solution, 5 c. c.;

distilled water, 95 c. c.). (c) Differentiator; the ferric alum mordant, preferably
diluted several times. An excellent stain, especially for cytological work. It
may be used after any fixer.

The steps necessary are: (1) Mordanting 1 to 24 hours; (2) rinse the, sec-
tions in water 10 to 30 minutes; (3) stain for 3 to 24 hours; differentiate slowly
and control it under the microscope. The slides may be alternately dipped into
the ferric alum solution for a few seconds and then into tap water. (4) Wash
in running water 15 to 60 minutes. The ferric alum mordant may be used several
times as well as the hematoxylin solution whose staining quality improves up to
a limit by use.

§ 95. Copper Hematoxylin (Weigert). Formula: Mordant; 3.5% aqueous
solution of copper acetate; Stain; 10% alcoholic solution of hematoxylin 10 c. c.;
distilled water, 90 c. c.; saturated solution lithium carbonate, 1 c. c. Differentia-
tor; potassium ferricyanide, 2.5 grms., borax, 2 grms.; distilled water, 200 c. c.
Designed for staining the myelinic sheath of medullated nerve fibers (§ 199) but
useful for other purposes.

The steps are: (1) mordanting for 1 to 24 hours, (2) rinsing the sections
in water 10 to 30 minutes, (3) staining for 3 to 24 hours, (4) differentiate slowly
controlling the action under the microscope. (5) Wash in running water 30
minutes or longer. The mordantage may be given in bulk if desired. As usually
employed with tissue fixed in chrome fixers, the stain is strictly a chrome-copper
hematoxylin. Other differentiators may be used if desired. Mount in neutral
balsam.

§ 96. For Mallory's phospho-molybdic hematoxylin, Heidenhain's vanadium
hematoxylin, and other hematoxylin formulae of occasional usefulness, consult
the works on technique (p. 90).

§ 97. Carmine Stains'. Like hematoxylin, the carmine stains depend
upon an acid staining principle, — carminic acid, — which in combination with
bases gives a red nuclear stain of value. The metals usually employed in carmine
formulae are aluminium, calcium, strontium, iron. Carmine itself is (Liebermann)
a combination of carminic acid with aluminium, calcium, and protein. It is
soluble in acids and alkalis. The necessary mordant is not so often employed
in a separate solution as is the case with hematoxylin. See, however, iron carmine
[6, 30].

§ 98. Carmalum (Mayer's) Formula: Carminic acid, 1 grm.; potas-
sium alum, 10 grms.; distilled water, 200 c. c. Dissolve with heat (if neces-
sary). Filter. Add 1 c. c. formalin as a preservative. It may be used for
in toto or for section staining.

Stain sections 5 to 30 minutes or as long as necessary. Rinse tissue before
staining with distilled (not tap-) water.

§99. Borax carmine. (Grenacher). Formula: Borax 4 grams;' carmine, 3
grams; water, 100 c. c.; allow the mixture to stand for several days, shaking
occasionally when most of the carmine will have dissolved; filter and add 100
c. c. of 70% alcohol. Let the mixture remain for several days, filter again and
the solution is ready for use.

This is a good carmine stain for in toto staining. Stain objects in toto for one
to several days, according to size; remove to 67% (70%) alcohol, acidulated

slightly with hydrochloric acid, (4 drops in each 100 c. c.), for a day and then
remove to 80% alcohol. It affords a bright red stain that is quite transparent.

§ 100. Paracarmine (Mayer). Formula: Carminic acid, 1 gram; alumi-
nium chlorid, 0.5 gram; calcium chlorid, 4 grams; 70% (67%) alcohol, 100 c. c.
Allow it to stand a day or so, shaking occasionally until the carminic acid has
quite dissolved, and then filter.

This is an excellent carmine stain for in toto staining. The tissue may be
stained one to several days (1 week), then washed in 67% and 82% alcohols to
remove the excess of staining fluid. A red nuclear stain, more opaque than borax
carmine. It does not over-stain readily, and since it is an alcoholic solution (70%)
it is quite penetrating and may be allowed to act for a greater length of time,
being thus suited for staining in toto objects of considerable size.

§ 101. Hcl. carmine. Formula: Carmine, 2 grams; concentrated hydro-
chloric acid, 3 c. c.; 70% alcohol, 100 c. c. Boil gently for 15 to 20 minutes to
dissolve the carmine; cool and filter.

This is a strong carmine stain, quite suitable for sections, especially such as
stain with difficulty. It may also be employed for staining in toto. Stain sec-
tions from alcohol or water for 5 to 15 minutes ; rinse away the superfluous stain
with 67% (70%) alcohol and differentiate for a few seconds to a minute with acid
alcohol (95% alcohol 100 c. c., concentrated hydrochloric acid 1/10 c. c.). Wash
away the acid alcohol with ordinary 95% alcohol. If a pure nuclear stain is not
desired the differentiation may be omitted.

Picric acid (§ 116) may be used as a counter stain, and in that case differentia-
tion is ordinarily not required.

§ 102. Alum Cochineal. Formula: Powdered cochineal, 75 grams; potas-
sium alum, 75 grams; distilled water, 1000 c. c. Boil the ingredients for half an
hour, or (better) macerate for a day or so, boiling up two or three times. Cool
and filter. Add to the filtrate distilled water to make up 1000 c. c. and a crystal
of thymol as a preservative.

This is an excellent stain, particularly for in toto staining of embryos, giving a
purpler stain than the carmine stains given above.

Stain objects, such as embryos, over night to 2 days or longer, depending on
size. Wash out with water, 2 to 6 hours and place successively in 50%, 67% and
82% alcohols several hours to a day in each.

§ 103. Mucicarmine. Formula: Carmine, 1. gram; alummium chlorid
(pure), 0.5 grams; distilled water, 2 c. c. Mix thoroughly together and heat over
a small flame for 2 minutes (in a test tube) until the mixture has become dark red.
Dissolve the whole in 100 c. c. 50% alcohol added gradually; after 24 hours
filter. For use, it may be diluted 5 or 10 times with water.

Stain sections 10 minutes or longer in the diluted stain, rinse in water, dehy-
drate, clear, and mount in neutral balsam.

If desired, the stain may be diluted with 50 or 67% alcohol instead of water.

§ 104. Gentian Violet. Formula: A concentrated solution in distilled
water. Stain (paraffin) sections from water for 5 to 10 minutes, rinse in water,
dehydrate and differentiate with 95% alcohol and complete the differentiation
with clove oil. When the differentiation is sufficient, clear with bergamot oil

and mount in balsam. This may be used alone to give a blue stain with tissue
fixed in Hermann's or Flemming's fluid. (See also § 112, 131).

§ 105. Methylene Blue. This valuable stain, used particularly in the
histology and pathology of the blood and nervous system, and in bacteriology,
is represented in a large number of formulae. For the staining of nuclei, basic
granules in the cytoplasm, neurochromatin granules, etc., simple aqueous solu-
tions may be employed. A 1% solution suffices for most purposes; in some
cases, a concentrated solution is to be preferred.

Stain from water for 5 minutes to as many hours, with or without heat, rinse
with distilled water, differentiate if desired in a l/10th% Hcl. in 95% alcohol,
or l/10th% alum solution. Wash, dehydrate, mount in neutral balsam.

§ 106. Alkaline Methylene Blue. Formula: Methylene blue, 2 grms.;
absolute or 95% alcohol neutralized with pure dry sodium carbonate, 50 c. c.;
add distilled water, 450 c. c.; 1% potassium hydroxid, 5 c. c. An excellent stain,
giving best results after mercuric chlorid fixers (incl. Zenker's, etc.).

§ 107. Eosin— Methylene Blue. Stain sections l/2 hour with a l/4% to 1%
aqueous solution of eosin, rinse in water, stain in alkaline methylene blue 10
minutes, rinse well in water. Differentiate and dehydrate rapidly with neutral
95% alcohol and absolute alcohol, clear in xylene, mount in neutral balsam.
Particularly useful for staining blood in the tissues (hemolymph) glands, etc..

§ 108. Toluidin Blue. This may be used, often to advantage, in place of
and for the same purposes as methylene blue. It gives a somewhat darker stain.

§ 109. Methyl green. This is a nuclear stain of much value, besides being
an important ingredient of triple stains (e. g., Ehrlich's triacid mixture and § 111).
In very dilute solutions it is serviceable in staining the nuclei of fresh tissue and
of isolated cells. A 1 % aqueous solution may be used with hematoxylin and picro-
fuchsin in differentiating the structure of the hair follicle. (Gage).

§ 110. Safranin. Formula (Babe's): Concentrated aqueous solution of
safranin, 1 part; concentrated alcoholic solution of safranin, 1 part.

Stain sections 1 to 4 hours, or over night; wash away excess of stain with
95% alcohol, differentiate with acid alcohol (95% alcohol, 100 c. c., hydrochloric
acid, 1/10 c. c.) for a few seconds, rinse with 95% alcohol and clear in carbol-
xylene or bergamot oil. If a pure nuclear stain is not required, the differentiation
may be omitted. This gives a good stain with tissue fixed in Hermann's or
Flemming's fluid. It is a brilliant, transparent red.

Other formulas may be employed (concentrated alcoholic solution, alcoholic
solution diluted with anilin water, equal parts concentrated solutions in alcohol
and anilin water, etc.). Differentiation may be accomplished with the use of
iodin-potassium iodid solution, or by counterstaining with an alcoholic solution
of light green or acid violet (§ § 121, 122).

§ 111. Ehrich-Biondi-Heidenhain Mixture. Formula: Saturated aqueous
solutions of Orange G. Rubin S. (Fuchsin acid) and Methyl green, 100 c. c., 20 c. c.
and 50 c. c. respectively. In preparing the mixture, only fully saturated solutions
should be taken which should be mixed in the order given slowly with constant
agitation. Only tissue that has been fixed in sublimate solutions (§11) should
be used, the sections should be thin and slightly acid. This may be secured by

treating with l/10th per cent, acetic acid before staining. The stain is useful for
some cytological work (nuclear degenerations, cytoplasmic transformations, etc.).
In staining use the stock solution diluted with distilled water 1 :60 and rendered
slightly acid by the addition of 0.2% acetic acid, drop by drop, until the red color
tone, due to the fuchsin acid, becomes slightly accentuated. The success of the
stain depends upon having the staining reaction right; if too acid, the fuchsin
acid predominates, otherwise the green and orange prevail. Stain 6 hours or
more; when sufficient, dehydrate rapidly with absolute alcohol, clear with xylene,
mount in xylene balsam.

§ 112. Flemming's Triple Stain. Stain sections in an alcoholic solution of
safranin diluted with an equal volume of anilin water, for a day or longer. Differ-
entiate in absolute (or 95%) alcohol with l/10th % Hcl until hardly any more
color comes away. Stain for 1 to 3 hours in a 1% aqueous solution of gentian
violet. Rinse in distilled water and treat with a strong (2%) solution of Orange G.
in distilled water, and while clouds of violet are still being given off, bring the
sections into absolute alcohol in which the differentiation is begun. Transfer the
sections to clove oil or bergamot oil which completes the differentiation and clears.
Mount in balsam before the last pale clouds of color have ceased to come away.

This stain is only recommended after such fixers as Flemming's fluid or Her-
mann's fluid. The stain is somewhat fickle, giving the best results only after some
practice. It is useful in some cytological work.

Several modifications have been proposed. A short method is frequently
employed as follows: (1) stain for a second or two in a mixture of equal parts of
saturated aqueous and saturated alcoholic solutions of safranin; (2) rinse in
water; (3) stain 2 to 10 minutes in 1% gentian violet; (4) rinse in water; .and (5)
stain for 10 seconds or longer in a 2% solution of orange G.. Dehydrate rapidly
with absolute alcohol; clear and differentiate with clove oil, controlling the
differentiation under the microscope. Remove the clove oil with xylene or toluene
and mount in balsam.

§ 113. Congo Red. A well known indicator, red in neutral or alkaline
solutions, turning blue in the presence of free mineral and many organic acids,
not affected by acetic, lactic or carbonic acid in the presence of ammonia. It is
useful as a plasma stain after hematoxylin, gentian violet, etc.

It may be employed in aqueous (^2 %) or alcoholic (2%) solution. With
subsequent differentiation in acid alcohol (§ 110) it is a useful stain with gastric
glands. Occasionally useful as an indicator with living organisms or tissue.

§ 114. Eosin. Formulas: (a) %% aqueous solution; (b) 2-% aqueous
solution; (c) 1/10% solution in water or 95% alcohol. Formula (a) is preferable
for most work; (b) affords a stronger and (c) a weaker stain. This may be used
as a counter-stain with hematoxylin to differentiate nucleus from cell-body.
Stain sections after hematoxylin for 10 to 30 seconds, wash away the excess of
stain with distilled water or 67% alcohol. Since alcohol tends to wash out the
eosin, unless the color is too strong it is advisable to hasten the process of washing
out and dehydration.

§ 115. Erythrosin. Formulas: (a) l/z to 1% solution in 67% alcohol,
(b) ^4 to 1% aqueous solution. This is a general stain similar to eosin in its

staining properties, but gives a redder color. Formulas (a) and (b) may be used
with sections and in the same way as eosin.

§ 116. Picric Acid. A counter-stain useful after carmine or hematoxylin.
Use ay£tol% solution in 67% or 95% alcohol, or simply add it to the alcohol used
in dehydration. It washes out the hematoxylin and is useful as a differentiator
of this stain, with which it is well to overstain somewhat if it is desired to counter-
stain with picric acid.

§ 117. Orange G. An excellent acid stain that may be employed as counter-
stain and differentiator after hematoxylin, gentian violet, etc.

Employ (a) 2 to 4% aqueous solution which may with advantage be slightly
acid (Hcl.) ; in the last event, sections should be rinsed well before the mounting.
It may with good effect be combined with eosin, erythrosin or fuchsin acid, (b)
Frequently a concentrated solution in 95% alcohol is to be preferred to the
aqueous solution.

§ 118. Fuchsin Acid (Rubin S., Magenta S.). Like Congo red, an indi-
cator, red in acid or neutral solutions, bleached by alkali. It is a valuable
plasma stain, but requires care in its use because of its sensitiveness to alkali.
Tap water should therefore be avoided. The staining solution and the mounting
medium should preferably be slightly acid, — at least not alkaline.

Employ a 2 to 4% solution; stain 1 minute to 24 hours.

§ 119. Picro-fuchsin. Formulas: (a.) General stain, — 1% aqueous solution
of fuchsin acid, 10 c. c.; saturated aqueous solution of picric acid, 75 c. c.; dis-
tilled water, 25 c. c. (b) For nervous tissue, — 1% aqueous solution of fuchsin
acid, 15 c. c. ; saturated aqueous solution of picric acid, 50 c. c. ; distilled water,
50 c. c. This is a valuable counter-stain to hematoxylin, especially serviceable
in the differentiation of white connective tissue fibers. The nuclei are a purplish
brown (hematoxylin stain), the connective tissue red, cell bodies and muscle
yellow-orange. In special cases the relative amount of fuchsin acid may be
decreased or increased, thus giving a preponderance to the yellow or red in the
general stain.

Stain well with hematoxylin, rinse in water, and stain with the picrofuchsin
15 to 30 seconds; wash away the excess of stain with distilled water or 67%
alcohol. Picro-fuchsin will gradually wash out the hematoxylin, therefore stain
strongly with hematoxylin and regulate carefully the time of staining with picro-
fuchsin. Picro-fuchsin is quite sensitive to alkalies, so that tap-water (unless
slightly acidulated) should not be used for washing out and the mounting medium
should be slightly acid or neutral, not alkaline.

§ 120. Mallory's Anilin Blue Connective Tissue Stain. Fix tissue in Zenker's
fluid (preferred) or in a mercuric chlorid fixer; imbed preferably in paraffin.

Sections are to be stained (a) for 5 minutes or longer in a 1/5% aqueous solu-
tion of acid fuchsin, and (b) without washing, stained about four times as long in
the following: Griibler's water soluble anilin blue, 0.5 gram; orange G., 2.0
grams; 1% aqueous solution of phosphomolybdic acid. 100 c. c. Wash, differ-
entiate and dehydrate with 95% and absolute alcohol. Clear in xylene and
mount in balsam.

A stain of quite general usefulness. The collaginous connective tissue fibers,,
reticular tissue, cartilage, osseous tissue, mucus, etc., are stained blue; nuclei,,
cytoplasm, muscle, red; red blood corpuscles, orange.

§ 121. Light Green. An acid stain sometimes useful as a counterstain with
safranin. A 0.2% alcoholic solution may be used in the differentiation of the
latter stain.

§ 122. Acid Violet may be used in similar cases and in the same manner^

§ 123. Bleu de Lyon (Lyons' Blue). A plasma stain frequently useful as a
counter stain after safranin or carmine (in toto). Employ an alcoholic (95%)
solution of about half saturation and stain sections for 10 to 15 minutes or longer.
Rinse with 95% alcohol and dehydrate more or less rapidly depending on the
differentiation desired. •

§ 124. Iodine. In addition to a certain value as a fixer and to facilitate the
removal of mercuric chlorid from tissue, usefulness as a differentiator after basic
aniline dyes, iodine has a place among the stains for the differential coloring of
starch, glycogen, amyloid, cellulose, etc. The following solutions may be men-
tioned: (a) saturated aqueous solution; (b) Iodine-potassium iodid solutions.
(Gram's, Lugols), — iodine 1 gram, potassium iodid 2 grams, distilled water 300 c. c
Lugol's solution is six times as strong, (c) 10% alcoholic solution, (tincture);
(d) for a formula useful for staining glycogen, amyloid, etc., see § 229.

§ 125. Resorcinfuchsin (Weigert). Prepare a concentrated solution of
the dry powder in a 1% solution of hydrochloric acid in 95% alcohol. Designed
for staining differentially elastic connective tissue fibers.

Stain sections 15 to 30 minutes, wash and dehydrate in 95% and absolute
alcohol, clear in xylene, mount in balsam.

The stain may be directly prepared according to a modified formula of Weigert,,
Formula: Fuchsin (basic), 2 grams; resorcin, 4 grams; water, 200 c. c. Boil
several minutes (10 or more); add 25 c. c. 30% solution ferric chlorid and boil
5 or 10 minutes longer. If the stain is not all precipitated, more of the ferric
chlorid solution may be added. Permit the liquid to cool. Let the precipitate
settle and decant fluid, or filter. Dissolve precipitate in 200 c. c. 95% alcohol,
employing heat if desired (boiling on a water bath). Filter. When the filtrate
is cool, add 4 c. c. strong hydrochloric acid.

The stain may be followed by picro-fuchsin or other similar reagent for stain-
ing the white connective tissue fibers.

§ 126. Orcein (Taenzer-Unna). Another standard method of staining
the elastic connective tissue fibers (elastin). Formula: Orcein D., 1 gram;:
95% alcohol, 100 c. c.; strong hydrochloric acid, 1 c. c.

Stain sections ]/2 to 1 hour, wash and differentiate in 95% alcohol and acid
alcohol (§ 82). Elastin stained dark brown. Other differential stains may be
used on the same preparation.

§127. Verhoeff's Elastin stain. [44] Formula: Hematoxylin, 0.15 grams;.
Absolute alcohol, 25 c. c.; dissolve by heat and add 1 drop of 5% ammonia solu-
tion. Permit it to stand for at least 5 minutes and add Lugol's solution (§ 124),.
22 c. c. ; filter and permit it to stand for 24 hours in a corked bottle. The solution
remains good for about 3 months. For use, add a 7% ale. sol. ferric chlorid, 1
drop per c. c.

Stain sections from alcohol, 12 to 24 hours; wash in water 1 to 5 minutes;
differentiate in a 1% ferric chlorid solution (ale. stock solution, 1 part, water 6
parts). Counter stain with picric acid (§116). A sharp stain, though possibly
not in all cases completely differential.

128. Jenner's Stain [27]. This is an eosinate of methylene blue. Formula:
Mix equal parts of 1.25% eosin (water soluble) and 1% methylene blue. Permit
the mixture to stand for 24 hours, then filter, wash the precipitate on the filter
with distilled water, dry it and dissolve it in methyl alcohol (0.5 grm. precipitate
in 100 c. c. alcohol).

This is primarily a stain for blood films. Sections may be stained in one of two
ways: (a) Stain 1 to 5 minutes, dilute the stain with an equal volume of water
and continue the staining for 5 to 10 minutes. Rinse with distilled water, differ-
entiate and dehydrate rapidly with 95% and absolute alcohol. Clear in xylene.
Mount in balsam.

(b) Dilute the stain with 1 to 4 volumes of distilled water and stain 1 to 24
hours. Further treatment as above. To insure permanency, it is well to thor-
oughly remove the alcohol with xylene in clearing, by 2 or 3 treatments.

§ 129. Wright's stain. Contains several staining principles, notably eosin-
ates of methylene blue, methylene azure, in solution in methyl alcohol, and is
made according to a complicated method, for which see original article [46].

Primarily for blood films (§ 214). Dissolve 0.2 grams of the dry stain in 100
c. c. of methyl alcohol. In staining sections follow the procedure given in §128.

§ 130. Nochts-Hastings Stain [22]. Similar in its composition to Wright's
stain and used in a similar manner. In staining sections, follow the procedure given
in § 128. Dissolve 0.3 grams of the dry stain in 100 c. c. of methyl alcohol. An
excellent blood stain (§ 214).

§ 131. Neutral Gentian Violet (Reinke, Bensley). Formula: Saturated
aqueous solution of gentian violet and saturated solution of orange G. are mixed
in equal proportions and permitted to stand 24 hours. The precipitate that forms
is removed by filtration, washed with distilled water, dried, and a concentrated
solution in methyl or ethyl alcohol is made. When ready to use it, dilute 1 to 4
times with distilled water. Stain 12 to 24 hours, drain away the stain, dehydrate
rapidly with 95% and absolute alcohol, continuing the differentiation if necessary
with oil of cloves, to the right degree. Remove the oil of cloves with xylene
(thoroughly) and mount in balsam.

PREPARATION FOR STAINING.

§ 132. After the sections are cut and before the process of
staining can begin, certain steps are necessary, such as the removal
of the paraffin in the case of paraffin sections. Furthermore, it is
usually advisable to fasten the delicate sections to the glass slides on
which they are to be finally mounted before beginning the series of
manipulations that are necessary in Staining and Mounting. To
consider the latter first:

§ 133. Handling the Sections. Sections may be carried on
through the staining and mounting processes either (a) not fastened
to the slide, — as free or loose sections, or, (b) fastened to the slide,
which is of the greatest advantage and practically necessary in the
case of serial sections. The methods of fastening the sections to the
slide are different for celloidin and paraffin sections.

§ 134. Free sections. It is seldom necessary or advantageous
to carry paraffin sections through the processes of staining and
mounting not attached to slides because- of their delicacy and the
readiness with which they tear or fray. Celloidin sections may be
more conveniently carried on in this way. Loose sections may be
carried on in watchglasses or larger glass vessels if there are many
of them, the sections either being transferred from vessel to vessel
by means of forceps or a section lifter, or the fluid decanted, care
being taken not to pour off the sections, and the succeeding medium
added. Single sections may be best carried on upon the slide which
must be kept horizontal. When the fluid is to be changed, place a
brush or needle gently on one corner of the specimen and pour off
the liquid, if necessary first absorbing most of it by means of a
pipette; in this way the section may be retained on the slide. If
many sections, not in series, are to be treated in the same manner,
they may be placed in a perforated container, — box or basket, —
and handled as a unit by transferring the container from fluid to
fluid (§ § 144—). When the final step is reached (§ 159) the sec-
tions may be transferred to slides and mounted.

§ 135. Fastening sections to the slide. 1. Celloidin sections.
Sections cut by either the alcohol or the clarifier methods may be
conveniently fastened to the slide in the following manner:

If the sections are transferred to the slide from clarifier or clearer,
absorb the fluid thoroughly by placing over the section some absorbent
paper and pressing it down gently and firmly, repeating the operation
several times with fresh paper. After the oil is well absorbed, with
a pipette drop upon the section enough ether-alcohol to thoroughly
wet it. This softens or dissolves the celloidin and on its evaporation
the section sticks to the slide. Allow the ether-alcohol to evaporate
until the celloidin has again set and the surface of the section looks
dull or glazed and then place it in a jar of 95% alcohol. Do not let
the sections dry.

If the sections are in series, it is better to put the ether-alcohol
on one end of the slide and let it run quickly over the sections and

drain from the other end of the slide, repeating the operation two or
three times if necessary. If the sections are well pressed down and
the clarifier thoroughly absorbed the sections will stick to the slide
under most rough manipulation. If, however, it is found that the
sections tend to float off of the slide in the process of staining, their
adhesion may be insured by using albuminized slides, or removing
the slides from the alcohol (§ 144) and again treating with ether-
alcohol.

Sections cut in alcohol may require more ether-alcohol.

§ 136. Celloidin Sections with Removal of the Celloidin. In
the above method, the celloidin is used in fastening the section to
the slide. As celloidin stains heavily with most basic stains, it is
sometimes desirable to have it removed as is paraffin. In most such
cases, the difficulty may be avoided by employing paraffin as the
imbedding method; where this is not possible, imbed in celloidin as
usual but harden and cut in alcohol. Use 67% alcohol on the
microtome knife, handle the sections with tissue paper as usual
(§ 70), but before transferring them to slide previously prepared
with albumen fixative (§ 138), let as much of the alcohol evaporate
as is possible without the sections drying. They should also be free
from wrinkles. Press the sections down well, remove the tissue
paper and place the slide in 95% alcohol which will harden the
albumen fixative cement. The celloidin may be removed now or just
before clearing by means of ether-alcohol, absolute alcohol, or clove
oil.

§ 137. Free-hand Sections and such made with the freezing
microtome may be fastened to the slide if desired by albumin fixa-
tive or M% celloidin or both. If the first is used, the sections should
be transferred to the albuminized slide from water which should be
largely absorbed and the sections pressed down by tissue paper. If
the latter, press sections to a clean slide with paper, pour on 95%
alcohol, dip cautiously into %% celloidin, drain and place slide in
95% alcohol.

§ 138. 2. Paraffin Sections are most conveniently fastened
to the slide by albumen fixative and spreading with water, as fol-
lows:— Place upon the slide a small drop of Mayer's albumin fixa-
tive* and with a (clean) finger tip spread it into a thin even film.
Place upon the albuminized surface enough distilled water to float

M*ayer's Albumen fixattie. White of egg, 50 c. c. ; glycerin, 50 c. c. ; salicylate
of soda, 1 gram. Shake them thoroughly and filter.

the sections freely. Cautiously and slowly warm the slide over a
small flame, as that of an alcohol lamp, or upon a metal warming table
until the sections begin to spread and straighten out. When the
wrinkles have entirely disappeared, allow the water to cool and then
drain it off, retaining the sections in position. The slide should now
stand 2 to 3 hours or better over night when the water beneath the
sections having evaporated » they will have been brought close to the
albumen fixative. They are now ready for the remaining steps.
Melt the paraffin by warming the slide over a low flame and place it
in xylene (§ § 143—).

§ 139. Albumin fixative and Heat. If the sections are free
from wrinkles or with few wrinkles that can be easily "ironed out,"
place the sections in position upon a slide prepared with albumin
fixative as above, and with a clean finger press the section into the
albumin fixative beginning at one edge of the section and by a rolling
motion of the finger, ironing out any wrinkles that there may be.
It is well to look upon the reverse side of the slide to see if the section
really adheres to the albumen fixative, as in some cases it does not.
Heat the slide gently and slowly over a small flame until the paraffin
melts and begins to run away from the specimen. Keep the paraffin
just melted for a minute or so, and then transfer to the xylene.
Should the paraffin section not adhere to the albumin fixative when
well pressed down, it can in many cases be made to do so by briskly
rubbing the reverse side of the slide with a woolen or silk cloth.

§ 140. J4% Celloidin (collodion). The adhesion of sections
that are particularly valuable or relatively thick may be ensured by
treating them with %% celloidin, as follows : Fasten the sections to
the slide by either of the above methods, remove the paraffin by
xylene (§ 143), and then after draining off the xylene from the slide,
10 to 13 seconds, it is put into a bottle containing J4% celloidin. In
a minute or more the celloidin displaces the xylene and penetrates
the sections. The slide is removed, allowed to drain for half a minute
and then put into a jar of 67% alcohol which sets the celloidin [17].
It is now ready for the staining processes (§ 144 — ).

PRELIMINARY STEPS.

§ 141. These are somewhat different for paraffin and celloidin
sections. In the case of the former, it is necessary to remove the
paraffin by means of a solvent (e. g., xylene), remove the paraffin

solvent by alcohol, and usually remove the alcohol with water. In
the case of celloidin sections the first step is unnecessary. If the
tissue was hardened and cut in alcohol, the second step may likewise
be omitted.

§ 142. In toto Staining. If the staining has already been
done, (§ 79) these "preliminary steps" are of course all unnecessary,
save the removal of the paraffin by xylene.

§ 143. Xylene. Leave paraffin sections in xylene until the
paraffin is entirely dissolved out, requiring usually only a few seconds.
A longer stay generally does no harm.

§ 144. Alcohol. Transfer paraffin sections from xylene to
95% alcohol leaving the sections in the alcohol 5 to 10 minutes; or
if you wish, shorten the period to a minute or so by waving the slide
gently to and fro in the alcohol.

Celloidin sections cut by the clarification method, are placed
in alcohol to remove the clarifier. This may take a longer time, and
if there are many slides it is well to use two changes of alcohol. A
longer stay in alcohol does no harm.

§ 145. Water. Remove the 95% alcohol with water if the
stain is an aqueous one.

§ 146. Staining. The following schema shows the general
steps in staining and mounting. In all the processes, seemingly
complicated, if it is remembered that the succession of media as in
histological technique generally depends upon their miscibility or
some special reaction, and the reason for the various steps is recog-
nized, much of the difficulty in remembering the order in which
they come will be avoided.

SCHEMA FOR THE STAINING OF SECTIONS.

Celloidin Sections Paraffin Sections

fasten to slide not fastened to slide

(a) absorbing clarifier

and

}

(b) flooding with a few
drops of ether-alcohol

xvlene*-

95 % alcohol

water

Aqueous stain
(e. g., hematoxylin)

water

Aqueous
Counter-stain
(e. g.,eosinor
picro-fuchsin)

water, or 67% alcohol

Alcoholic

Counter-stain

(e.g., orange G.)

alcohol

Dehydrate
(95%-99% alcohol)

1
Clear

1
Mount in balsam

fasten to slide

albumen fixative,

spreading with

water and drying

Alcoholic stain
(e. g., Hcl. carmine)

alcohol

MOUNTING.

§ 147. Whether stained or unstained, prepared for micro-
scopical examination by isolation or sectioning, and especially if it
is desired to keep the preparation, it is necessary to mount it in
some way, — i. e., so arrange it upon some suitable support (glass
slide) and in some suitable mounting medium that it may be satis-
factorily studied with the microscope.
Mounting may be

I. Temporary, or
II. Permanent, — as

A. Dry7, or in air,

B. In a medium miscible with water, or

C. In a resinous medium, in which case it is necessary
first to remove all water by either (a) drying — Desiccation, or (6)
a series of displacements, i. e., I. Removing the water with strong
alcohol — Dehydration; %. Removing the alcohol with clearer —
Clearing:, 3. Replacing the clearer with balsam or other resinous
mounting medium.

§ 148. Temporary mounting. Illustrations may be found in
the examination of blood corpuscles and living ciliated cells (§ 35).
Temporary examination of tissues is quite simple, though important,
and for this it is only necessary to place the teased tissue or section
on the slide in a drop of the fluid in which it is at the time, normal
salt solution, dissociator, or alcohol, and cover. The examination of
preparations intended for permanent mounts during the staining
or before mounting will often serve to detect faulty treatment at a
time when it may be remedied without great expenditure of time, or
to discard the specimen as worthless.

§ 149. Permanent mounting. These usually include (a) mount-
ing dry on a ring or in a cell, (6) in glycerin or glycerin jelly, media
miscible with water, and (c) in Canada balsam or damar, resinous
media.

§ 150. Mounting dry. The preparation, may be either upon
the under side of the cover-glass (best if possible) or rest upon the
bottom of the cell.

In the first case a shallow cell made by a shellac ring will be
sufficient; in the second, a shellac ring may not give a deep enough
> cell and a paper, hard rubber, or metal ring may be cemented to the
slide.

(a) When the preparation is on the cover. Prepare a shellac
cell (§ 156) on the slide of a size slightly smaller than the cover to
be used, and allow it to dry for a day or so. Warm the cover bear-
ing the preparation to remove the last traces of moisture, and place
it film side down upon the ring. Warm the slide until the edge of
the cover may be made to adhere to the shellac ring and press the
cover down until it adheres all the way round. Seal the cover with
shellac and label (§ § 163, 165).

(b) Mounting in a paper or rubber cell. With a brush, cover
one side of the ring with a layer of shellac and place it on the center
of the slide, shellac side down; place within the cell the prepara-
tion, arranging it in the manner desired, and place upon the ring a
cover-glass of a suitable size, and seal it with shellac; label.

§ 151. Mounting in glycerin media, (a) Pure glycerin; (b)
glycerin and acetic acid, 1%; (c) glycerin and a stain. As glycerin
extracts most stains (Ex. Carmine) it is sometimes advisable to
have a small amount of stain dissolved in the glycerin used for
mounting. Of such combined mounting and staining mixtures may
be mentioned (1) Glycerin and Congo red, (2) Glycerin and car-
mine. Other combinations may be used.

§ 152. Congo Glycerin. Formula: J/2% aqueous solution,
Congo Red, 1 part; glycerin, 1 part.

§ 153. Carmine Glycerin. Formula: — Carmalum, 25 c. c.,
Glycerin, 75 c. c.

§ 154. Methyl green and Eosin Glycerin. Formula: — 1%
aqueous solution methyl green, 2 c. c.; >£% aqueous eosin, 1 c. c.,
glycerin, 97 c. c.

Glycerin and glycerin-jelly are most serviceable in mounting
isolation preparations. For both of these mounting media the ob-
ject must be mounted from water or an aqueous solution.

Arrange the section or teased tissue in the center of the slide,
drain off the water or aqueous solution in which the preparation is
and add a small drop of glycerin. Take a clean cover in the forceps,
breathe on the under side and carefully lower it upon the object;
gently press it down. It is best to use only a small drop of glycerin
so as not to get it outside the cover, as it is hard to clean
away satisfactorily. Clean carefully and seal with shellac in
accordance with § 163.

§ 155. Mounting in glycerin-jelly. The preparation should
be mounted from some aqueous solution. Warm the slide gently
and put it upon the centering card; in the center of the slide place
a drop of warmed (melted) glycerin- jelly. Remove the object from
the water or aqueous solution and arrange it in the glycerin -jelly.
Grasp a cover-glass with the fine forceps, breathe on the lower side,
gradually lower it upon the object and gently press it down. Allow
the glycerin-jelly to set, keeping the slide horizontal meanwhile.
Scrape away the superfluous glycerin- jelly around the cover-glass
and seal with shellac (§ 163).

§ 156. Preparation of shellac mounting cells. Place the slide
upon the turn-table and center it (i. e., get the center of the slide
over the center of the turn-table). Select a guide ring on the turn-
table which is a little smaller than the cover-glass to be used; take
the brush from the shellac, being sure that there is not enough
cement adhering to it to drop. Whirl the turn-table and hold the
brush lightly on the slide just over the guide ring selected. An
even ring of the cement should result. If it is uneven, the cement
is too thick or too thin or too much was on the brush. After a ring
is thus prepared, remove the slide and allow the cement to dry
spontaneously, or heat the slide in some way. Before the slide is
used for mounting, the cement should be so dry when it is cold
that it does not dent when the finger nail is applied. A cell of con-
siderable depth may be made with shellac by adding successive lay-
ers as the previous one dries.

§ 157. Mounting in balsam: by desiccation. Certain prepara-
tions may be mounted in balsam, by drying, the method of desiccation
(§ 147), e. g., cover-glass preparations of bacteria, stained cover-glass
preparations of blood, etc. For this is it only necessary that the prep-
aration be absolutely dry, a small drop of balsam placed upon it or
upon the under side of the cover-glass, which is carefully placed
over the specimen and pressed down.

Mounting in balsam: by displacement. Mounting in balsam by
desiccation is serviceable for but few preparations in histology, and
in most cases the removal of the water by a series of displacements is
resorted to (§ 147). For this the following steps are necessary:
Dehydration, Clearing, Mounting in balsam.

§ 158. Dehydration. The sections are entirely freed from
water by the use of 95% or absolute alcohol. The slide or free sec-

tion may either be placed in a jar of alcohol or alcohol from a pipette
be poured over it. Treat the preparation to be mounted for 5 to 15
minutes. The thicker the section the longer the time required;
celloidin sections require a longer time than paraffin sections. In
any case, be sure that the dehydration is complete, giving a longer
rather than a shorter time, and then clear.

§ 159. Clearing. This is accomplished by putting the slide in
a jar of clearer or dropping the clearer upon the section from a
pipette. When the section is cleared it will be transparent. Test
it by holding it against a dark background; if it is not cleared it
will be cloudy, white, and opaque. Carbol-xylene (melted carbolic
acid, 1 part; xylene, 3 parts); xylene; or certain essential oils (ori-
ganum, thyme, cajuput, bergamot) are used.

§160. Mounting in balsam. Drain off the clearer and allow
the section to stand until there appears the first sign of dullness
from evaporation of the clearer from the surface. Then place a
small drop of balsam upon the section or upon the cover-glass \vhich
is then inverted over the specimen.

Remember that in mounting in this way you must always "De-
hydrate, Clear and Mount in Balsam,''' and that the three steps are
inseparable.

§ 161. Natural balsam is acid in reaction due to organic acids contained.
As these bleach basic dyes, notably hematoxylin, it is well to use for most purposes
balsam that has been dried out and redissolved in a known solvent, such as
xylene or benzene. Neutral balsam solutions are to be preferred. Alkaline
balsam is sometimes preferable for some hematoxylin stains; acid balsam in
certain other cases (fuchsin acid, injection masses, § § 118, 217 — ). Furthermore,
according to Mann [36] , such solvents as xylene readily oxidize with the formation
of acid products. For delicate work, therefore, it is probably advisable to use as
thick balsam as is convenient, and avoid inclusion of air bubbles in the mounting.
Benzene balsam from this standpoint is preferable to xylene balsam.

Certain resinous mounting media, — Camsal Balsam, Euparal, — have been
recently prepared in which specimens may be mounted direct from 95% alcohol;
clearing is then unnecessary, of course.

SEALING THE PREPARATIONS.

§ 162. Sealing glycerin mounted specimens. Wipe away the
superfluous glycerin as carefully as possible with a moist cloth or a
piece of lens paper. Place four minute drops of cement carefully at
the edge of the cover at the four quarters and allow them to harden
for half an hour or more; these will anchor the cover-glass and the

preparation may then be placed upon the turn-table and a ring of
shellac cement put round the edge while revolving the turn-table.

§ 163. Sealing glycerin- jelly mounts. Allow the glycerin-
jelly to harden for 12 hours or longer. With a knife scrape away
the superfluous jelly and then carefully wipe around the coyer-glass
with a cloth moistened with water. Place the slide on a turn-table,
carefully center the cover-glass, and with a brush seal the edge of
the cover by a ring of shellac while revolving the turn-table. A
second coating may be given subsequently if needed, after the first
coat has dried.

§ 164. Sealing balsam mounts. This is necessary only with
special preparations, and should in any case be done only after the
preparations have dried out for several weeks. With a knife scrape
off all superfluous balsam from around the cover-glass and wipe it
carefully with a cloth moistened with alcohol or benzin (or xylene).
Seal as with glycerin- jelly mounts.

LABELING MICROSCOPIC SLIDES.

§ 165. Every permanent microscopic preparation should be
carefully and neatly labeled in ink, the label being placed upon the
right hand end of the slide. The label should furnish at least the
following information:

EXAMPLE.

(1) The number of the prepara-
tion, the thickness of the
cover-glass and of the sec-
tion.

(2) The name, kind, and source
of the preparation.

(3) The fixer and the stain.

(4) The date of the specimen.

No.

Ileum of Cat.
Transection.

Z.*H.&E.

November, 1898.

C. 15

In the case of specimens with which it is advantageous to have
more information at hand a second label may be placed upon the
other end of the slide, and it may bear the following information:

*It is convenient to adopt a standard system of abbreviations, thus: — Z. =
Zenker's fluid; He. = Kelly's fluid; M. = mercuric chlorid, etc. ; H. = Hema-
toxylin; E. = eosin, etc.

(1) Mode of fixation (detail) .

(2) Imbedding method.

(3) Stains employed (detail).

(4) Mounting medium (generally not necessary).

(5) Special purpose of the preparation.

A catalog giving the full data of the specimen, — age, condition
of the animal, mode of preparation in detail, special points illus-
trated, etc., is valuable particularly in special investigations and
with standard specimens.

Paper labels are very convenient but possess the disadvantage that they are
very apt to come off. In this laboratory it is the custom to label permanent
preparations by means of a writing diamond with the record number at least.

§ 166. A carbon ink label written upon the slide itself is fairly durable and
may be made as follows: First coat the end of the slide with a thin solution of
balsam (xylene or benzene), and permit this to dry thoroughly. Upon this surface
it is possible to write neatly with carbon ink and after the ink is dry a second coat-
ing of the thin balsam or of shellac may be given. This method is recommended.

SLIDES AND COVERS.

The slides and cover-glasses used in histological work are often
slightly greasy and should be cleaned before using. After they are
cleaned they should be handled by the edges only, or with forceps.

§ 167. Cleaning Slides. For ordinary work it is enough to
wipe the slides out of clean water to which about 5% ammonia has
been added. A clean glass towel free from lint should be used and
after the slides are cleaned they should be stored in covered glass
jars away from the dust.

§ 168. Cleaning Cover-glasses. Place them in 95% alcohol
to which 1% of hydrochloric acid has been added. A clean, soft
cloth such as an old linen handkerchief, gauze, etc., should be used
for wiping them. In wiping a cover-glass "grasp it by the edge with
the left thumb and index. Cover the right thumb and index with
the cleaning cloth; grasp the cover between the thumb and index
and rub the surfaces keeping the thumb and index well opposed on
directly opposite faces of the cover so that no strain will come upon
it, otherwise the cover is liable to be broken." (Gage, 17).

Cover-glasses when cleaned should be stored in covered glass
boxes, or in Petrie dishes.

§ 169. Cleaning Mixture for Glass. For special purposes, such as when the
slide or cover-glass is to be used in the preparation of blood smears, the cleaning

mixture whose formula is given below may be used. Place the cover-glasses in
this mixture, one by one, and permit them to remain over night or longer. Rinse
them thoroughly in running water until all color of the dichromate has disappeared
rinse them again in distilled water and transfer to 95% alcohol, out of which they
may be wiped.

§ 170. Dichromate Cleaning Mixture. Formula: — Potassium dichromate,
200 grams; water, 800 c. c.; strong sulphuric acid, 1200 c. c. Dissolve the
dichromate in the water by the aid of heat, and to the solution add slowly the
sulphuric acid. The two fluids should be mixed in a lead-lined kettle [17].

§ 171. Used slides and cover-glasses, vials and other glassware that have
been used with balsam, cedarwood oil, or other oily substance, etc., may be
cleaned by boiling them with a solution of strong soap, such as "gold dust," one
or more changes. Used xylene or toluene is sometimes useful. Slides and covers
may require a second cleaning with the cleaning mixture. If only water, glycerin
or glycerin- jelly has been used on them, they may be cleaned with water, prefer-
ably warm water, and then, if necessary, wiped out of 50% alcohol.

§ 172. Measuring the thickness of the cover-glasses. With the cover-
glass measurer determine the thickness of the cover-glasses and sort them into
three groups: (a) those with a thickness of .13-.17 mm., (b) those less than
.13 mm., and (c] those thicker than .17 mm. Groups (a) and (b) only should be
used; (c) should be discarded or used only with objects for low magnification.

It is advantageous to know the thickness of the cover-glass on an object
for the following reasons: (a) That one do not try to use objectives in studying
the preparation of a shorter working distance than the thickness of the cover-
glass [17] ; (b) In using adjustable objectives with the collar graduated for different
thicknesses of cover, the collar might be set at a favorable point without loss of
time; (c) For unadjustable objectives the thickness of cover may be selected
corresponding to that for which the objective was corrected [17]. Furthermore if
there is a variation from the standard one may remedy it in part at least by
lengthening the tube if the cover is thinner and shortening it if the cover is thicker
than the standard [17].

THE MICROTOME KNIFE.

Finally, the microtome knife or section razor should receive a
passing word as upon it depends far more than may be at first sus-
pected the excellence of results. Scrupulous care should be taken
to maintain a keen edge, smooth and free from nicks or corrosion.
Never touch the edge with anything hard or metallic. Keep it clean.
Before and after using it is advisable to strop it upon a strop backed
with wood, and occasionally it will be necessary to hone it.

§ 173. For microtome knives two grades of hones are service-
able; the yellow Belgian for first sharpening and either a blue-green
water hone or a fine Arkansas oil stone for finishing. These should be
kept clean and free from dust. If a section knife is used a great

deal, it is best to put it on the fine hone for a short time before each
day's sectioning. Unless experienced in sharpening section knives,
it is well for the first few times to work under the direction of one
skilled in the manipulation. There are, however, excellent accounts
to be found in works on technique [e. g., 17, 21a].

Do not be satisfied with any but a smooth edge, keen enough to
cut without "pulling," a hair held in the fingers, a quarter of an
inch or more from the fingers. When once a good edge is secured,
take pains to preserve it.

With the yellow hone, use a lather of olive oil soap; the blue-
green soap stone is rubbed up with water; with the Arkansas stone
apply a good thin oil.

SPECIAL METHODS.

THE CELL.

The technique of the Cell is almost coextensive with that of his-
tology as a whole, at least as far as concerns the application of the
more exact and delicate methods. It is necessary therefore to give
here only the more salient points and accepted methods.

§ 174. General Methods. Hermann's fluid, Flemming's fluid,
Zenker's fluid, Mercuric chlorid, Carney's fluids, and Picro-aceto-
formol are standard fixers, although special problems may demand
other combinations. Iron Hematoxylin, the Ehrlich-Biondi-Heiden-
hain triple mixture, Safranin, Gentian Violet and Orange G. are per-
haps the most serviceable stains. Of these iron hematoxylin is
particularly and universally useful. The Ehrlich triple stain is
valuable in the more analytical work and should follow Mercuric
chlorid (or Zenker's fluid, Carnoy's fluid) fixation. Safranin and
gentian violet, separately as red or blue stains, or successively fol-
lowed by orange G (Flemming's triple stain) only after Hermann's
fluid, or Flemming's fluid, or similar mixtures.

§ 175. Chromatin. While regressive stains of the iron hema-
toxylin type give valuable chromatin stains, they are not as a rule
analytical or selective. In accordance with the recommendation of
Heidenhain it is better to stain progressively with dilute solutions
if a pure chromatin (basichromatin) stain is desired.

Methyl green is one of the most delicate and precise of chro-
matin stains. It may be used alone or in combination, as in the
Ehrlich-Biondi mixture (§111). A very dilute hematoxylin (§ 88,
92) is excellent.

§ 176. Nucleoli. The Ehrlich-Biondi may be recommended
to bring out other nuclear structures, such as nucleoli, although
other combinations of basic and acid stains may be used. Mont-
gomery recommends Ehrlich's hematoxylin followed by a strong
aqueous eosin.

§ 177. Cytoplasm. Quite different pictures are obtained by
the use of fixers as similar in composition as Flemming's fluid and
Hermann's fluid, the difference seeming to be due to a varying
preservation of cytoplasmic "granules." The ground work of the
cell body, the so-called Spongioplasm, may be preserved by such
fixers as Zenker's fluid, Flemming's fluid, Hermann's fluid, Carnoy's
fluid, etc., and iron hematoxylin and the Ehrlich-Biondi mixture give
satisfactory stains. The granules that may be present are of different
kinds and often not easily interpreted; they include, — (a) "re-
serve" material (yolk granules, fat granules, etc.), (b) basophile,
acidophile, neutrophile granules, (c) granules less easily preserved
which include many secretion granules, etc.

(a) Yolk granules. If present in large amount a special fixer
may be indicated (§ § 20, 21, 22).

Fat granules', see § § 224.

(b) Basophile granulations (Granoplasma, Unna), may be
demonstrated by simple alcohol fixation and subsequent differential
staining with basic dyes. Compare § 195 and the special technique
of blood (§ 21 1-).

(c) These require special technique: in general, (1) the employ-
ment of oxidizers such as dichromates, osmic acid, formalin, (2)
no acid, or a minimum amount. This seems to indicate the presence
of reducing substances usually lipoid in nature whose combinations are
soluble in (or rendered soluble by) acid. Here belong the Mitochon-
dria of Benda.

§ 178. Mitochondria. Benda's Method.

1. Fix in Flemming's fluid with the acid reduced to 3 drops; 2.
Rinse in water, 1 hr.; 3. Place in equal parts pyroligneous acid and
1% aqueous solution of chromic acid, 24 hours; 4. 2% potassium

dichromate, 24 hours; 5. Running water, 24 hours; 6. Alcohols;
paraffin imbedding; 7. Sections 5^; 4% ferric alum for 24 hours;
8. Rinse in distilled water and — 9. Place in a mixture of saturated
alcoholic solution of sodium sulphalizarinate 1 c. c., distilled water, 80
to 100 c. c., 24 hours; 10. Rinse away the stain with water; 11.
Stain on the slide with a freshly prepared mixture of saturated alcoholic
solution of crystal violet and aniline water, equal parts, warming
until it steams; 12. Rinse in w^ater and differentiate in 30% acetic
acid (1 minute or less) ; 13. Rinse in running water 5 to 10 minutes
to remove the acid, drying with blotting paper; 14. Dehydrate
rapidly in absolute alcohol, (dip); 15. Clear in bergamot oil and
xylene; 16. Mount in neutral balsam.

The above method is capricious. The following method is recom-
mended: (1) fixing 12 to 24 hours in Zenker's fluid or the Copper
dichromate mixture (§ 16) with 1/10% acetic acid only; (2) mor-
danting 3 to 4 days in 2.5% dichromate (or Muller's fluid or Erlicki's
fluid respectively); (3) paraffin imbedding; (4) using iron or copper
hematoxylin as the stain (§ § 94, 95). Benda's fluid may be substi-
tuted for the fixer, and a number of other stains may be used; con-
sult [6, 30].

§ 179. Secretion Granules in the cytoplasm, such as the zymogen
granules in the stomach and pancreas, granules of the suprarenal
medulla and Islands of Langerhans, etc., may be preserved and
demonstrated in the same manner and probably for the same general
reasons ; thus : —

1. Trypsinogen granules. Fix in 1% osmic acid which pre-
serves and browns them; or as under 2 below.

2. Pepsinogen granules. Fix in Kelly's fluid or one of sim-
ilar composition. Iron hematoxylin or Weigert's copper hema-
toxylin, or neutral stains (§ § 107, 128, 131) may be used.

3. Medulla of suprarenal. Apply similar fixation methods, e. g.,
Kelly's fluid, and regressive staining with basic stains (e. g., iron or
copper hematoxylin, toluidin blue, etc.).

CONNECTIVE TISSUE

§ 180. White (Collagenous) Fibers. Fuchsin acid is particularly
valuable. Three methods of applying it for the differential staining
of connective tissue follow :

(a) Picro-fuchsin. See § 119. This may be used with or with-
out a basic counter-stain which should precede it. If a counter-
stain is used remember to overstain and use the picro-fuchsin to dif-
ferentiate it.

(b) Orange-fuchsin. Formula: Fuchsin acid, 2 grams; Orange
G., 1 gram; Glycerin, 7 c. c.; distilled water, 100 c. c. Fix tissue in
Flemming's fluid. Stain sections 30 seconds; dehydrate, clear, and
mount in balsam (not alkaline) . Suitable also for staining the reticu-
lar tissue (lymphatic tissue).

Somewhat more delicate than the picro-fuchsin.

(c) Mallory's connective tissue stain. See § 120. While not a
differential stain for collaginous fibers, it is nevertheless a valuable one,
and frequently to be preferred.

§ 181. Elastic Fibers (Elastin). Employ either the Weigert
ResorchvFuchsin or the Orcein methods (§ § 125, 126), or the Verhoeff
[44] method (§ 127).

Both the white and elastic fibers may be stained in the same
preparation, the elastic fibers being stained first.

In a mixture composed of white and elastic fibers, picro-fuchsin
(§ 119) will stain the elastic fibers a light yellow, the white fibers being
colored red.

In studying the connective tissues, it should be remembered
that acetic and the mineral acids cause swelling or solution (gelatini-
zation) of the white fibers, depending upon their strength. While
this improves the cutting quality of organs rich in connective tissue,
it also causes vagueness in outline of the white connective tissue
fibrils. It may therefore be advisable to decrease the percentage of
acetic acid in the fixer when the connective tissue is under investi-
gation.

§ 182. Reticular Tissue. Fuchsin acid, Mallory's connective
tissue stain, or the orange-fuchsin acid mixture may be chosen for
staining this form of connective tissue. As however the cellular ele-
ments usually mask the fiber relations, if a view of the latter is desired
the cells must be removed, — by a mechanical method, such as cau-
tiously brushing the section with a camel's hair brush, or by digestion.

§ 183. Digestion Method. The organ to be examined (e. g.,
lymphatic node, spleen, etc.), is preserved in 67% alcohol, cut into
sections 2 mm. or more in thickness, washed thoroughly in water to
remove the alcohol, and digested with pancreatic solution in 1%

sodium carbonate solution in an incubator at 38 C. until examination
under the microscope shows that the cells have become disintegrated
and digested. The digestion fluid should be changed every day or
2 or 3 days. A varying length of time is necessary, — sometimes a
month or more. Paraffin sections may also be submitted to digestion,
with or without the removal of the paraffin.

The following method is recommended for the further preparation
of digested tissue. After washing thoroughly in running water to
remove the digestion fluid, the tissue is carefully imbedded in celloidin
(§ 61 — ) and sectioned, the sections being 20 to 30^ thick. These
sections are not fastened to the slide, but after having been brought
into 95% alcohol are placed in a concentrated solution of acid fuchsin
in 95% alcohol to which a drop or two (per 50 c. c.) of glacial acetic acid
has been added. After several minutes, the sections are rinsed in
95% alcohol to remove the excess stain, cleared in carbol-xylene and
mounted in balsam. By this method the delicate morphology is
preserved and a sharp stain is secured.

The method of artificial digestion with trypsin or pepsin has
other applications in the histological analysis and it possesses a dis-
tinct value.

CALCIFIED STRUCTURES — BONE AND TEETH.

(A) . Decalcification .

§ 184. For the purpose of investigating the soft structures of
tissues containing lime salts, such as bone, teeth, and calcified carti-
lage, it is necessary to remove the lime salts before sections can be
prepared in the usual way by a process known as decalcification.
Solutions of a large number of acids, combined or uncombined with
other substances, may be used as decalcifiers. Very satisfactory
are: (1) Hydrochloric acid, 1 c. c., 67% alcohol, 100 c. c.

(2) nitric acid, 3 c. c.; 70% (67%) alcohol, 97 c. c., and

(3) nitric acid, 5 c. c.; saturated aqueous solution of (potash)
alum, 50 c. c.; water, 50 c. c.

In the first and second formulas the alcohol, in the third the
alum acts as a restrainer of the acid. The first or second of these
formulas is, perhaps, better for bone; the second has a more rapid
action and is possibly a better decalcifier for teeth. It is better to
let the decalcification proceed slowly for a longer time in an abundance
of fluid changed often, in order that the carbon dioxid may not be

formed too fast, accumulate in the tissues, inflate and distort them.
Many fixers contain acid (e. g., see § § 12, 14, 17, etc.), and in their
action give decalcification enough in the case of small calcified
objects.

§ 185. Directions for use. [10.] The tissue to be decalcified
had best be first thoroughly fixed and hardened by one of the approved
methods, and should be in 82% alcohol. In fixing, structures not
needed should be removed, — muscles trimmed away from the bone,
etc. Bones or teeth should be opened with nippers or a saw, so that
the fluid may reach the marrow or pulp cavity.

Place the hardened tissue in the decalcifier, where it should
remain until the lime salts have been entirely removed, as may be
ascertained by inserting a fine needle ; if any calcified matter remains
there will be a gritty feeling on using the needle. The time neces-
sary for complete decalcification will depend upon the size and den-
sity of the calcified tissue, and will vary from 3 to 15 days or longer.
The decalcifier should be changed after the first day, and if the tis-
sue is large it is best to change it subsequently two, three or more
times at intervals of several days.

When decalcification is complete rinse the tissue well in water or
67% alcohol for a few minutes and place it in 67% alcohol for one or
two days and then in 82% alcohol for several days, or until ready to
imbed. The 82% alcohol should be changed once or twice in order
that the nitric acid may be well wrashed out. Although paraffin in
many cases may be employed for imbedding, the celloidin method is
generally more satisfactory.

Hematoxylin with eosin, hematoxylin with picro-fuchsin, and
hematoxylin with picro-carmine afford good stains; by staining
thoroughly with hematoxylin a differential staining of bone and
cartilage may be obtained. Mallory's connective tissue stain fre-
quently gives interesting pictures.

(B). Sections of Dry Bone or Tooth.

§ 186. Though the general structure of bone and tooth is shown
moderately well when the tissue has been decalcified (§ 184). the
Haversian canals, canaliculi and lacunae of bone and the dentinal
tubules of the teeth are shown much better in sections of dried,
non-decalcified, tissue rendered sufficiently thin for microscopic ex-
amination by grinding or filing.

§ 187. Directions for procedure. Prepare thin transverse sec-
tions of dried bone in accordance with the directions below. Longi-
tudinal (radial) sections and tangential (surface) sections may also
be prepared in the same manner, the former to show the Haversian
canals and their anastomoses, the latter to indicate the shape of the
lacunae as seen in a different plane.

1. Sawing the section. Make an exact transection of a part of
the shaft of a long bone. The section should be about 1 cm. long
and include the thickness of the shaft from the surface to the medul-
lary cavity. Make the sections about 1 mm. thick.

2. Grinding the sections. Place the piece of bone on a cork or
piece of soft wood and wet it with water. File it on one side until
smooth and then turn it over. Continue the filing till the piece is
from .05 to .10 mm. thick, using the cover-glass measurer to deter-
mine the thickness. In the beginning one can press quite hard in
filing; as the section thins, more care should be exercised and the
pressure should lessen.

A grinder, such as a fine carborundum wheel or emory wheel con-
nected with a variable speed electric motor is very useful and greatly
expedites the preparation of the sections. The carborundum wheel
should be horizontal and the sections ground on the flat surface of the
wheel, water being used to carry away the bone dust.

3. Washing and drying the section. When the section is thin
enough, rinse it and dry it with lens paper.

4. Mounting the sections in hard balsam. To prepare the bal-
sam, put two or three large drops on the middle of a slide and heat
the slide in some way to drive off the volatile constituents. Do not
heat the balsam hot enough to produce bubbles. When the balsam
chips after cooling, it is ready for use.

In mounting, have the section and a clean cover so placed that
they may be easily and quickly grasped. A cork somewhat smaller
than the cover-glass should be within reach, and also a stone or
piece of glass upon which to quickly cool the specimen as soon as it
is mounted.

Heat the slide until the balsam is well melted. Put the slide
upon a piece of paper, grasp the piece of bone with the forceps and
plunge it into the melted balsam; put on the cover as quickly as
possible and press it down with the cork; finally put the slide on
the stone or glass to cool the balsam quickly. All of this should be

done as rapidly as possible, and if done rapidly, the air will be retained
in the lacunae and canaliculi, and cause them to stand out as black
spots and lines. If soft balsam were used it would soon drive out
the air, and being of nearly the refractive index of bone, it would
obliterate the lacunae and canaliculi. Further, if the hot balsam
were not cooled quickly, the air would be driven out and balsam
would take its place in the spaces.

MUSCLE.

§ 188. Fresh. Much of the investigational work on muscle
has been done on fresh muscle, or frozen sections. For examination
fresh it is advantageous to have very thin muscles. Of the several
muscles that have been recommended, one of the most available is
the M. cutaneus pectoris of the frog. This may be prepared by
cutting the skin in the midventral line, cutting at right angles to
the first cut across to the angle of the jaw, thence caudally parallel
to the first cut. The skin flap so formed contains the insertion of
the muscle which may now be easily dissected free and removed to-
gether with some of the tissue at insertion and origin to handle it by.
It may be used for examination fresh, with the polarization micro-
scope, etc.

§ 189. Isolation Methods. Nitric acid (§ 44) may be used for
plain muscle and for skeletal muscle. Potassium hydroxid (§ 45)
is suitable for heart muscle.

(a) Nitric acid. Place in the nitric acid dissociator the fresh
striated muscle, gland or organ containing the muscle, — (plain
or striated,) — that it is desired to isolate. If it is the intention
to w^ork out the anatomy of the muscle or the relation of the muscular
coats in an organ, the entire muscle or organ should be taken; other-
wise, portions will suffice. At the ordinary temperature of the
laboratory the dissociating action will have been sufficient in from
1 to 3 days; test at intervals with needles to ascertain whether
the fascicles and fibers can be easily separated; or fragments may
be shaken in a test tube or vial with water in order to separate the
fibers.

When the dissociation is sufficient pour off the acid and wash
the muscle gently but thoroughly with water. If the tissue is to be
stained with hematoxylin or carmine, or kept for any length of time,
drain off the water and add a half -saturated solution of alum. For

permanent storage, pour off the alum solution and place successively
in 67% and 82% alcohol.

For temporary examination, tease out a portion of the muscle in
water, separating the fibers carefully by means of needles ; cover and
examine.

Permanent preparations, (a) unstained. After teasing out with
the needles drain off the water and add a small drop of glycerin
or glycerin-jelly; cover, and seal after first properly cleaning (§ § 162,
163) . (b) stained. — Either employ the nitric acid method given above
or (better; Badertscher) dissociate at incubator temperature (about
38° C.) in a sat. solution of mercuric chlroid with 10% of strong nitric
acid added. Test at intervals and when dissociation has proceeded
far enough, wash out the dissociator with water and 67% alcohol.
Stain with carmine or hematoxylin. Mount in glycerin (§ 154-),
glycerin -jelly (§ 155), or dehydrate, clear and place in dilute balsam
(§ 157- Cf. § 215).

(b) Potassium hydroxid. Place in the fluid small pieces of
the heart muscle of a fetal, new-born or young animal; after 10 or 15
minutes, the tissue should be tested with needles at intervals of about
five minutes, so that the action may not be too prolonged; probably
15 to 30 minutes will suffice. As soon as the elements separate
readily, pour off the caustic potash solution and add an abundance
of 60% solution of potassium acetate (potassium acetate, 60 grams;
distilled water, 40 c. c.). Take small fragments and tease them in
this solution, or shake them in a vial, until the 'cells' are separated
from each other.

For temporary examination, cover, in a drop of the potassium
acetate solution. For permanent preparations, drain off the potas-
sium acetate solution and add a small drop of glycerin or glycerin-
jelly.

Stained preparations. Pour off the potassium acetate solution
and add a half saturated solution of alum, letting it remain for 24
hours or longer. Tease in water, stain with hematoxylin or carmine,
wash away the stain with water, and add a drop of glycerin or gly-
cerin-jelly. Cover and seal (§ 162 — ).

If a large amount is desired, the tissue may be carried through
the various steps in a vial.

§ 190. Sections. To bring out the structure of the fibrillae
picro-aceto-formol (§ 17) or an alcoholic fixer (§ § 23, 24) is preferable

although the sarcoplasm is not so well preserved. The muscle should
be moderately distended upon cork, before fixing, the ends secured
by small pins. 10% formalin or Zenker's fluid or Mercuric chlorid
may also be used, the sarcoplasm being much better fixed in di-
chromate, osmic acid or formalin mixtures. Iron hematoxylin is
particularly indicated as a stain for muscle (§ 94). Mallory's con-
nective tissue stain (§ 120) is found to give an excellent differentiation
(Kingery).

To differentiate muscle in situ, picrofuchsin may be used, which
stains muscle yellow or orange, the surrounding connective tissue
red. Mallory's connective tissue stain is also frequently useful.

To differentiate the intercalated discs of cardiac muscle, fix in
mercuro-nitric mixture (§ 20) ; imbed in paraffin ; stain sections with
haemalum (§ 89), diluted, 12 to 24 hours; differentiate with acid
alcohol; dehydrate, clear, mount in balsam.

THE NERVOUS SYSTEM.

An analytical grouping of the numerous methods used in the
study of the finer structure of the Central Nervous System is pre-
mature. The most salient point is the prominent part that reduc-
tion processes seem to play. The more important methods here pre-
sented deal (a) with the finer structure of cell and fiber; — demonstra-
tion of the tigroid substance and fibrillae ; (b) the differential staining
of the myelinic nerve fiber (Weigert and Marchi methods); (c) the
morphology of the elements (neurones) as revealed by the chrome-
silver impregnation methods (Golgi methods) or the use of intra-
vitam (methylene blue) methods.

§ 191. Isolation of Nerve Cells. Employ formaldehyde disso-
ciator for the isolation of the nerve cells of the spinal cord and of
the cerebral cortex, proceeding as follows :

Split the spinal cord along its median plane, separating thus
the two halves, and place it in an abundance of the dissociating
fluid. The cerebral cortex should be cut into small pieces by sec-
tions vertical to the surface. Allow it to remain in the dissociator
from 2 to 24 hours; for the best results a stay in the fluid of more
than 24 hours is not so satisfactory; although isolated cells are
readily obtained their processes are broken off much nearer the cell
body.

Place a fragment of the gray matter of the spinal cord or the
cortex of the cerebrum on a clean slide in a drop of J^% Congo Red
(§ 113) or 1/10% eosin in formaldehyde dissociator; with the blade
of a scalpel crush the tissue, grinding it thoroughly with a rotary
movement, which will reduce it to small pieces. Gather the debris,
drain off the fluid, and add a drop of glycerin containing stain.
Cover and examine, tapping the cover sharply with the handle of the
scalpel to shake out the processes of the cells and free them from
surrounding matter. Examine, searching for cells with many and
long processes, and if a satisfactory preparation, seal according to
§ 119.

§ 192. Isolation of Nerve Fibers, (a) For isolation of myel-
inic nerve fibers, with the preservation and blackening of the myelin,
and for amyelinic fibers, employ osmic acid dissociator (§ 42).

(b) For the isolation of myelinic nerve fibers with the removal
of the myelin for the demonstration of the axis cylinder, neuro-
lemma and framework of the sheath, fix nerves in Dichromate-
acetic or similar fluid one or more days, wash in water, pass up
through the alcohols, dehydrate, remove the myelin by placing the
tissue in a fat solvent, — chloroform, for one or more days, 95% alco-
hol 1 day, pass through the alcohols to water, stain in Delafield's
hematoxylin 12 to 24 hours, wash in water, pass up through the
alcohols, dehydrate and place in clearer. Out of this the small bun-
dles of the nerve fibers may be teased apart with needles, care being
taken to keep the fibers as nearly parallel as possible. Mount in
balsam.

§ 193. Gold Chlorid Methods. These methods, which are
widely serviceable, depend upon the reduction of gold chlorid solu-
tions by the tissues through the agency of (a) sunlight or (b) various
chemical substances of which the acids, formic, acetic, citric, etc.,
are particularly used. Either one or both of these agencies may be
used. Usually fresh tissue is used although the method may be
applied to fixed tissue, particularly as a neurofibrilla stain [6, 30].
It is the method par excellence for staining motor nerve terminations
for which purpose the following method is serviceable.

§ 194. [21]. Fresh tissue or (better) tissue fixed in 10% formalin may be
used. Place small pieces of muscle containing the endings for 30 minutes in 10%
formic acid solution. Remove to 1% gold chlorid for 30 to 40 minutes, avoiding
direct sunlight; the tissue becomes yellow. Transfer again to a 2% formic acid
solution in which the tissue should remain for 1 or 2 days in the dark (rich purple

color). A bluish-purple indicates too great a reduction. Wash in distilled water
for an hour or so.

If it is desired to make a teased preparation, transfer to glycerin in which
the fibers may be cautiously teased apart, taking care not to separate them too
much. Permanent mounts may be made in glycerin or glycerin-jelly.

If sections are called for, dehydrate, clear (xylene method, § 52), and imbed in
paraffin. Celloidin may also be used.

With fresh tissue the treatment by formic acid may cause too
marked a swelling and distortion ; in which case the Ranvier method
[30] may preferably be used.

§ 195. Tigroid substance (Nissl's bodies). The stainable sub-
stance in the cell body of nerve cells resembles the chromatin of
the nucleus in its reactions, staining with basic stains of which a
number are suitable. Alcohol, Carnoy's fluid (§ 26, 27), mercuric
chlorid, or formalin may be used as fixers, preferably one of the first
three. Methylene blue or toluidin blue are the usual stains.

§ 196. NissPs Method (modified). Imbed in paraffin or cel-
loidin tissue that has been fixed in 95% alcohol or Carnoy's fluid;
cut the sections rather thick, 15 to 20^. The sections may either be
fastened to the slide or carried on as free sections. Stain the sec-
tions in a 1% aqueous solution of methylene blue or toluidin blue
for 5 to 10 minutes, heating it until it steams. Permit it to cool,
rinse in water, dehydrate and differentiate in absolute alcohol, clear
with oil of cajuput and xylene and mount in xylene balsam. The
nerve cells and nuclei will be stained blue, all else colorless. In the
cell-bodies of the nerve cells, the tigroid substance will be stained.
Should the stain not be selective enough, differentiate for a few
seconds before dehydrating with a mixture of anilin 1 part, 95%
alcohol 9 parts.

If celloidin is used for imbedding, it should be remembered that
it should be dissolved away in the differentiation (absolute alcohol).

§ 197. H eld's Method (modified). Fix in mercuric chlorid
(or as for Nissl's method), imbed in paraffin, cut sections 5 to 10^
and stain 15 minutes in 1% solution of erythrosin in 67% alcohol,
rinse, stain 10 minutes in 1% aqueous solution of toluidin blue, dif-
ferentiate briefly in 1 /10th% alum solution, dehydrate rapidly in
95% and absolute alcohol, clear in xylene, mount in balsam. Alka-
line methylene blue or Nissl's soap solution (y&% methylene blue in
1/5% soap solution) may be used in either of these two methods if
desired.

§ 198. Neurofibrillae (Simarro-Cajal Methods). Three of Ca-
jal's methods may be given : Formula 3a. 1 . Fixation in ammonia-
cal alcohol (2 to 10, usually 4 to 5, drops of ammonia per 50 c. c. of
95% alcohol), 20 to 48 hours; 2. Mop up with absorbent paper and
3. Place in !>£% silver nitrate solution for 4 to 5 days at 32 to 40° C. ;
the tissue when ripe should be light gray; 4. Wash for a few minutes
in distilled water; 5. Reduce in a solution of 1 to 2 grams hydro-
chinon or py rogallol ; water, 100 c. c.; formalin, 5 c. c.; for 24 hours;
6. Wash in wTater; 7. Imbed by the paraffin method; 8. Section
and mount in balsam or damar. Recommended for spinal cord,
cerebellum, spinal ganglia.

Formula l^a. 1. Fix small pieces of tissue (5 mm. thick or less)
for 6 to 12 hours in formalin, 15 c. c., water 85 c.c.; 2. Wash 6
hours or longer in running water; 3. Place for 24 hours in 50 c. c.
of alcohol with 5 drops of ammonia added; 4. Mop with absorbent
paper; 5. Place in 1^% silver nitrate solution for 4 to 5 days (35
to 38° C.); The remaining steps as in formula 3a. Recommended
for sympathetic ganglia, cerebrum, cerebellum.

Formula 5a. 1. Fix small pieces for 6 to 8 hours in water and
pyridin, each equal parts; then for 18 to 24 hours in pure
pyridin; 2. Wash for several hours in running water; 3. Place
in 90% alcohol for 1 day ; 4. Mop with absorbent paper; 5. Place
in \Yi% silver nitrate solution for 4 to 5 days (35 to 38° C.); The
remaining steps as in 3a and 4a. Recommended for embryonic and
fetal tissue (neurogenesis), regeneration, cerebrum.

By these "photographic" methods, fibrillae, fibrillar networks,
changes in histogenesis, etc., may be demonstrated.

For other methods of demonstrating the neurofibrillae, — Biel-
schowsky's, Bethe's toluidin blue method, Apathy's hematein
method, etc., consult larger wwks on technique [6, 30],

§ 199. The Weigert Method for staining differentially the
myelin of myelinic nerve fibers. This method in all its various
forms, depends upon the power of the myelin, probably through the
reducing fatty acid present, to combine writh and hold in (nearly)
insoluble form the chromium (oxid), which thus serves as a pri-
mary mordant for a copper or iron hematoxylin stain, which is sub-
sequently differentiated by an oxidizer as a bleacher. The important
steps are: — (1) fixing and mordanting in dichromate solutions; usually
the potassium salt is chosen; (2) a second mordantage in copper
(acetate), (3) the staining; (4) the differentiation. The point at
which the imbedding and sectioning are introduced is of secondary

importance. It should be remembered, however, that the fatty sub-
stances (lipoids) of the myelin upon which the method depends are
soluble in the reagents of both the paraffin and cello idin methods, less
so in the latter — and in acids, and that even the dichromate mordan-
tage does not preserve them perfectly. The dichromate mordant age
must thus be given before the imbedding is begun (alcohols) , and pref-
erably with the fresh tissue. Practically the only fixer that is indif-
ferent in this respect and after which the dichromate may first be used
is formalin. Other dichromate fixers such as Zenker's fluid with the
acetic acid reduced to about 1/5% may be used, but it is well to let
them act only a relatively short time, and continue the mordantage
with simple dichromate solutions. Aside from this the point at which
the imbedding and sectioning are introduced is of secondary im-
portance; thus, Strong [41] combines the copper and chromium
mordantage by using copper dichromate; the former may also follow
the dichromate treatment before imbedding is begun, in case of
celloidin imbedding it may be applied to the celloidin imbedded
block, or after the sections are cut. Street er [40] stains (in toto,)
4 to 6 days as well as mordants before the imbedding. Weigert has
added to both the primary and secondary mordants chromium
fluorid to (1) hasten the process and (2) prevent precipitates; the
formulas are: (a) 5% potassium dichromate, 100 c. c.; chromium
fluorid, 2 grams; (b) 5% copper acetate, 100 c. c.; chromium fluorid,
2 grams; glacial acetic acid, 4 c. c. (b) is especially indicated if the
secondary mordantage is given before the sectioning. Whatever
modification of the method is employed, the reduction of the dichro-
mate by the tissue in the primary mordantage should fully reach
the dark brown stage, but not pass it (i.e., become green). Sheldon
[4] gives a good resume of the method. The following method is
serviceable : —

1. Fix tissue for 1 to 2 days in Zenker's (see above) Orth's, or
Kelly's fluid, 10% formalin, Muller's fluid, or potassium dichromate
solution. —

2. Mordant until dark brown in 3% and 5% aqueous solution
of potassium dichromate. This usually takes about 4 weeks; or,
shorten the period by using the dichrornate-chromium fluorid mix-
ture (above) when about 5 days should suffice ;

3. Wash in running water 1 or 2 days, and

4. Pass up through the alcohols, preferably keeping the tissue
in the dark.

5. Imbed in celloidin or paraffin.

6. Stain and differentiate sections by the Weigert copper hema-
toxylin method (§ 95).

7. Mount in neutral or alkaline balsam.

Large sections are usually best carried on as free sections. The
differentiation of the stain should be carefully watched and stopped
when the fibers are a rich dark blue on a yellow-brown-background.
The reagents used in dehydrating, clearing and mounting should
be neutral or alkaline, — not acid.

§ 200. Pal's method may be used if it is desired to stain the
nerve cells subsequently.

Fix and mordant, in the dichromate as above; omit the copper
mordantage; imbed and section, staining the sections with the strong
hematoxylin used in the Weigert method (§ 199) until the sections are
a blue-black.

Rinse the sections in tap water and differentiate by treating for
a short time (20 to 30 seconds) with a 1 /10% aqueous solution of
potassium permanganate and for a few seconds with a mixture of
1% oxalic acid and 1% potassium sulphite, equal parts. The action
will be very rapid and must be carefully watched. Wash the sec-
tions % hour in running water. * Counter-stain with a red stain
(eosin, erythrosin, carmine, etc.) if desired.

§ 201. Marchi Method. This method of staining differen-
tially degenerating myelinic nerve fibers depends upon the fact that
potassium dichromate (or chromic acid) is able to satisfy the re-
ducing power of myelin but does not oxidize the globules of free
fatty acid (?) formed in the degeneration of the myelinic sheath
of the fiber, which may be subsequently blackened by the reduction
of osmium tetroxid. Important points in the successful application
of the method are: (a) the length of time the degeneration should
be allowed to proceed before treating with potassium dichromate,
(b) the time in the dichromate mordant, (c) the time in the osmic
acid mixture (sufficient and complete penetration), (d) the preser-
vation in situ and final mounting of the osmicated fat granules; the
difficulties here are those of fat preservation in general (§ 224).

(a) The optimum will vary and must often be determined
experimentally: in general, — for cold blooded animals; (Toad), 30
to 40 days; for mammals, 12 days.

(b) 8 to 10 days in Muller's fluid or 3% potassium dichromate
is usually enough; a longer time does no harm.

(c) The osmic acid is useful in 1% aqueous solution usually
mixed with a potassium dichromate solution. 6 to 10 days suffice; a
longer time does no harm (brittleness) .

(d) See below.

§ 202. As employed by van Gehuchten [18] : —

1. Harden in 3% potassium dichromate solution for 3 weeks;

2. Transfer to a mixture of 1% osmic acid solution, 1 part;
3% potassium dichromate, 4 parts; for 3 weeks, blocks of tissue
not more than 2 mm. thick. Use abundance of the fluid and change
2 to 3 times if deemed necessary.

3. Wash in running water for 12 to 24 hours.

Avoid the paraffin and celloidin imbedding methods if pos-
sible. Of these two the celloidin method is preferable. For further
treatment, see § 227.

§ 203. Flemming's or Benda's fluids (§ § 19, 227) may be used
instead of the Marchi method for the same purpose with small objects,
and peripheral nerves, etc.

§ 204. The Golgi Methods, whose field of application is not
confined to the nervous system (gland ducts, bile capillaries, blood
capillaries, secretory canaliculi, muscle, etc.) consist in (a) mordant-
ing the fresh or living tissue for a sufficient length of time in a dichro-
mate solution, usually containing as well osmic acid or formalin,
and then — (b) transferring to a silver nitrate solution, whereupon
certain of the nervous elements become outlined more or less com-
pletely by impregnation with a chrome-silver combination.

The reaction probably depends on the presence, in certain "phy-
siological states" of the elements of a substance or substances which
combines with the chromium salt (with reduction?) and through it
with the silver salt. These hypothetical substances, — or possibly
physical states,— seem to disappear more or less rapidly after the
death of the animal and their power to hold the silver in combination
to decrease with the progress of the dichromate mordantage beyond
a certain point. If successful, certain of the cells and their processes,
— amyelinic, and to a certain extent, myelinic nerve fibers, are out-
lined by an impregnation, black by transmitted, brown by reflected
light.

§ 205. The method is, however, capricious; success depends on
(a) the kind of animal; different parts and tissues react more satis-
factorily in some animals or classes of animals than in others, (b)

The age of the animal; some regions of the nervous system give bet-
ter results in young or fetal animals ; other parts take the stain
better in older animals, etc. (c) The time of mordantage; it is neces-
sary that the tissue be mordanted a certain length of time, constant
(relatively) for a certain kind of tissue under the conditions above
(a and b). It is necessary that the best amount of dichromate mor-
dantage be given, (d) Different organs and regions of the central
nervous system vary greatly in the ease with which they can be made
to furnish satisfactory impregnations. Almost certain impregna-
tions of hippocamp can be gained; cerebral cortex is likewise quite
easy to stain. With the olfactory bulb the action is not constant
though fairly complete. The optic lobes and retina of birds and
large reptiles are more satisfactory than those of mammals. The
my el (spinal cord) of embryo birds (7 to 14 day chick best) is gen-
erally more satisfactory than that of mammals; in any case, fetal or
new-born animals should be employed. Difficult are satisfactory
impregnations of sympathetic ganglia, organs of special sense and
the intrinsic nerves of the viscera.

The important forms of the method are: — (a) the Slow Method;
mordantage in dichromate solutions of preferably increasing
strength, 2, 3, 5%, for 1 to 4 months depending on the temperature,
strength of solution, etc. (b) The Rapid Method; in this another
oxidizer, osmic acid, is combined with the dichromate with a reduc-
tion in the duration of the treatment to a few days. Combination of
(a) and (b) are sometimes serviceable; (c) Double (or triple) Im-
pregnations, obtained by repeating (b). Important modifications
are: (1) substitution of formalin for the osmic acid in (6), (2)
mercuric chlorid instead of silver nitrate (Cox's Method). The
Golgi methods have been widely applied and for details the individual
papers may be consulted [6].

§ 206. Golgi's Rapid Method. This is the most generally
serviceable of the different methods.

Directions jor use. Tissue of a (preferably) yoiing animal is
placed in a mixture of 4 parts of 3% potassium dichromate and 1
part of 1% osmic acid. The amount of the fluid should be at least
twenty times the bulk of the tissue and should be changed as soon
as it grows turbid or loses the strong characteristic odor of the osmic
acid.

After the action has proceeded to the right degree (§ 207), rinse
the tissue in water for about 5 minutes and place for 15 minutes in

a ]/4C/c solution of silver nitrate, and then for 2 or more days in a
%% solution of silver nitrate, preferably keeping it in the dark.
Without washing, imbed rapidly in celloidin as follows:
(a) Dehydrate 2 to 3 hours in 95% alcohol, changed two or
three times; (b) place in thin celloidin for 20 minutes, in thick cel-
loidin for 20 to 30 minutes; (c) imbed in thick celloidin, on a block
of wood (best ; § 65a) ; (d) harden the mass in chloroform for 20 to
30 minutes, and (e) place the block in clarifier and cut, sections
being 50 to 100 ^ thick according to the nature of the tissue and the
character .of the impregnation.

(/) Place the sections in 95% alcohol for a few minutes; clear
in carbol-xylene and mount in balsam by placing the section on the
slide, absorbing the clearer thoroughly by means of tissue paper and
spreading over it thick xylene balsam. Do not cover. Later, when
the balsam has hardened somewhat, it may be melted by heat and
much of the superfluous balsam drained from the section and scraped
away with a knife, and a cover glass added if desired.

§ 207. Time oj hardening. From results of Cajal, van Gehuch-
ten, and others, and from general laboratory experience, the follow-
ing periods will probably be found approximately correct. In
general: The best results are to be obtained with kittens 3 to 20
days old, puppies 2 weeks old, rats 8 to 10 days, rabbits 8 days, (a)
For cerebral cortex (and hippocamp) : New-born kitten, 1 to 2 days;
kitten half grown (3 to 4 months), 3 to 4 days; new-born rabbit, 24
hours; rabbit one month old, 2 to 3 days; adult mice, 3 to 4 days.

(b) For spinal cord : Chick of 5 to 6 days' incubation, 24 hours;
chick, 14 to 15 days' incubation, 3 days; new-born kitten or puppy,

2 to 3 days. Frog tadpoles (large) 3 to 5 days.

(d) Sympathetic system: Chick of 14 to 18 days' incubation,

3 days.

(e) Retina : 1 to 3 days.

(/) Olfactory mucous membrane: 3-4-7 days.

§ 208. Intra vitam Methylene Blue. Methylene blue shares
with a number of coal tar dyes the power of staining during life
nerve cells and fibers and certain cytoplasmic granules. Like the
Golgi methods, "intra vitam" staining of nervous tissue is capricious;
applicable only to living or fresh tissue and depends upon unknown

substances or conditions that become changed after death. In the
reaction reduction of the methylene blue to its leucobase (colorless)
by the nervous tissues appears to play an important part. In general,
the technique involves: (1) Bringing a methylene blue solution of
sufficient strength in contact with the (essentially) living nervous
elements, (2) permitting it to remain a sufficient length of time for
the staining reaction, (3) exposing the tissue to the action of the
oxygen of the air until the stain is fully developed, — re-oxidation
of any leuco-methylene blue and satisfying the reducing reaction of
the tissue; then — (4) either examining at once or fixing the stain
in situ by its precipitation in an insoluble form for its preservation
(imbedding and sectioning) .

1. The methylene blue may be brought in contact with the
neurones by injection, (a) through the vascular system, — aorta if
the animal is small, artery supplying the part, if large; — (b) into
the body cavities ; (c) subcutaneously : or, by immersion of the organ
or part, or the entire animal if small (many invertebrates). Keep-
ing in mind the end result desired, the best method will suggest itself
in a particular case. Cajal cut parallel slits in the cerebral cortex
and inserted the methylene blue in powdered form or as a saturated
solution. In general, the solutions should be as dilute as possible;
of 1/15 to 1/4% strength in physiological salt solution, the more
direct the application the weaker. It is well to have on hand a 1%
stock solution in physiological salt solution and dilute it (with salt
solution) as desired. Of the methylene blue preparations that are
available, Ehrlich's or "B. X." are more generally used. A combi-
nation of injection and immersion is often advisable. In introduc-
ing the stain by injection, first remove the blood by washing out with
physiological salt solution or by bleeding and let the injection be
full, i. e., through the capillaries into the veins. If a mammal is be-
ing dealt with, salt solution and staining solution should be warmed
to body temperature (35 to 38° C.).

2. It is difficult to give any general rules as to the time the
methylene blue solution should remain in contact with the tissue
before exposure as it is best to determine it experimentally in each
case. The time should be shorter for warm blooded, longer for cold
blooded animals. If introduced by injection, 20 to 30 minutes for
a mammal, 2 to 12 hours for a cold blooded form, may be suggested;
the organ or part should then be removed, wet with the dilute stain
(perhaps 1 /15% strength) for another period of time, — ^ to 1 hour,

access of oxygen to the point desired being kept in mind. At inter-
vals, free hand sections should be examined under the microscope to
determine the state of the reaction. If the tissue is mammalian, it
should be kept protected from evaporation during this time and
warm, as in an incubator.

If the stain was applied by immersion, a shorter time suffices; —
up to 15 minutes or so with a subsequent exposure to the air of l/2
to 1 hour wet with the dilute solution.

Small aquatic animals may be immersed in very dilute solu-
tions (1/100 to 1/1,000%), the optimum strength and time of im-
mersion being experimentally determined.

In some instances (particularly parasitic worms) the tissue
hold the methylene blue in reduced form in spite of exposure to the
air and the color is only developed when placed in the fixer (below).
In any event it is better to fix the stain earlier than later.

4. Two methods are standard for preserving tissue stained
intra vitam with methylene blue, the first of these (Dogiel's) is suit-
able only for such tissue as may be exposed for study by teasing;
the second (Bethe's) may be used both for such preparations and
those which it is desired to imbed and section.

Dogiel's Method. Immerse the tissue in a saturated solution of
ammonium picrate (orange-yellow needles) for 2 to 24 hours according
to the size of the piece, using abundance of fluid. If maceration
occurs, Dogiel suggests addition of 1% of 1% osmic acid. Transfer
tissue to equal parts of the ammonium picrate solution and glycerin in
which the tissue may be preserved, teased, and mounted.

Bethe's Method. Immerse the tissue in a 5 to 10%aqueous solu-
tion of ammonium molybdate for 1 to 24 hours according to the size of
the piece, using abundance of the fluid. Trim the tissue as desired,
removing all unnecessary parts, dividing it into smaller pieces, etc.
Wash in distilled water, changed several times, for 1 to 3 hours.
Dehydrate rapidly in 95% and absolute alcohol, — 4 to 6 hours,
shortening the time if possible. Imbed rapidly in celloidin (§ 206)
which may be hardened in 67% alcohol and sections cut.

It may be advisable, particularly in summer work, to have the
water and alcohol specially cooled to prevent dissolving of the stain
in the alcohol. Tissue already fixed in the ammonium picrate may
be refixed in the ammonium molybdate solution. Indeed, this
double fixation is recommended by Bethe as particularly suitable for
invertebrate material.

The paraffin method is not advisable if the above method suf-
fices. If it is desired, however, after the dehydration, clear thor-
oughly with oil of cloves followed by xylene and infiltrate in paraffin
(§ 52). In treating the sections, avoid alcohol as much as possible.

-Section or in toto staining may be applied, preferably carmine
(not alkaline or acid formulae) .

This important method has been elaborated largely through the
work of Bethe, Cajal, Dogiel, Huber, Retzius and others, for which
consult [6].

The method may be used for the staining of Neurofibrillae ; for
its applications for this purpose, consult the special articles.

§ 209. Neuroglia Stain. Tissue is fixed for 24 hours in copper
dichromate-sublimate-acetic (1/5%) mixture (§ 16) and subsequently
mordanted 3 or 4 days in 2.5% copper dichromate. Imbed in paraffin,
Sections (5 to 10^) are fastened to the slide and the Benda stain is
used, as follows: — (1) 4% ferric alum for 24 hours; rinse well in dis-
tilled water and (2) place for 24 hours in a dilute solution (amber
yellow) of sodium sulphalizarinate (concr. sol'n. in 70% alcohol
added to distilled water). Rinse in distilled water, blot with absorb-
ent paper, and stain (3) in a 1/10% aq. sol'n. of toluidine blue,
heating it until it steams, cooling and staining 15 minutes. (4) Rinse
with distilled water and treat for a few seconds with acid alcohol (70%
alcohol, 100 c. c.; concr. Hcl, 6 drops). (5) Blot with absorbent
paper and dehydrate rapidly with 95% and absolute alcohol. (6)
Differentiate carefully with creosote, to the right degree; (7) blot
with absorbent paper, and rinse in several changes of xylene. Mount
in balsam. Neuroglia fibers, a dark blue, neuroglia 'cells' a light
blue, axis cylinder and myelinic sheath red to brown, nuclei dark blue,
etc.

This method may be used with fresh or formalin material, human
or animal. (Kingery).

THE BLOOD.

Special methods in the examination of the blood include (1)
Examining fresh; (2) Technic of staining blood films; (3) Deter-
mination of the number of red and white corpuscles per cubic milli-
meter; (4) differential counting of the white corpuscles; (5) De-
termination of the relative amount of hemoglobin; (6) Spectroscopic
examination of blood (hemoglobin), etc. (1) and (2) are briefly
given here; for (6) see [17].

§ 210. Examining fresh. This consists in covering a drop
on a slide, and immediately sealing the cover-glass to prevent evapo-
ration, observing the following cautions: (1) The drop of blood
(from the finger or the lobe of the ear) should flow freely and not
be obtained by pressure. The drop should be a medium-sized one,
which will spread out in an even, thin layer under the cover. (2)
The drop should be received upon a cover or slide, covered, and
sealed at once with castor-oil.

Examination of fresh blood may be used in clinical examination
for the detection of some abnormal conditions, and it is of value in
the rough diagnosis of many others.

§ 211. Stained preparation of blood, (a) Preparing the blood
film. This may be best done in one of two ways: (1) The edge
of a slide is first drawn through a drop of fresh blood and then moved
quickly across the surface of a clean cover-glass or slide, in this
way spreading the blood in a thin, even layer upon the glass. Success
depends upon getting the right amount of blood upon the edge of the
slide and the quick, even movement by which it is spread upon the
cover-glass or slide. Preparing the film on a slide is simpler and to be
preferred if a differential count of the leucocytes is to be made. A
second, possibly better, method is the following:

(2) Have ready twro thin clear cover-glasses (or slides) and
obtain a drop of fresh blood. Take one of the covers in the forceps,
touch it to the drop of blood and place it upon the second cover-
glass eccentrically, with one edge projecting slightly. Slip the two
covers apart in the plane of their surfaces and dry them quickly by
waving them in the air or by passing them rapidly over the tip of
a flame. The lower cover-glass will have the better film.

(b) Fixing the hemoglobin with (a) ether-alcohol or heat, or (b)
at the time of staining (methyl alcohol) (§ 214).

§ 212. Fixing the film. When the blood films on the covers
are dry, place them in the fluid for J/2 to 1 or several hours. Let
them fix for a longer rather than a shorter time, as the quality of
the stain (with triacid mixture) will be improved. After they have
fixed a sufficient time remove and again dry them in the air. They
may now be stained, immediately or at convenience.

§ 213. Staining unfixed films. Eosin-Methylene Blue stains
(below). Fixed films may be stained writh hematoxylin and eosin as
well as with other stains. If the film is on the slide balsam and a

cover-glass are unnecessary if it is to be examined with the oil immer-
sion objective.

§ 214. Eosin-Methylene Blue. The most of the formulas are
made on the principle of neutral stains, — eosinates of methylene blue
dissolved in methyl alcohol, the staining solution being diluted with
water during the staining (§ 85, b). As polychrome methylene blue
is generally used, the range of selectivity is increased by the presence
of methylene azure.

(a) Nochts-Hastings Stain. (§ 130). Stain blood smears 1
minute with the undiluted stain, then dilute with distilled water
until a metallic film begins to appear and the diluted stain appears
reddish at the edge. Stain 5 minutes more, rinse quickly with dis-
tilled water, absorb excess with absorbent paper, dry in the air;
when dry mount in balsam.

(b) Wright's stain (§ 129) is of similar composition and the
staining process is carried out in the same manner. Stain blood
smears 1 minute with the undiluted stain ; dilute drop by drop with an
equal volume of water and stain for 3 minutes; rinse, dry, etc., as
above.

(c) Jenner's stain (§ 128) is a simpler stain, and easier to use.
The simple (i. e., not polychrome) methylene blue is used, and
the differentiation of the stain is secured in the washing out. Stain
blood films 3 minutes or more, rinse a short time with distilled water
(until the best portions of the film are pink) . Absorb excess of water
with absorbent paper and dry in the air. When dry, mount in
balsam.

§ 215. It is sometimes advantageous, — as for class work and with
non-mammalian blood, — to handle blood in bulk. The following
method has been used with good results. Fix for 1 to 6 hours by
having the blood drop into a vial of 1% osmic acid. The blood-cells
are allowed to settle and the supernatant fluid removed with a pipette.
By this method the blood is passed through, successively, — 2 or 3
changes of distilled water, 50%, 67% alcohols, paracarmine, 67%,
82%, 95% and absolute alcohols, xylene, to thin xylene balsam (§ 161)
in which the blood is stored. By gently agitating, the corpuscles are
evenly distributed and a drop of the balsam mounted contains
numerous blood cells. This is an excellent method for preparing
isolated epithelial and muscle cells for class use. Such material may
be kept for years and is always ready to use.

FINE INJECTION.

For the purpose of examining microscopically the finer arteries
and veins and the capillaries in a tissue, and their relation to the
other parts, it is necessary to fill them with some colored injection
mass, or otherwise stain or color them. Numerous injection masses
are in use; the following meet the general needs. For injection
fluids for special purposes, consult the literature [6].

§ 216. Carmine gelatin mass. Formula: Dry gelatin, 75
grams; carmine (No. 40), 10 grams; water, 90 c. c.; ammonia, 10
c. c.; acetic acid, q. s.\ chloral hydrate, 10 grams.

Soak the gelatin in water until it is soft; pour off the superflu-
ous water and melt it (in an agate or porcelain dish) over a water
bath. Grind the carmine to a paste with water; add all the am-
monia and water; filter, warm to 80° or 90° C., and add to the warm
gelatin. Then add slowly the acetic acid diluted with an equal vol-
ume of water, while constantly stirring the mass, until the mass
smells very slightly of the acid. Filter through fine flannel. If the
mass is acid, the chloral hydrate may be safely added (as a preserva-
tive) ; if any ammonia is present it will decompose it forming chloro-
form, and a granular precipitate. If too much acid is added, the
gelatin will not set.

§ 217. Berlin blue injection mass. Formula: Dry gelatin,
75 grams; saturated aqueous solution of Berlin blue, 150 c. c.; chloral
hydrate, 10 grams. Prepare the gelatin in the manner given above
(§ 216); warm the Berlin blue solution (to 80° or 90° C.), and add
it to the hot gelatin. Heat the mixture for 10 minutes or more,
stirring it occasionally, and filter it through fine flannel and add the
chloral hydrate.

§ 218. For securing the best results in injecting the following
conditions should be observed: (1) A young but nearly mature,
lean animal is to be preferred. (2) Kill the animal with an anes-
thetic (chloroform) and leave it in the anesthetic at least half an
hour before beginning the injection; do not, however, wait until
rigor mortis sets in. (3) Inject only the part desired, tying all an-
astomosing vessels and all vessels to other parts. Inject into the
artery of the part, leaving the vein open until nearly pure injection
mass escapes, then tie it and continue the injection until the part
feels hard and is the color of the injection mass. (4) When the
injection is finished cool the part injected by means of cold water,
ice, or snow.

(5) Harden the injected tissue 1 or 2 days in 50% alcohol, 2
or 3 days in 67% and 82% alcohols. The acidity of the alcohols
should be insured by adding to the 50% alcohol a few drops of acetic
acid. The tissue may be stored in 82% alcohol until ready for sec-
tioning. Formalin (10%) may also be used as the fixer and preserving
fluid. For sectioning the celloidin method is usually preferable.

§ 219. Silvering blood vessels. Silver nitrate may be used
for coloring blood vessels, and thus differentiating them. See § 222.

§ 220. Dense Masses, such as do not pass through the capil-
lary network are useful in giving double injections, the veins and
capillaries one color, arteries another. Two such may be mentioned:
(a) Lampblack Gelatin Mass; (b) Ultramarine Gelatin Mass (Spal-
teholz.).

(a). Lampblack and gelatin mass in the proportion of about
1:12.

(b). A 10% gelatin mass to which is added ultramarine in
proportion of 30:100.

In injecting, inject through the artery with one of the gelatin
masses given in § § 216 — , and follow it up with one of the above
which will push the first mass through into the capillaries and veins.

SILVER NITRATE IMPREGNATIONS.

§ 221. The preparations stained by means of nitrate of silver
are prepared as follows; The fresh tissue is washed for a minute or
so in distilled water to remove from the surface all albuminous sub-
stance, and then transferred for 2 to 5 minutes or longer to a 1 to
Yf/o aqueous solution of silver nitrate and exposed to direct sunlight
until a light brown. When, by examination with the microscope, the
stain was found to be sufficient it was again rinsed in water and placed
in glycerin or alcohol. Employed in this manner with fresh tissue,
silver nitrate stains the cell cement, affording thus negative images of
the cells. If a membrane such as mesentery is to be silvered removed
from the body, it should first be cautiously stretched, as over a ring,
to avoid creases.

§ 222. Silvering Vascular Epithelium. In order that the
vascular epithelium of small arteries, veins, and capillaries should be
well demonstrated, silver nitrate solutions of J^ to //2% strength
must be injected into the vessels.

§ 223. Procedure. Connect a canula with the artery supply-
ing the alimentary canal (superior mesenteric) or the brain (caro-
tid) and inject distilled water until the water flows out of the re-
turning vein colorless. Then immediately inject the silver solution
until it runs from the vein. After a minute or two follow writh
distilled water or physiological salt solution. Place the intestines
and mesentery in water and expose them to the light until they become
slightly browned. Strips of the muscular coat of the intestines,
especially of the rabbit, will show capillaries well. Veins and arteries
side by side may be found in the mesentery. If the brain vessels are
injected one can get admirable preparations showing nuclei as well as
cell outline by staining in hematoxylin. Mount in glycerin, or, if
desired, dehydrate and mount in balsam. The tissue may be kept in
50% alcohol or in 50% glycerin for several months before mounting
if it is kept in the dark.

For large vessels and endocardial epithelium open the vessels
or the heart and silver as directed above for mesentery. It may be
necessary to make thin free-hand sections so that the preparation
will be thin enough for high powers.

HISTO-CHEMICAL METHODS.

There are special chemical substances which it is often desirable
to preserve and differentially stain. In most cases, the staining re-
actions are not specific enough to come under the category of micro-
chemical tests, the evidence gained being circumstantial or indirect,
the application of two or three different methods being sometimes
necessary for confirmation. Such methods may be spoken of as
histo-chemical rather than micro-chemical.

A. Fats. (Lipoids).

§ 224. Free fats and lipoids are soluble in ether, chloroform,
absolute alcohol, xylene, benzene, and essential oils. As these are
necessary for paraffin and celloidin imbedding methods, especially
the former, the satisfactory preservation of these substances pre-
sents some difficulties. The use of the freezing microtome is there-
fore particularly called for. See, however, § § 227, 228.

The stains applicable to the demonstration of fats are (a) stains
soluble in the fat solvents, e. g., Sudan III, and scarlet red; and (b)
such as depend upon the reduction of salts by the fats (e. g., osmic
acid and potassium dichromate) [29].

§ 225. Sudan III. Fix tissue in formalin or Miiller's fluid.
Cut free-hand sections, employ the freezing microtome method, or
isolation (§ § 35 — ). Rinse sections in 82% alcohol and transfer
sections to a strong solution of the stain in 82% alcohol;
leave several minutes covered from evaporation; rinse with 82%
alcohol and transfer to water. Mount in glycerin or glycerin jelly
(§ § 151, 155). Fat is stained red.

§ 226. Herxheimer's Stain. Preparation for staining is as above
(§ 225). Pass sections into 67% alcohol. Transfer to a strong solu-
tion of Scarlet Red in 67% alcohol rendered alkaline by 2% of sodium
hydroxid. Stain for several minutes; rinse with 67% alcohol and
transfer to water. Mount in glycerin or glycerin-jelly. Fat globules
stained red. It affords a more intense stain than Sudan III.

Indophenol (saturated solution in 67 or 82% alcohol) may be
used as a blue stain for fat in a similar manner.

§ 227. Osmic Acid. (§ § 30, 18, 19). Osmic acid is reduced
by the unsaturated fatty acids (e. g., oleic acid) which are blackened
by it. The saturated fats and fatty acids (e. g., stearic, palmitic,)
are not so blackened [1] but the black color subsequently appears
when the tissue is placed in alcohol [41]. The fat so oxidized and
impregnated with osmium (?) becomes less soluble in fat solvents
(§ 224) and may be retained in tissue imbedded in either paraffin or
celloidin. There is, however, a difference in fats; adipose tissue is
easily preserved, while some of the fat granules found in the organs
require the special precautions mentioned below.

Fix sections of tissue 2 to 3 mm. thick in Flemming's or Benda's
fluid for 2 days; dehydrate in 95% alcohol, and transfer to thin
celloidin, — and subsequently thick celloidin — made up with 95%
alcohol (not absolute). The sections may be stained in safranin,
quickly dehydrated and cleared with carbol-xylene and mounted in
balsam, either without a cover-glass, or in thick balsam melted by heat
and applied warm. Unless such precautions are taken, the solvent of
the balsam may in time dissolve out the granules of blackened fat.

Paraffin does not afford as good a preservation of the more labile
fat globules. Dehydrate in equal parts of 95% and absolute alcohol,
clear before the infiltration in carbol-xylene or chloroform. Paraffin
in the sections should be dissolved out by carbol-xylene in preference
to xylene (§ 143).

§ 228. Bichromate mordantage, with subsequent copper or
iron hematoxylin stain, appears to rest upon the power of the fat or
lipoid to reduce the dichromate and thus take on a mordantage which
gives the basis for the subsequent staining. So far, it has not been
possible by this technique to preserve, in paraffin or celloidin sec-
tions, the individual fat granules, but it is nevertheless a useful
method for the differentiation of lipoid-containing cells.

Fix tissue 2 days in Zenker's with but 1 /10 or 1 /5% of acetic acid
or Helly's fluid, mordant 4 days or longer in Muller's fluid at 35 to
38° C., wash in water, imbed in paraffin using chloroform as the clearer
(§54). Stain sections with the copper hematoxylin (§ 95). Lipoid-
containing cells, myelinic nerve fibers, erythrocytes, etc., a dark blue.
It should be also remembered that other structures may also be
stained by this technique.

The freezing microtome may be used with such tissue and stain,
as has been done by Benda and Fischler [6].

B. Glycogen.

Glycogen is soluble in aqueous media, and while it may be retained
by a short fixation in several fluids, the best preservative of it is 95%
alcohol (82%— absolute).

§ 229. The Iodine Method. [16]. Fix tissue in 95% or 82%
alcohol. Imbed in paraffin. Spread the sections using instead of the
water, the iodine stain for glycogen, which is made up as follows:
iodin, \^/2 grams; potassium iodid, 3 grams; sodium chlorid, \}/<i
grams; distilled water, 300 c. c. Spread sections may be stained or
restained by immersing in the iodin solution which will color the
glycogen a mahogany red. For very soluble glycogen, 50% alcohol
may be employed instead of the water in making up the stain. In
mounting, dissolve the paraffin with xylene, drain, place on the
preparation melted yellow vaseline, cover, seal with shellac or balsam.

§ 230. Best's Method. [5]. While rather complicated, this
is generally recognized as the best method for the demonstration of
small quantities of glycogen, especially when it is desired to see the
relation of the granules to the protoplasm.

Fix tissue in 95% alcohol; imbed (preferably) in celloidin (§67);
if paraffin is used, after dehydration (§ 51) place the pieces of tissue
in pure acetone for 15 minutes, xylene 20 minutes, xylene paraffin
1 hour, pure paraffin (§ 52) 1 hour. Stain sections in Delafield's
hematoxylin (§ 91) strongly, rinse and differentiate (if necessary)
and stain in the following special carmine stain which had been

' 87

previously prepared, (a). Carmine, 1 gram; ammonium chlorid,
2 grams; lithium carbonate, 0.5 grams; distilled water, 50 c. c.
Bring the mixture to a boil. When it is cool, add 20 c. c. 10% am-
monia. Preserve the solution in the dark; after 2 to 3 days it is
ready for use and retains its staining quality for a few weeks only,
(b) When ready to stain, filter the above solution (a), and add to
2 parts of the stain, 3 parts of 10% ammonia solution, and 6 parts
of methyl alcohol.

Stain sections 1 hour, differentiate in a mixture of 2 parts of
methyl alcohol, 4, parts of absolute alcohol, 5 parts of water; rinse
with 82% alcohol, dehydrate, clear and mount in balsam. Glycogen
stained an intense red.

In working with glycogen it is sometimes necessary to apply as
a control the digestion of the glycogen in one or more sections or a
part thereof, by means of saliva.

C. Amyloid.

Amyloid, a form of connective tissue degeneration, resembles
glycogen in some of its physical (not chemical) properties and stain-
ing reactions. Two methods for its demonstration may be men-
tioned :

§ 231. Iodine Method. Practically any fixer may be used
(Zenker's fluid). Paraffin, celloidin, or (better) frozen sections
may be used. Stain sections with the iodine solution (§ 229) for
several minutes; rinse in distilled water and transfer to glycerin or
glycerin-jelly (§ 151, 155) in which they may be mounted. Seal the
preparations (§ 162 — ). The amyloid a reddish-brown; the stain,
however, will fade in the course of a few months.

§ 232. Gentian Violet, among other anilin stains, colors amyloid
differentially (metachromasia). Stain paraffin sections for several
minutes in a 1% solution of the stain; rinse and differentiate in 1%
acetic acid; wash thoroughly with distilled water, and (a) mount in
glycerin- jelly, or (b) dry in the air, treat with xylene and mount in
balsam. In the latter procedure, the staining may be applied before
removing the paraffin.

D. Mucus.

Mucous substances (mucins, mucinoids) possess acid properties
combining with alkalis and bases (heavy metals), such combinations
swelling up or dissolving in water. Acetic acid, alcohol and picric
acid also precipitate these substances. For the fixation of mucus,

most fixers may be used, — Mercuric chlorid, Zenker's fluid, Flem-
ming's fluid or Hermann's fluid. If, however, it is desired to pre-
serve the mucus granules (mucinogen?) in the cell-body, alcoholic
or picric acid fixers are usually required, and the tissue at no time
subsequently should be placed in water or aqueous solutions. Be-
cause of its acid character mucus stains wTith basic stains, the first
two methods given below being very selective. If cell granules are
desired, remember the caution as to the avoidance of water.

§ 233. Mucicarmine. (§ 103). Fix as recommended above.
Stain paraffin sections 1 to 24 hours; wash in water, dehydrate,
clear and mount in balsam. It may be used diluted with 50% or
67% alcohol.

§ 234. Muchematein. (§ 93). Employ in the same manner as
mucicarmine. The alcoholic formula is indicated if cell granules
are to be preserved.

Staining of the nuclei with hematoxylin (§ 88, 89) or carmine (§ 98)
or elastin stain (§ 125) may be previously given; a picro-fuchsin
stain may follow it. Excess of alum (aluminium salts) or presence
of acid must be avoided, hence wash thoroughly with water if there
has been previous staining.

§ 235. Basic anilin stains usually give sharp, often differen-
tial stains (metachromasia) for mucus. Methylene blue, toluidin
blue, or gentian violet may be recommended. Safranin after Flem-
ming's fluid fixation often gives a delicate stain for mucus.

E. Micro-chemical Tests.

Micro-chemical tests for inorganic elements must occasionally
be employed, particularly in cytological work. The methods have
been largely elaborated by Macallum to whose papers references are
given.

§ 236. Iron. (a). Berlin Blue Reaction. Alcohol fixation and
the freezing microtome should be employed. Rinse sections in
distilled water, 2% potassium ferrocyanate, 4 to 5 hours, 1% Hcl
alcohol several hours, rinse in water, dehydrate, clear with clove oil;
balsam. A carmine stain may be given, — before or afterwards.

(b) Ammonium Sulphide Reaction. Place sections in am-
monium sulphide solution (freshly prepared) for 5 to 20 minutes;
(dark green color); rinse quickly with water, dehydrate, clear in
clove oil, mount in balsam.

Free iron may also be demonstrated by the use of a pure aque-
ous solution of hematoxylin (dark-blue color); other metals, how-
ever, also unite with hematoxylin (e. g., calcium).

§ 237. Masked Iron does not give the reaction with pure aqueous
hematoxylin [31, 34]. Macallum applies the ammonium sulphide at
higher temperature (60° C.) and for several days.

§ 238. Phosphorus. See Macallums methods [31].

§ 239. Potassium. See Macallum's method [33].

§ 240. Chlorides. See Macallum's method [32]. Cautions!

§ 241. Calcium stains strongly with aqueous hematoxylin and
may often be so demonstrated. To identify the calcium and dis-
tinguish from iron, usual chemical tests may be applied [31, 28].

REFERENCES.

1. Altmann, R. Die Elementar Organismen. 2te. Aufl. Veit & Co.,

Leipzig, 1894.

2. Bell, E. T. The Staining of Fats in Epithelium and Muscle Fibers. Anat.

Record. Vol. IV. No. 5, 1910.

2. Benda, C. Die Mitochondria. Ergeb. d. Anat. u. Entwicklungsges.
Vol. XII, pp. 743-781. 1901.

A. Berg, W. Die Fehlergrosse bei den histologischen Methoden. Berlin,
1908. Also: Arch. f. mikr. Anat., Vol. LXII, 1903; Vol. LXV, 1905.

5. Best, F. Ueber Karminfarbung des Glykogens und der Kerne. Zeitschr.

f. wiss. Mikr. Vol. XXIII, 1906.

6. Ehrlich, P., Krause, R., Mosso, M., Rosin, H., & Weigert, P.; Editors.

Enzyklopaedie der Mikroskopischen Technik. Urban & Schwarzen-
berg. Berlin, 1910.

7. Fischer, A. Fixierung, Farbung und Bau des Protoplasms. Kritische

Untersuchungen iiber Technik u. Theorie in den neuen Zellforschung.
Jena, 1899.

8. Gage, S. H. & Susanna P. Staining and permanent Preservation of his-

tological Elements isolated by means of Caustic Potash and Nitric Acid
Proc. Amer. Soc. Micr. Vol. XI, 1889. pp. 34-45.

9. Gage, S. H. Notes on albuminizing slides for the more certain fixation of

serial celloidin sections. Proc. Amer. Soc. Micr. Vol. XIII, 1891, pp.
79-84.

10. Gage, S. H. Methods of Decalcific'ation in which the Structural Elements

are preserved. Proc. Amer. Soc. Micr. Vol. XIV. 1895. pp. 121-124.

11. Gage, S. H. Ah aqueous Solution of Hematoxylin that does not readily

deteriorate. Proc. Amer. Soc. Micr. Vol. XIV. 1892. pp. 125-127.

12. Gage, S. H. Agnes Claypole's method of fixing paraffin sections to the

slide. The Observer, Vol. VI. 1895. pp. 60-61. See also: Agnes
Claypole; Amer. Micr. Soc. Trans. 1894. pp. 66 & 127.

13. Gage, S. H. On the Use of Formalin as a dissociating Medium. The

Micr. Bull. & Sci. News. Vol. XII, 1895. pp. 4-5.

14. Gage, S. H. Improvements in oil-sectioning with celloidin. Proc. Amer.

Soc. Micr. Vol. XVII, 1895. pp. 361-370.

15. Gage, S. H. Notes on the Isolation of the tissue Elements. Proc. Micr.

Soc. Vol. XIX, 1897. pp. 179-181.

16. Gage, S. H. Permanent Preparations of Tissues and Organs to show

Glycogen. lodin stain; Mounting in yellow Vaseline and Xylene
balsam. Proc. Amer. Micr. Soc. 1906. Vol. XXVIII, pp. 203-205.
See also: Amer. Journ. Anat. Vol. IV, No. 2, 1904.

17. Gage, S. H. The Microscope. Comstock Publ. Co., Ithaca, N. Y. 10th

Ed. 1908.

18. v. Gehuchten, A. Anatomic du Systeme Nerveux de I'Homme. 1906.

4me. Ed. p. 340.

19. v. Guyer, M. F. Animal Micrology. Univ. of Chicago Press. Chicago,

1906.

20. Hall, W. J. & Herxheimer, G. Methods of Morbid Histology and Chemical

Pathology. Edinburgh & London, 1905.

21. Hardesty, Irving. Neurological Technique. Univ. of Chicago- Press.

Chicago, 1902.

21a. Hardesty, Irving. Laboratory Directions for Histology. Blakiston.
1908.

22. Hastings, T. W. A modified Nochts Stain. The Johns Hopkins Hos-

pital Bull. Vol. XV. 1904. pp. 122-123. See also: Journ. Exper.
Med. Vol. Ill, 1905. p. 265.

23. Heidenhain, M. Ueber Kern und Protoplasma. Leipzig, W. Engel-

mann. 1892.

24. Heidenhain, M. Ueber die zweckmassige Verwendung des Congo und

anderer Amidoazokorper sowie iiber neue Neutralfarben. Zeitschr. f.
wiss. Mikr. Bd. XX. 1903. p. 179.

25. Heidenhain, M. Ueber Vanadiumhaematoxylin, Picro-blauschwarz und

Kongo-Korynth. Zeitsch. f. wiss. Mikr. Vol. XXV. 1908. p. 397

26. Heidenhain, M. Plasma und Zelle. Erste Abt.: Allg. Anatomie des

lebendigen Masses. Jena. 1907.

27. Jenner, A., A new Preparation for Rapidly Fixing and Staining Blood.

Lancet, 1899, 1. p.'370.

28. v. Kahlden-Giercke. Technik der histolgischen Untersuchungen. 8.

Ed. Fischer, Jena, 1909.

29. Faure-Fremiet, A. Mayer, G. Schaeffer. Sur la microchimie des corps

gras; application a 1'etude des mitochon dries. Arch. d'Anat. mi-
croscopique. Vol. XII, Fasc. L, 1910. pp. 19-102.

30. Lee, A. B. The Microtomist's Vademecum. 7 Ed. Blakiston, Phila.

1913.

31. Macallum, A. B. Die Methode und Ergebnisse der Microchemie in die

biologische" Forschung. Ergeb. d. Physiol. Vol. VII, pp. 552-652.
1908.

32. Macallum, A. B. On the Nature of the Silver Reaction in Animal and

Vegetable Tissues. Proc. Roy. Soc. Vol. LXXVI. 1905. p. 217.

33. Macallum, A. B. On the Distribution of Potassium in Animal and Vegeta-

ble Cells. Journ. Physiol. Vol. XXXII. p. 95. 1905.

34. Macallum, A. B. On a new Method of distinguishing between organic

and inorganic Iron. Journ. Physiol. Vol. XXII. 1897. p. 92. Also:
On the Demonstration of the Presence of Iron in Chromatin by Micro-
chemical Methods. Proc. Roy. Soc. Vol. L, p. 277. 1891.

35. Mallory & Wright. Pathological Technique. 5th. Ed. 1911. W. B.

Saunders & Co., Philadelphia and London.

36. Mann, Gustav. Physiological Histology. Oxford. 1903.

37. Moore, V. A. A Note on the Use of Anise Oil in Histological Methods

with special Reference to its Value in Cutting Serial Sections on the
Freezing Microtome. Amer. Month. Micr. Journ. Vol. XV. 1894.
p. 373.

38. Myers, B. D. The Chiasma of the Toad (Bufo lent.) and of some other

Vertebrates. Zeitschr. f. Morphol. & Anthropol. Vol. III. 1900.
pp. 183-207.

29. Pappenheim, M. Grundriss mikroskopischer Farbetechnik. Berlin. 1901.

40. Sheldon, R. E. Paraffin-Weigert methods for the staining of nervous tis-

sue, with some new modifications. Folia Neuro-biologica. Vol. VIII.
No. 1, 1914. pp. 1-28.

41. Starke, J. Ueber Fettgranula und eine neue Eigenschaft des Osmium

tetraoxydes. Arch. f. Anat. u. Physiol. Physiol. Abt. 1895. p. 70.

42. Streeter, G. L. Ueber die Verwendung der Paraffineinbettung bei Mark-

scheidenfarbung. Arch. f. mikr. Anat. Bd. LXII. 1903. p. 734.

43. Strong, O. S. Notes on the Technique of Weigert's Method for Staining

Medullated Nerve Fibers. Journ. Comp. Neurol. Vol. XIII. 1903.
p. 291. Also: Ibid. Vol. XVI. 1906.

44. Verhoeff, F. H. An improved differential Elastic Tissue Stain. Journ.

Amer. Med. Ass'n. Vol. LVI, No. 18, pp. 1326-1327. 1911.

45. Wright, J. H. A rapid Method for the differential Staining of Blood

Films and Malarial Parasites. Journ. Med. Research. Vol. VII.
1902. p. 138.

46. Wright, J. H. Revised Directions for making and using the Wright's blood-

stain. Journ. Amer. Med. Ass'n. Vol. LV. pp. 1979. Dec. 3, 1910.

INDEX

Acetic acid, picric, formol, 12
Acetic acid, 5, 7

Acid hematoxylin (Delafield's), 39
Acid, formic, 15, 69

fuchsin, 44

hydrochloric, 36, 63

nitric, 15, 1 8, 63, 66

osmic, 15, 18,69,73,75,85

sulphuric, 18
Acid stains, 35
Acid violet, 45
Action of fixer, 7, 8
Adjective staining, 35
Albumin fixative, 49
Alcohol, absolute, 95%, 82%, 67%,

50%, ii, 14,22,50,51
Alcohol-acetic, 14
Alcohol-acetp-chloroform, 14
Alcohol-acetic-formol, 14
Alcohols as fixers, 14, 60, 70, 71
Alcohol, dehydration in, 22, 27, 51, 54
Alcohol, grades, preparation of, 1 1
Alum cochineal, 41
Alum differentiator, 36, 70
AlkaUne methylene blue, 42
Ammonium molybdate, 78
Ammonium pi crate, 78
Ammonium sulphid (iron test) , 88
Amyloid, stains for, 87
Anilin blue, Mallory's connective tissue

s£ain, 44

Anise-seed oil, 33
Babes' safranih, 42
Balsam, Canada, 54, 55
Balsam, camsal, 55

Damar, 52

Euparal, 55
Basic stains, 35, 88
Benda's fluid; see Flemmings fluid
Benda's stain for neuroglia, 79
Bensley's neutral gentian, 46
Berlin blue, 82

Berlin blue reaction (iron), 88
Best's method for glycogen, 86
Bleaching; peroxid, perhydrol; see

Hermann's fluid
Bleu de Lyon, stain, 45
Blocks, wooden, for imbedding, 28
Blood films, fixing and staining, 80, 81
Blood methods for, 79
Blood, preserving in bulk, 81
Blood vessels, silvering, 83
Bone, 63, 65
Borax carmine, 40

Bouin's fluid, see picro-aceto-formol
Boxes, paper, for imbedding, 24, 29
Cajal, methods for neurofibrillae, 71

Calcium, tests for, 89

Camsal balsam, 55

Carbol-xylene, 55

Carmalum, 40

Carminates, carmine, stains, 40, 86

Carmine gelatin mass, 82

Carminic acid, 40

Carnoy's fluids, 14, 70, 79

Castor-xylene, 29, 30

Caustic potash, dissociator, 19, 67

Cedar wood oil, clearer, 23

Cell, methods for, 59

Celloidin, staining, of 37

Celloidin method, 20, 26-,

Celloidin, solutions, 2%, 6%, 12%, 27,

K%,34,49

removal of , 48
Chick blastoderm, fixing; see nitric

acid

Chloral hematoxylin, 88
Chlorides, test for, 89
Chloroform, clearer, 23, 69

fixer, 14

hardener, 29
Chromatin, stains for, 59
Chromo-aceto-osmic; see Flemming's

fluid

Chromo-nitric; see Perenyi's fluid
Clarifier, clarifying, 29, 30
Class, isolated tissue for, 81
Cleaning mixture for glass, 58
Cleaning slides, cover-glasses, glass-
ware, 57, 58

Clearers, clearing, 22, 23, 55
Cochineal, alum, 41
Collagen, staining, 61
Collodion; see celloidin
Congelation mass; see freezing
Congo red, 43, 69
Connective tissue, 61
Copper dichromate-sublimate -acetic,

12,61,72,79

Copper hematoxylin, 40, 73, 86
Cover-glasses', 57, 58
Crystallization of paraffin mass, 25
Cytoplasm, fixation, 60
Cytoplasm, granules, 60, 6 1
Damar balsam, mounting medium, 52
Decalcification, 63
Dehydration, 22, 27, 54
Delafield's hematoxylin, 39, 69
Dichromate as fixer, 8, 1 6, 72, 73, 74, 86
Dichromate-acetic, 12
Differentiation, 36, 73 '
Digestion method (reticular tissue), 62
Dishes, for paraffin imbedding, 23

Dissociation, dissociators, 6, 16-
Double staining, double stains, 37
Dry mounting, 52
Ehrlich's acid hematoxylin, 39
Ehrlich-Biondi-Heidenhain triple stain,

Elastic fibers (elastin), stains for, 45, 62
Eosin, 43

Eosin-methylene blue, 42
Eosinate of methylene blue, 46
Epithelium, silvering, 83

isolation (dissociation) of, 17, 1 8
Erlicki's fluid, 15
Erythrosin, 43
Ether-alcohol, 27
Euparal, 55

Fastening sections to slide, 47-
Fat, fixation of, 72, 84

freezing microtome, use, 84

stains for, 84
Fixation, 7, 8
Fixation by injection, 9
Fixation, rules for, 9
Fixers, action of, 7

formulae for, 11-16

list of, 8
Fixing, 6, 7

Flemming's fluid, 13, 74, 85
Flemming's triple stain, 43
Formaldehyde, formalin, 14, 15

Dissociator, 18, 68
Formol-dichromate, 12
Free- (hand) sections, 47, 48
Freezing method for sectioning, 19, 33,

48,84

Fresh tissue, examination of, 6
Fuchsin acid, 44
Gage's dissociators, 18
Gage's hematoxylin, 38
Gage's method for glycogen, 86
Gelatin masses, 82
Gentian violet, 41, 43, 46, 87, 88
Gilson's fluid; see mer euro-nitric
Glycerin, 53, 55
Glycerin- jelly, 54, 56
Glycogen, fixation, staining, 86
Gold chlorid, methods, 69
Golgi methods, 74
Gram's solution; see iodine, 45
Granules, cytoplasmic, 60

secretion, 61
Ground sections, 64
Gum arabic, 33
Hsemalum, Mayer's, 39
Hair, dissociation of, 18, 19
Hardening of tissues, 10

celloidin mass, 29
Hasting's stain, 46, 81
Heidenhain's iron hematoxylin, 39
Held's method (nerve cells) , 70

Kelly's fluid, 12
Hemateates, hematein, 38
Hematoxylin stains, formulae, 38, 39, 40

stock solution, 38
Hermann's fluid, 13
Herxheimer's stain, 85
Horn, dissociation of, 18, 19
Hydrochloric acid decalcifier, 63
Hydrochloric acid carmine, 41
Imbedding, celloidin, methods, 20, 26

paraffin, 20, 22
Impregnations, 36
Indifferent fluids, 16
Indophenol, 85
Infiltration, (imbedding methods), 22,

26, 33

Injection, fine, dense, masses, 82, 83
In toto staining, 35, 41 , 50
Intra vitam methylene blue, 76
lodin, ii, 45, 86, 87
Iron hematoxylin, Heidenhain's, 39
Iron, tests for, 88, 89
Isolation ,16-, 66, 68, 69
Jenner's stain, 46, 8 1
Labeling slides, 56
Lampblack, gelatin mass, 83
Light green, 45
Lipoids ; see fats
Lipoids; foot note, 7

demonstration, 84
Lugol's solution; see iodine, 45
Lyon's blue, stain; see Bleu de Lyon,

Maceration, 16

Mallory's connective tissue stain, 44
Marchi methods (degenerating nerve

fibers), 73

Mayer: see carmalum, haemalum
mucicarmine, Muchematein, par-
acarmine

Mercuric chlorid, 1 1
Mercuro-nitric, 13
Methylene blue, 42, 70, 76
Methyl green, 42
Micro-chemical tests, 58
Microtome knife, care of, 58
Mitochondria, fixation and staining, 60,

Mordantage, 35
Mounting media, methods, 6, 52-

cells, 53, 54
Muchematein, 39, 88
Mucicarmine, 41, 88
Mucus, 87
Miiller's fluid, 15

dissociator, 17
Muscle, isolation of, 66
Muscle, methods for, 66-
Myelin, myelinic nerve fibers, 69
Nail, dissociation of, 18, 19

Nerve cells, isolation, staining, 68
Nerve fibers, isolation, degenerated,

69,73

Nervous system, methods, 68-
Neurofibrillae, 71
Neuroglia, stain for, 79
Neutral gentian, 46
Neutral stains, 37, 35, 36
Nissl's methylene blue, nethod, 76
Nitric acid, decalcifier, dissociator,

fixer, 14, 1 6, 63
Nochts-Hastings stain, 46, 81
Normal, salt solution; see physiological

salt solution
Orange-fuchsin acid, 62
Orange G., 44
Orcein, 45, 62

Orth's fluid; see formol-dichromate
Osmic acid, dissociator, fixer, 15, 18, 69,

73, 75

Osmium tetroxid ; see osmic acid
Pal's method, 73
Paracarmine, 41

Paraffin, grades, method, 20, 21-, 23
Paraffin sections, handling of, 48
Per enyi's fluid, 14
Peroxid, as bleacher; see Hermann's

fluid

Phosphorus, 89

Physiological salt solutions, 16
Picric acid, stain, 44
Picro-aceto-formol, 12
Picro-fuchsin, 44, 62
Picro-nitric, 14
Platino-aceto-osmic : see Hermann's

fluid

Potassium dichromate, fixer, 16
Potassium hydroxid dissociator, 19, 67
Potassium, tests for, 89
Progressive staining, 35
Pyridin; see Cajal's silver methods, 71
Pyroxylin; see celloidin
Quadruple stains, 89
Radium, to prevent electric sections, 26
Regressive staining, 35
Resorcin fuchsin, 45, 62
Reticular tissue, 62
Safranin, 42, 85
Scarlet red, 85
Sealing preparations, 55

Secretion granules, 61
Sectioning, methods of, 6, 19-
Section staining, 35
Serial sectioning in celloidin, 32
Shellac, cells, rings, 54

sealing, 55, 56
Silvering blood vessels, 83
Silver nitrate, 83, 71, 74
Slides, cleaning, 57
Soluble cotton ; see celloidin
vStaining, forms of, rules of, 35
Staining, isolated cells, sections, etc.,

18, 51, 53, 66, 67
Stains, 35-, 38
Stock solutions, 1 1
Storing of tissues, 10
Substantive staining, 35
Sudan III, 85

Sulphalizarinate of sodium, 61 , 79
Sulphuric acid dissociator, 18
Tellesnicky's fluid; see dichromate

acetic

Tigroid substance, 70
Tissues, fixing, 6, 7 -

hardening, storing, 10
Tissue paper, for handling sections, 31
Toluene, as clearer, 22, 21
Toluidin blue, 42
Tooth, 63, 64

Transferring celloidin sections, 31
Triple staining, 37
Triple stain; Ehrlich-Biondi, 42

Flemming's, 43

Ultramarine, gelatin mass, 83
van Gehuchten's, 74
Vascular epithelium, silvering, 83
Verhceff's elastin stain, 45
von Luko's fluid; see alcohol-aceto-

formol

Weigert methods, for nerve, 71
Weigert's copper hematoxylin, 40, 71
White connective tissue fibers, 61
Wooden blocks, for celloidin imbed-
ding, 28, 30
Wright's stain, 46, 81
Xylene, 23, 50
Xylene, as clearer, 22, 23
Xylol; see Xylene, 22 footnote
Zenker's fluid, 1 1
Zenker-formol; see Helly's fluid, 12

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