T EC H N I Q U ES OF

HISTO- AND CYTOCHEMISTRY

G "/J

TECHNIQUES OF

HISTO- AND CYTOCHEMISTRY

A Manual of Morphological and Quantitative
Micromethods for Inorganic, Organic and
Enzyme Constituents in Biological Materials

By DAVID CLICK, Ph.D.

ASSOCIATE PROFESSOR OF PHYSIOLOGICAL CHEMISTRY
THE MEDICAL SCHOOL, UNIVERSITY OF MINNESOTA

With a Foreword by Robert R. RENSLEY, M. B., D. Sc.

PROFESSOR EMERITUS OF ANATOMY, UNIVERSITY OF
CHICAGO

1949

INTERSCIENCE PUBLISHERS, INC., NEW YORK
INTERSCIENCE PUBLISHERS LTD., LONDON

Copyright, 1949, by
Interscience Publishers, Inc.

All Rights Reserved

THIS BOOK or any PART THEREOF MUST NOT
BE REPRODUCED IN ANY FORM WITHOUT PER-
MISSION OF THE PUBLISHER IN WRITING. THIS
APPLIES SPECIFICALLY TO PHOTOSTATIC AND
MICROFILM REPRODUCTIONS.

PRINTED IN THE UNITED STATES OF AMERICA
BY MACK PRINTING CO., EASTON, PENNSYLVANIA

This Book is Dedicated to

KAJ LINDERSTR0M-LANG and HEINZ HOLTER

of the Carlsberg Laboratory, Copenhagen, in appreciation of
their scientific achievements, which have contributed gener-
ously to the development of histo- and cytochemistry, and in
appreciation of their fine human qualities of integrity, under-
standing, and a truly civilized sense of values and of humor.

^\GAi

X

FOREWORD

To the older biologist, microchemistn.- meant the application of
appropriate reagents to sections of tissue or to intact cells to enable
him to recognize mider the microscope the nature and localization
of compounds in li^-ing substance. To the chemist, on the other hand,
it signifies the application of instruments of great precision and
rigorous methods to the accurate determination of the composition
of extremely small amounts of material. To the modem biologist,
it includes all of those methods which may aid him to delineate in
the exiguous confines of the cell that elusive and mysterious
chemical pattern which is the basis of life. To the extent that it
requires isolation and purification of compounds, microchemistn.* is
but a significant extension of the usual methods of biochemistry-,
but with the discovery- of methods of isolating microscopic and
submicroscopic and even ultramicroscopic components of lining
substance, and the application of physical and chemical methods of
analysis to them, the new microchemistrf promises to become the
most important tool we possess for elucidation of the fundamental
chemical pattern of protoplasm.

In our enthusiasm for these methods, however, we must not forget
that by far the most sensitive instrument for microchemical analysis
is the li\'ing organism itself. The methods of immunology*, for
example, suffice to discriminate between compounds so closely
related that the chemist is at a loss to distinguish between them.
The genes revealed by genetic experiment exceed by an infinite
multiple the meager number of nucleoproteins revealed by bio-
chemical research. The bioassay methods depend on the exquisite
sensiti\'ity of the li^-ing organism to minute changes in its chemical
en\-ironment. These are also microchemical methods.

In this volume, the author has chosen to follow the historical
pattern and to present the methods of microscopical analysis first.
In this field, the difficulty is not so much to find suitable reagents
as to prepare material in a form susceptible to microscopic study.
Botanical material for a long time possessed definite advantages,
since the support aft'orded by cellulose walls, and. in higher plants,

Vlll FOREWORD

by the vascular bundles, enabled the worker to obtain sections with-
out previous preparation. The introduction of the freezing-drying
technique by Gersh removed to some extent this advantage and
enabled the worker to obtain sections of animal material without
previous chemical treatment. This valuable method has been wisely
chosen as the first subject of the first section.

This section is devoted to the methods for recognition of chemical
substances in microscopic preparations. The author has contented
himself with presenting accurately and without prejudice the many
methods so far suggested for the detection of various chemical sub-
stances in tissues. In this field there are three purposes to be served
— namely, recognition, localization, and quantitation. The first of
these may be accomplished equally well by macrochemical methods
and the third has only recently achieved importance through the
introduction of the newer physical methods of measurement. Accord-
ingly, localization becomes the chief function of microchemistry of
this order. On the user rests the responsibility for perceiving and
avoiding the many pitfalls which are inevitable. Gross blunders have
been made in the past and can only be avoided in the future by
meticulous care and high critical ability on the part of the worker.
The belief that in the complex colloidal matrix of protoplasm reac-
tions occur as they do in more simplified systems in vitro is respon-
sible for many mistakes. No assumption as to solubility or insolu-
bility is admissible. Otherwise soluble substances may be firmly
adsorbed to submicroscopic surfaces or incorporated into molecular
aggregates. In those methods which require fixing, imbedding, and
cutting in order to prepare the material for microchemical tests, the
disparity between the mass of material and the volume and variety
of solvents employed makes the data on insolubility equally untrust-
worthy. The worker must examine critically each step of the process
and seek to api)raise its effect on the final result. Reaction time, drift
of highly dispersed reaction products to other locations due to surface
charges, and translocation of reaction owing to differences of ion
mobility, all must be carefully considered.

Microchemical reactions yielding colorless products and requiring
the use of a second reagent to visualize them should be accepted only
with reservation and should be used only as a first approach, subject
to the results of later confirmatory tests. To this category belong,
for example, the molybdate reactions for the detection of phosphate,

FOREWORD IX

Macallum's reaction for potassium, and the popular methods for
the detection of phosphatases. In all of these, the possibility for the
adsorption of molybdic, cobalt, or lead ions, respectively, independ-
ent of the reactions supposed to occur, should be entertained.

Negative results should not be accepted; the limitations imposed
by the microscope as to the thickness of preparations both from the
standpoint of transparency and of dispersion of light make any
negative conclusions inadmissible.

It may be inferred from the preceding remarks that this writer
views microchemical methods of this category with suspicion. This
is not the case. He simply wishes to insist that the worker scrutinize
critically every phase of his technique and consider seriously what
the value of the method may be for recognition, quantitation, and
localization, respectively.

The second, third, and fourth parts of the volume are devoted to
the methods for accurate physical and chemical microanalysis as
applied to biological problems, and to the newer methods for the
mechanical separation of the morphological constituents of proto-
plasm. This writer has already indicated above his belief that in
these methods rests our chief hope of progress in the solution of the
mystery of life.

The road to this goal is a long and laborious one. It is the writer's
hope and belief that travelers along this difficult highway will find
their burdens lightened by the collection into one volume of so many
useful methods of investigation.

R. R. Bensley

PREFACE

"But not by nature is the man of science more critical
or careful than his colleagues are. What gives him an
advantage over them is that when issues rise in his
domain they can be settled with a sureness and
dispatch which elsewhere are unknown; for science
has a priceless touchstone here to seek out truth — the
technique of measurement."

Edmund W. Sinnott in Science and the Education of
Free Men, American Scientist 32: 209 (1944).

Like a lens gathering diverse rays and concentrating them in a
new beam which penetrates depths hitherto unilluminated, each
new border science reveals a new threshold of knowledge. We who
work in the life sciences stand at such a threshold today. The lights
of histology and cytology are being joined with those of chemistry,
and the brilliant new beams converging to a focus probe beyond
the old limitations. The day is past when our vision cannot penetrate
beyond the architecture of the cell. The wealth of knowledge that
has been, and can still be gleaned from purely descriptive microscopic
anatomy is not to be minimized, but under the new illuminations
we can begin to discern the chemical patterns in the cellular archi-
tecture. From this knowledge an understanding of the functions of
the patterns will follow. Thus, from histology and cytology, as we
have known them in the past, the new field of histo- and cyto-

XI

Xll PREFACE

chemistry is arising, and from this new field, a histo- and cyto-
physiology will develop — and so on in the expanding and exciting
quest into the nature of living processes. These new instruments and
techniques which carry our vision deep into the living unit, to the
molecules and the atoms — they include the beautiful ingenuities
which have been refined out of the mountain of past scientific
experience, and some which have been newly created for the purpose.
It is these instruments and techniques with which we shall be
concerned in this book.

In a discussion on cytological technique, J. R. Baker (1942) of
Oxford stated, "It was once remarked to the writer that biochemists
like to have their substances in test-tubes. The cytologist wants to
have his exactly where they were in life, and to know, as precisely
as he can, what they are. When substance and structure are known,
the way is clear for the elucidation of the main problem of cytology,
which is to discover what a cell does to keep alive and to perform
its functions for the body as a whole or for the next generation." It
isn't so much "that biochemists like to have their substances in test-
tubes" as it is that, until not long ago, biochemists had no means of
dealing with substances except in "test-tubes." That biochemists
have been fundamentally dissatisfied with this limitation is apparent
in their growing efforts to refine their techniques to enable investiga-
tions in situ. We can be sure that future campaigns designed to as-
sault the present horizons of cytology will follow the strategic lines
made possible by the development of equipment and procedures
which bring chemical investigations directly to the cell, and the parts
of the cell, existing in natural milieu. In the following pages we shall
examine the techniques and devices already elaborated for this
purpose, and no one with imagination will fail to be impressed and
excited by the possibilities engendered.

Lison's Histochemie Animate, a book dealing mainly with micro-
scopic techniques of chemical morphology, was published in Paris
in 1936. Many notable advances have occurred since then, and the
present volume has been designed to bring together in a compact
and readily available form detailed descriptions, not only of the
morphologic, but also of the quantitative techniques. Insofar as
it has been possible, the material presented has been brought up to
date as of January 1, 1947. Pertinent publications which have ap-
peared between January 1 and September, 1947, have not been

PBEFACE XIU

discussed, but a bibliography appendix containing these references
has been included at the end.

To those who treat with condescension all science that cannot be
quantitated, it might be said, "True, morphology is only a stepping
stone, and while as a stepping stone it is not a place to stop, it is
none the less basic." And to those whose comprehension is confined
to mere morphology, it might be said, "True, morphology is basic,
but it is only a stepping stone in science." And to both it should be
said, "All fields of science are stepping stones, and in science there
are no places to stop."

The author wishes to express his appreciation to the following for
critically reviewing various sections of this volume and offering many
helpful suggestions: Dr. R. R. Bensley, Dept. of Anatomy, University of
Chicago; Dr. W. L. Doyle, Dept. of Anatomy, University of Chicago;
Dr. 0. H. Lowry, Dept. of Pharmacology, Washington University; Dr.
G. H. Scott, Dept. of Anatomy, Wayne University; and Dr. J. M. Tobias,
Dept. of Physiology, University of Chicago. The encouragement in this
undertaking of Dr. M. B. Visscher, Dept. of Physiology, University of
Minnesota; and Dr. B. Sullivan, Director of Laboratories, Russell-Miller
Milling Co., Minneapohs, is also gratefully acknowledged. The author is
grateful for the invaluable assistance of the pubhsher's staff.

David Glick
October, 1948
Minneapolis

CONTENTS

Foreword vu

Preface • xi

Abbreviations xxiv

Microscopic Techniques 1

I. Freezing-Drying Preparation of Tissue 3

Parker-Scott Method for Freezing-Drying Tissues 5

II. Cliemical Methods U

A. Requirements 11

B. Inorganic Elements and Radicals 14

Potassium 14

Gersh Modification of Macallum Method for Potassium 14

Carere-Comes Siena Orange Method for Potassium 15

Calcium 16

Calcium Sulfate Test for Calcium in Plant Tissue 17

Cretin Color Test for Calcium and Other Minerals 17

Von Kossa Silver Test for Calcium 17

Magnesium ■ 18

Broda Method for Magnesium 18

Zinc 18

Mendel and Bradley Method for Zinc 19

Iron 19

Prussian-Blue Test for Ferric Iron and TurnbuU's Blue for

Ferrous Iron 20

Humphrey Dinitrosoresorcinol Test for Iron 21

Thomas and Lavollay Hydroxyquinoline Test for Iron 21

Nickel 22

Cretin and Pouyanne Method for Nickel 22

Lead and Copper 22

Mallorj' and Parker Hematoxylin Method for Lead and Copper 23
•• Mallory and Parker Methylene Blue Method for Lead and

Copper 24

Mercury 24

Method of Hand et al. for Mercurous and Mercuric Mercury . . 25

Silver 25

Okamoto et al. Procedure for Silver 26

Gold 27

Roberts Method for Gold 27

Elftman and Elftman Method for Gold 28

Okamoto et al. Method for Gold ; 28

Platinum 29

Palladium 29

Uranium 29

Arsenic 30

Castle Cupric Salt Method for Arsenic 30

Bismuth 30

Wachstein and Zak Method for Bismuth 31

Castel Method for Bismuth 32

XV

63013

Xvi CONTENTS

Microscopic Techniques (continued)

Chloride and Phosphate-Carbonate 32

Gersh Method for Chloride and Phosphate-Carbonate 33

Iodide 34

Phosphate 34

Serra and Queiroz Lopes Modification of the Molybdate

Method for Phosphate 34

Nitrate 35

Cramer Method for Nitrate 35

Sulfhydryl and Disulfide Groups 36

Nitroprusside Test of Rapkine 36

Bourne Nitroprusside Test 37

Hammett and Chapman Nitroprusside Test 37

Organic Substances and Groups 38

Lipids and Cholesterol 38

Kay and Whitehead Procedure for the Sudan IV Stain
for Lipids 39

Jackson Procedure for Lipids Using Acetic-Carbol-Sudan III . . 39

Telford Govan Technique for Sudan Dye Staining in
Aqueous Media 40

Schultz Cholesterol Test 41

Carotene, Carotenoids, and Vitamin A 42

Steiger Method for Carotene in Leaves 42

Riboflavin 43

Chevremont and Comhaire Method for Riboflavin 43

Polysaccharides in General 43

Method of Hotchkiss for Polysaccharides 44

Acid Polysaccharides— Hyaluronic Acid 45

Hale's Method for Acid Polysaccharides 45

Mucoproteins 46

Lison's Method for Polysaccharide Sulfate Compounds (after

Sylven) 47

Glycogen and Mucin 47

Bauer-Feulgen Stain for Glycogen (after Bensley) 48

Best Carmine Stain for Glycogen (after Bensley) 49

Gomori Procedure for Glycogen and Mucin 50

Starch 51

Milovidov Method for Starch 51

Cellulose 52

Post and Laudermilk (1942) Iodine Stain for Cellulose 52

Chitin 52

Murray Method for Softening Chitin 53

Diaphanol Method for Softening Chitin 53

Methods for Staining Chitin 54

Ascorbic Acid 54

Bourne Silver Stain for Ascorbic Acid 55

Protein Reactions 56

Arginine and Arginine-Containing Proteins 56

Serra Method for Arginine and Arginine-Containing Proteins. . 57

Thomas Method for Arginine and Arginine-Containing Proteins 58
Tryptophane in Proteins 58

Romieu Reaction for Tryptophane in Proteins 59

Tyrosine in Proteins 59

Millon Reaction for Tyrosine in Proteins (after Bensley

and Gersh) 59

a-Amino Acid Groups in Proteins 60

Berg Ninhydrin Test for o-Amino Acid Groups 60

CONTENTS XVll

Microscopic Techniques (continued)

Melanin 61

Dublin Application of the Bodian Method to Demonstration

of Melanin 61

Hemoglobin 62

Ralph Method for Hemoglobin 62

GouUiart Method for Hemoglobin 62

Dunn Method for Hemoglobin 63

Bile Pigments and Acids 63

Stein Test for Bile Pigments 63

Forsgren Test for Bile Acids 64

Aldehydes, Nucleic Acids, and "Plasmal" 65

Coleman Preparation of Feulgen Reagent 67

Whitaker Feulgen Technique for Plant Tissues 67

Cowdry Modification of Feulgen Reaction for Paraffin

Sections of Animal Tissues 68

Oster Modification of Feulgen Reaction for Fresh-Frozen

Sections of Animal Tissues 68

Method of Turchini et al. for Nucleic Acids 69

Water-Insoluble Carbonyl Compounds 69

Bennett Use of Phenylhydrazine Reaction for Water-Insolu-
ble Aldehydes and Ketones 70

Albert and Leblond Use of 2,4-Dinitrophenylhydrazine Re-
action for Water-Insoluble Aldehydes and Ketones 71

Bennett Use of Semicarbazide Reaction for Water-Insoluble

Aldehydes and Ketones 71

Purines - 72

Murexide Test for Certain Purines 72

Hollande Modification of the Courmont-Andre Method for

Uric Acid and Urates 73

Indole and Related Compounds 73

Phenols 74

Lison Modification of the Chromaffin Reaction 74

Urea 75

Sulfonamides 75

Method of MacKee et al. for Sulfonamides 76

D. Enzymes 77

Urease 77

Alkaline Phosphatase 78

Gomori Revised Method for Alkaline Phosphatase 79

Acid Phosphatase 80

Gomori Revised Method for Acid Phosphatase in Animal

Tissues 81

Glick and Fischer Adaptation to Grains and Sprouts of

Gomori Method for Acid Phosphatase 82

Other Phosphatases 85

Zymohexase (Aldolase plus Isomerase) 86

Allen and Bourne Method for Zymohexase 87

Lipase 88

Gomori Revised Method for Lipase 89

Peroxidase 90

McJunkin Method for Peroxidase in Tissue Sections 90

Armitage Method for Peroxidase in Blood or Bone Marrow

Smears 91

Dopa Oxidase 91

Laidlaw Method for Dopa Oxidase 92

Amine Oxidase 93

Oster and Schlossman Method for Amine Oxidase 93

Xviii CONTENTS

Microscopic Techniques (continued)

Cytochrome Oxidase 94

Graff Method for Cytochrome Oxidase in Fixed Tissue

("M. Nadi Oxidase") 94

Graff Method for Cytochrome Oxidase in Fresh Tissue

("G. Nadi Oxidase") 95

Loele Method for "a-Naphthol Oxidase" 96

Succinic Dehydrogenase* 96

Semenoff Method for Succinic Dehydrogenase 96

III. Physical Methods 99

A. Fluorescence Microscopy 99

1. Apparatus 99

2. Preparation of Tissues 102

3. Photomicrography , 102

4. Characterization of Substances 104

Direct Observation of Fluorescence 104

Spectroscopic Analysis of Fluorescence 108

B. Emission Histospectroscopy 109

1. PoHcard Technique 109

2. Scott and Williams Technique 110

C. Visible and Ultraviolet Absorption Histospectroscopy 113

1. Caspersson in Situ Technique 114

2. Norberg Technique 120

Apparatus 120

Manipulations 121

The Sample Slide for Absorption Measurements 122

Accessories for Norberg Technique 123

Methods 124

Phosphorus 124

Norberg Method for Phosphorus 124

3. Stowell Technique 125

D. Roentgen Absorption Histospectroscopy 127

1. Some Theoretical Aspects 128

2. Thickness of Sections 132

3. Apparatus 133

4. Measurement of Density of Photographic Film 138

Nomogram 140

E. Microincineration 140

1. Preparation for Incineration 141

2. The Incineration Furnace 142

3. Scott Incineration Procedure 143

4. Microscopic Examination and Interpretation 144

5. Quantitative Estimation of Ash 146

F. Analytical Electron Microscopy 147

The Scott-Packer Analytical Electron Microscope 148

Manipulation 151

G. Radioautography 152

Preparation of Radioautographs 156

Belanger and Leblond Technique 157

Discussion 160

Chemical Techniques 163

Introduction 165

I. General Apparatus and Manipulation 165

A. Vessels, Stoppers, Holders, etc 165

B. Microliter Pipettes 170

CONTENTS XlX

Chemical Techniques (continued)

C. Filters 176

D. Stirring Devices 178

E. Heating Devices 180

F. Moist Chambers 181

G. Electrodes 183

H. Conductivity Apparatus 188

I. Balances 189

II. Colorimetric Techniques 195

A. Capillary Tube Technique 195

1. Apparatus 196

2. Manipulations 197

3. Methods 198

Preparation of Protein-Free Supernatants 198

Tungstic Acid Supernatants 198

Trichloroacetic Acid Supernatants 199

Zinc Sulfate-Sodium Hydroxide Supernatants 200

Chloride 200

Westfall, Findley, and Richards Method for Chlorides 200

Sodium 203

Bott Method for Sodium 203

Phosphate 208

Walker Method for Phosphate 208

Phosphatase 209

Weil and Russell Method for Phosphatase 209

Reducing Substances 210

Walker and Reisinger Method for Reducing Substances 211

Creatinine 211

Method of Bordley et al. for Creatinine 211

Uric Acid 213

Bordley and Richards Method for Uric Acid 213

Urea 214

Walker and Hudson Method for Urea 214

Hydrogen Ion Concentration 216

B. Cuvette Technique 216

1. Apparatus '. 216

2. Methods 219

Calcium 219

Sendroy Method for Calcium 219

Chloride 224

Sendroy Method for Chloride 225

Phosphate and Phosphatase 226

Lundsteen and Vermehren Method for Inorganic Phosphate

and Phosphatase 227

Lowry and Lopez Method for Inorganic Phosphate in

Presence of Labile Phosphate Esters 228

Bessey, Lowry, and Brock Method for Phosphatase 229

Nitrogen and Ammonia 230

General ; 230

Digestion of Sample for Determination of Total Nitrogen ...234

Le\'y Nesslerization Method for Nitrogen 237

Russell Phenol-Hypochlorite Method for Ammonia 238

Uric Acid, Creatine, Creatinine, and AUantoin 239

Borsook Method for Uric Acid 240

Borsook Method for Creatine and Creatinine 242

Sure and Wilder Method for Creatine and Creatinine 243

Borsook Method for AUantoin 243

XX CONTENTS

Chemical Techniques (continued)

Ascorbic Acid 245

Lowry, Lopez, and Bessey Method for Ascorbic Acid 245

Glycogen 247

Boettiger Method for Glycogen 248

Van Wagtendonk, Simonsen, and Hackett Method for

Glycogen 249

Vitamin A and Carotene 250

Method of Bessey et al. for Vitamin A and Carotene 250

III. Titrimetric Techniques 255

A. Microhter Burettes 255

B. Photometric End Points 265

C. Methods 265

Sodium and Potassium (Combined) 265

Linderstr0m-Lang Method for Sodium and Potassium 266

Potassium 268

Norberg Method for Potassium 268

Sodium 270

Lindner and Kirk Method for Sodium 271

Calcium 272

Siwe lodometric Permanganate Method for Calcium 273

Siwe Acidimetric Method for Calcium 274

Lindner and Kirk Cerimetric Method for Calcium 275

Iron 276

Kirk and Bentley Method for Iron 277

Ramsay Method for Iron 278

Phosphorus 280

Chloride 281

Linderstr0m-Lang, Palmer, and Holter Electrometric

Method for Chloride 282

Nitrogen and Ammonia 283

Brliel, Holter, Linderstr0m-Lang, and Rozits Method for

Nitrogen and Ammonia 283

Urea 286

Kinsey and Robison Method for Urea 286

Urease 287

Amide, Peptide, and Nitrate Nitrogen 288

Borsook and Dubnoff Methods for Amide, Peptide, and

Nitrate Nitrogen 288

Acid, Alkah, Amino, and Carboxyl Groups 290

Lipid 291

Schmidt-Nielsen Method for Lipid 291

Extraction and Fractionation of Lipids 292

Schmidt-Nielsen Method for Extraction and Fractionation

of Lipids 293

Iodine Number of Lipids 294

Schmidt-Nielsen Method for Iodine Number 295

Reducing Sugars 296

Holter and Doyle Modification of Linderstr0m-Lang and
Holter lodometric Method for Reducing Sugars 296

Heck, Brown, and Kirk Cerimetric Method for Reducing

Sugars 297

Glycogen 298

Heatley Method for Total Glycogen 299

Heatley and Lindahl Method for Desmo- and Lyoglycogen 300
Ascorbic Acid 300

GHck Method for Ascorbic Acid 301

CONTENTS XXI

Chemical Techniques (continued)

Amylase 302

Proteolytic Enzymes 302

Linderstr0m-Lang and Holter Acidimetric Acetone Method
for Proteolytic Enzymes 303

Linderstr0m-Lang and Duspiva Alkalimetric Alcohol
Method for Proteolytic Enzymes 304

Weil Formol Method for Tryptic Activity 306

Arginase 306

LinderstrcSm-Lang, Weil, and Holter Methods for Arginase. .307
Esterases and Lipases 308

Glick Acidimetric Method for Esterase and Lipase 309

Glick Acidimetric Method for Cholinesterase 310

Catalase 310

Holter and Doyle Method for Catalase 310

IV. Gasometric Techniques 313

A. Volumetric 313

1. Capillary Respirometry 314

(a) Cunningham-Barth-Kirk Differential Respirometer 314

(b) Cunningham-Kirk Open-Tube Respirometer 319

(c) Tobias-Gerard Respirometer 322

(d) Scholander Micrometer-Burette Differential Respirometers 324

2. Gas Analysis 326

(a) Scholander Micrometer Burette Gas Analyzer 326

(b) Berg Simplified Gas Analyzer 328

(c) Scholander-Roughton Syringe Gas Analyzer 329

Oxygen 331

Roughton and Scholander Method for Oxygen 331

Carbon Monoxide 334

Scholander and Roughton Method for Carbon

Monoxide 334

Nitrogen 335

Edwards, Scholander, and Roughton Method for

Nitrogen 336

Carbon Dioxide 337

Scholander and Roughton Method for Carbon Dioxide. .337

3. Membrane Interferometer Volumetry 340

B. Manometric 342

1. Cartesian Diver Manometry 342

(a) Microliter Diver Technique 343

(b) 0.1 Microliter or Capillary Diver Technique 382

(c) Methods Other Than for Respiration 393

Cholinesterase 393

Lindestr0m-Lang and Glick Method for Cholinesterase . 393

Thiamine and Cocarboxylase 394

Westenbrink Method for Thiamine and Cocarboxylase 395

Diphosphopyridine Nucleotide 396

Anfinsen Method for Diphosphopyridine Nucleotide ...396

2. Optical-Lever Respirometry 399

C. Polarographic 404

Microelectrode Measurement of Local Oxygen Tension in Tissue 404

V. Dilatometric Techniques 413

Dilatometric Apparatus and Its Use 414

Peptidase 417

Method of Linderstr0m-Lang and Lanz for Peptidase 419

Density and "Reduced Weight" 420

XXll - CONTENTS

Chemical Techniques (continued)

VI. Determination of Amount of a Biological Sample 423

A. Preparation of Frozen Tissue Sections of Accurate Thickness 427

Linderstr0m-Lang and Mogensen Method for Accurate Cutting
and Special Handling of Frozen Tissue Sections 428

B. Microscopic Examination and Chemical Analysis of the Same

Tissue Section 430

Method of Anfinsen et al. for Microscopy and Analysis on
the Same Tissue Section 431

C. Volume of Irregularly Shaped, Small Biological Samples 431

Holter Method for Measurement of Volume 432

VII. Deductive Methods 435

Microbiological Techniques 43

Introduction 439

Riboflavin 439

Lowry and Bessey Method for Riboflavin 440

Mechanical Separation of Cellular Components 445

Introduction 447

I. Types of High-Speed Centrifuges 448

II. Separation of Components of A. punctulata Eggs (after Harvey) 452

III. Isolation of Cell Nuclei 454

A. Nuclei from Avian Erythrocytes 454

Laskowski Procedure for Isolation of Nuclei 454

Dounce and Lan Procedure for Isolation of Nuclei 455

B. Nuclei from Other Cells 456

Stoneburg Procedure for Isolation of Nuclei 456

Dounce Procedure for Isolation of Nuclei 456

Lazarow Procedure for Isolation of Liver Nuclei as Used by

Hoerr 458

Behrens Procedure for Isolation of Nuclei from Thymus and

Lymph Cells (as Modified by Gulick et al.) 459

IV. Isolation of Chromatin Threads from Cell Nuclei 461

Claude and Potter Procedure . .v 461

V. Isolation of Cytoplasmic Particulates 462

A. Mitochondria 462

Bensley and Hoerr Procedure for Guinea Pig Liver 462

Claude Procedure for Certain Neoplastic Cells of the Rat 463

Claude Procedure for Isolation of "Large Granules" from Liver 464

B. Submicroscopic Particulates 465

Lazarow Procedure for Separation and Isolation of Lipoprotein

and Glycogen Particles from Guinea Pig Liver 466

Claude Procedure for Isolation of "Microsomes" 466

VI. Isolation of Chloroplasts from Leaf Cells 468

Granick Procedure 469

Neish Procedure 469

VII. Isolation of Other Particulates from Cells 471

Bibliography 473

Bibliography Appendix 504

List of Manufacturers 508

Index 511

ABBREVIATIONS

1. liter

ml. 10~^ liter

lA. 10-6 liter

gal. gallon

g. gram

mg. 10~^ gram

[xg. 10-6 gram

m/xg. 10"*^ gram

lb. pound

oz. ounce

D.C. direct current

R.P.M. revolutions per minute

E.M.F. electromotive force

m. meter

cm. 10-^ meter

mm. 10"^ meter

fi 10-6 meter

mix. 10~^ meter

A 10-^0 meter

( angstrom unit)

X.U. 10"^ angstrom

ft.

foot

in.

inch

yr-

year

hr.

hour

min.

minute

sec.

second

amp.

ampere

/xamp.

10-6 ampere

/.F.

10-6 farad

Kev

10^ electron volts

M

molar

mil/

10-^ molar

N

normal

cone.

concentrated

dil.

dilute

soln.

solution

sp. gr.

specific gravity

C. P.

chemically pure

c.p.

candle power

m.p.

melting point

b.p.

boiling point

All temperatures are given in degrees centigrade.
If not otherwise indicated, all solutions are understood to be aqueous.
If not otherwise indicated, the term alcohol refers to 95% ethyl
alcohol.

xxiv

4

MICROSCOPIC TECHNIQUES

"She (Science) warns me to be careful how I adopt a
view which jumps with my preconceptions, and to
require stronger evidence for such a beUef than for
one to which I was previously hostile. My business
is to teach my aspirations to conform themselves to
fact, not to try and make facts harmonize with my
aspirations. — Sit down before fact as a little child,
be prepared to give up every preconceived notion,
follow humbly wherever and to whatever abysses
nature leads, or you shall learn nothing."

Thomas Huxley
in a letter to Charles Kingsley, September 23, 1860.

The microscopic techniques which are treated in the present vol-
ume are those designed to establish the distribution of elements,
groups, substances or activities in microtome sections of tissue by
means of examinations under some form of microscope. This re-
quires that the factor in question be made apparent by character-
istic optical or photographic properties such as a specific color,
fluorescence, or radiation. With the exception of absorption histo-
spectroscopy, these microscopic techniques are limited to observa-
tions which are essentially qualitative in nature. However, the
microscopic techniques permit a much greater degree of localization
of particular chemical constituents in histologically defined cells, or
cytologically defined parts of cells, than is possible by means of the
quantitative chemical techniques. Thus, one is often forced to
choose between degree of localization and quantitation. Obviously,
it would be preferable to establish both the cellular disposition of
biologically significant factors and their quantitative relationships.

/. FREEZING DRYING PREPARATION

OF TISSUE

Since the microscopic techniques that will be discussed are almost
all based on the use of microtome sections of tissue, it is pertinent
that the freezing-drying preparation for sectioning be described in
detail. This technique of sudden cooling to low temperatures and
rapid dehydration of the frozen material in vacuo has many advan-
tages over the usual histological methods employing fixing and
dehydrating solutions. The chief of these advantages are a minimum
of chemical change in the tissue (there is an almost instantaneous
cessation of metabolic activity and no chance for other chemical
changes to occur), a mimum of shifting of diffusible constituents
(fluid is not used and the fixation is immediate), a greater preserva-
tion of cytoplasmic inclusions than is possible with the use of fixing
solutions, the possibility of direct paraffin infiltration of dehydrated

4 MICROSCOPIC TECHNIQUES

tissue, and the absence of cell shrinkage. These are no inconsider-
able advantages, and the freezing-drying technique should be given
the preference wherever possible.

Scott ( 1943) has been careful to point out that, while distortion
of mineral distribution might be expected to occur as the result of
ice crystal formation during the freezing and that artifacts might
be occasioned by the paraffin infiltration, neither of these appears to
be a serious difficulty in the more recent improved techniques. As
Scott indicated, on the one hand ice crystal formation could be
readily recognized should it occur in a manner that might influence
interpretation, and on the other it has been impossible thus far to
find evidence of distortion resulting from the infiltration, although
various control experiments have been performed to test this
possibility.

Gersh (1932) extended the Altmann method of dehydrating tissue
in vacuo at liquid-air temperatures, and the improved procedure is
known as the Altmann-Gersh technique.* In a critical study Scott
(1933a) pointed out that the dehydration temperature of —20°
used by Gersh was not low enough to prevent a certain amount of
ion diffusion since this temperature is above the eutectic point of
certain naturally occurring salt systems. In the improved cryostat
of Packer and Scott (1942), to be described in detail later, the
temperature is maintained below — 54.9°, the eutectic point of
CaCl2.6H20. When he first indicated the desirability of using lower
temperatures, Scott ( 1933a) recommended the use of alcohol cooled
to — 177°, instead of liquid air, since the latter gives rise to a gas
envelope around the tissue which retards the rate of freezing. A
further improvement was effected by Hoerr ( 1936) , who found that
more rapid freezing (hence smaller ice crystals) was obtained by
placing tissue in isopentane cooled to — 160° to — 195° by means of
liquid nitrogen. Among others, Simpson (1941) confirmed the
advantages of the isopentane method and, in addition, pointed out
the desirability of employing small pieces of tissue for treatment
since the centers of larger pieces do not yield sections of the highest
quality.

After the appearance of the Gersh (1932) vacuum dehydrator,
other types were described by Goodspeed and Uber (1934) and
Scott and Williams (1936). However, since none of these was

* The Gersh apparatus is available from A. S. Aloe & Co.

FREEZING-DRYING PREPARATION OF TISSUE 5

wholly satisfactory, Packer and Scott (1942) developed a cryostat
of a new design that is the finest instrument yet devised for the
freezing-drying of tissues. An important feature of this apparatus
is that the frozen and thoroughly dried tissue can be brought
gradually to the temperature of the melted paraffin, and then it
can be embedded without contact with the moisture of the air.
Previous practice was to transfer the very cold tissue from the
cryostat to the air, and then plunge it directly into melted paraflEin,
thus subjecting it to a sudden temperature change of about 100°.
Sjostrand (1944) described a freezing-drying apparatus somewhat
simpler than the Packer-Scott instrument but it was not designed to
permit paraffin infiltration within the apparatus.

PACKER-SCOTT METHOD FOR FREEZING-
DRYING TISSUES

The Freezing-Drying Apparatus. In the diagram of the ap-
paratus, which is made of Pyrex glass (Fig. 1), the drying chamber
(C) is a 2.5 in. tube 12.5 in. long surrounded by a jacket of about
3.5 in. diameter that can be exhausted through stopcock B connected
by glass tubing to S. The 3 gal. Pyrex Dewar flask (Di), containing
solid carbon dioxide in butyl alcohol is used to cool the drying
chamber, and it is arranged so that it can be easily lowered away
from the apparatus. A commercially built refrigerator has also been
employed in place of solid carbon dioxide for the cooling by Hoerr
and Scott (1944). When a pressure of 1 mm. of mercury, or less,
is maintained in the space E, and paraffin is in tube C, the equi-
librium temperature over the paraffin is about — 66°. As used at
present, there is no occasion to employ temperatures higher than
— 66°, but Packer and Scott point out that a thermocouple sealed
in space E could be used to operate a thermostat which in turn could
control the current in the paraffin heater (D) in order to maintain
temperatures above — 66°. The heater (D) is required to melt the
paraffin in tube C so that the tissues held in the copper gauze basket
(Ci) can be embedded in vacuo.

The heater (D) is constructed by covering a thin-walled copper
cylinder with liquid porcelain (Insa-lute), and, after dry, winding
No. 18 Nichrome wire (about 70 ohms) over it and applying another
layer of liquid porcelain over the wire. A thin-walled sleeve of cop-

6 MICROSCOPIC TECHNIQUES

per is then fitted over the whole and, after testing the unit, elec-
trical connections to the outside are made through tungsten glass
seals. Small glass projections on the outside of the drying tube
near the bottom serve to support the heating unit in its proper
position.

Two glass boats {H) contain the phosphorus pentoxide used as
the drying agent. They are placed in tube G, which is 4 in. in di-
ameter, through the opening at J. The closures at A and J are
grease joints fitted with springs in the usual manner. All connect-
ing tubes on the low pressure side of the apparatus have a diameter
of 1.5 in. The vapor trap {K) has a 1.5 in. inner tube and a 0.5
in. annular space, and the inner tube projects about 5 in. below the
level of the solid carbon dioxide in butyl alcohol used as a re-
frigerant to surround the trap.

The two-stage diffusion pump (L) and the single-stage booster
pump {N) employ Octoil-S {Distillation Products Co.) instead of
mercury since the former has a very low vapor pressure (claimed
to be < 10"*^ mm. of mercury at 15°), effects a very high pumping
speed, and is much cheaper than mercury. The pump (L) has an
intake speed of 10 liters per second ; it was designed by Professor A.
L. Hughes of the Washington University Department of Physics.
The boilers of the vapor pumps were protected from drafts, etc. by
a covering of several layers of wet asbestos paper and over these
a thin aluminum cone (M) . Turned-under tabs of the cone support
a flat circular 300 watt heater ( Chromolox) . A small electric blower
is used to cool the upper chamber of the boiler of pump L for
maximum efficiency. A single-stage Welch mechanical pump is con-
nected by rubber pressure tubing at U to the booster pump {N).
The phosphorus pentoxide trap (0) prevents contamination of the
oil when the vacuum is broken by stopcock P. The stopcock Q
has a 1 cm. bore and is used to isolate the high-vacuum section
from the mechanical pump while the space E is being exhausted.
The two-way stopcock R connects to the air at T and to E through
S.

The type of support employed for the glass apparatus is shown in
Figure 2. The main standard is a rectangular aluminum box 4 ft.
high bolted to a % in. iron base plate 2 ft. square. The aluminum
box is made by welding together at W two heavy 8 in. aluminum
channels (V). The glass is supported by copper rings silver-

FREEZING-DRYING PREPARATION OF TISSUE 7

soldered to Vie in. brass rods which are threaded into the aluminum
standard. An instrument panel is mounted under G to carry the
rheostats and meters for controlling the heating current for the
diffusion pumps and the paraffin chamber.

Detail of
joints A and J

Detail of
paraffin heater D

Fig. 1, The Packer-Scott (1942) freezing-drying apparatus.

Fig. 2. Arrangement for supporting the Packer-Scott (1942)
freezing-drying apparatus.

In order to ascertain something of the state of dehydration of
the tissue the ionization pressure gauges F and I in Figure 1 are
fitted into tube G. When the rate of evaporation falls to a very

8 MICROSCOPIC TECHNIQUES

low value, the pressure gradient between F and / disappears. How-
ever, since water may diffuse out, it has been found desirable to
continue the pumping for a day or more after the pressures at
F and / are coincident and remain so.

The ionization gauges used, of the type described by Montgomery
and Montgomery ( 1938) , are No. 47 radio tubes of Radio Corpora-
tion of America, which are sealed to the vacuum system with black
vacuum wax, care being taken to avoid knocking off the filament
coating during the sealing. A power pack is employed consisting
of a half-wave rectifier, filter system, and power transformer. An
additional filament transformer is included to supply the filament
of the second gauge. In order that the same meters and galva-
nometer may be used for both gauges a double-throw triple-pole
switch is employed. For pressures less than 3 X 10"^ mm. of mer-
cury a student type of wall galvanometer is used to measure positive
ion current in the gauges; this current is directly proportional to
the pressure. For higher pressures a microammeter with a range
of 0-50 may be used. Since the usual calibration constant (1
yuamp. for 7 X lO"*' mm. of mercury) is for air, a different constant
would apply in the presence of water vapor, hence only relative
pressures are obtained.

PROCEDURE

1. Place paraffin in apparatus; melt and degas it in vacuo by
means of the mechanical pump alone.

2. Let the paraffin solidify and raise the cooling Dewar flask
around the drying chamber. When equilibrium is attained the
system is ready for the tissue.

3. Either freeze the tissues in liquid air or, preferably, in isopen-
tane at liquid air or liquid nitrogen temperatures. Violently agitate
the isopentane to speed the heat extraction.

4. Place frozen tissue in the copper gauze basket and transfer
immediately to drying chamber of apparatus.

5. Start the mechanical pump at once; then start the diffusion
pumps and run for 12 hr. before taking pressure readings. The
gauge filaments must be heated and gas allowed to escape for
several hours before reliable readings are possible. After the read-
ings of both gauges are equal, continue pumping for some time
depending on the size, shape, and character of the tissue. No rule

FREEZING-DRYING PREPARATION OF TISSUE 9

can be applied here since the time for total dehydration is a function
of a number of poorly defined variables.

6. When dehydration is considered complete, lower away the
Dewar cooling flask and allow the tissue to come to room tempera-
ture.

7. Start the paraffin heater and keep the temperature of the
paraffin just above its melting point. The top of the paraffin will
melt first; regulate the heating so that a portion of solid paraffin
remains at the bottom of the chamber. Conditions of — 66° and
4 X 10"^ mm. of mercury are obtained routinely during operation.
The tissue will sink into the melted paraffin without causing a single
bubble to rise if the tissue is properly dehydrated and the paraffin
completely degassed; otherwise bubbling will occur.

8. Break the vacuum after infiltration is complete and the par-
affin has been allowed to solidify; remove and block for cutting.

//. CHEMICAL METHODS

A. REQUIREMENTS

The microscopic technique employing chemical methods depends
in almost every case on the direct observation of an insoluble
product of a microchemical reaction between the substance or group
whose distribution is being investigated and a suitable reagent.
Since the whole purpose of these methods is to visualize the presence
of a cellular or intercellular constituent in situ, it is essential that
the tissue be handled in a manner that will not permit the con-
stituent to diffuse or change its anatomical position during the pro-
cedure. The minimum requirements of the chemical method then
may be listed as:

1. The preparation of microtome sections in which there has
been no significant alteration in the position of the constituent
being investigated.

2. A reagent which is specific for this tissue constituent.

3. A reaction between the reagent and constituent which is of
such a nature, and rapid enough, to obviate diffusion of the con-
stituent or of the reaction product.

4. A reaction product, thus trapped in situ, which is capable of
being visualized.

The frequency with which these requirements can be met is,
unfortunately, still very low. The problem is most difficult in the
case of those constituents which are diffusible in solution, e.g.,
inorganic ions. While it is possible to prepare tissue sections without
the use of solutions by means of the freezing-drying technique, the
chemical formation of a substance in these sections for purposes of
visualization involves the use of a reagent in solution. One might
imagine that, if the interaction of the reagent in solution with the
ion in the tissue were rapid, the ion would be bound as an insoluble

11

12 MICROSCOPIC TECHNIQUES

substance before serious diffusion could occur. Still, in the case
of the precipitation of tissue chloride by silver nitrate solution, Scott
and Packer (1939) pointed out that differences in ionic mobilities
and the effects of ionic charges at cellular interfaces might easily
produce precipitations in regions different from those in which
the chloride originally existed. On the other hand, Gersh (1941)
claimed that the results he obtained from chloride distribution,
using silver nitrate as the reagent, were valid as borne out by
related data obtained with entirely different biochemical methods.
Regardless of the merits in this particular instance, the dangers
indicated by Scott and Packer cannot be ignored, and no way has
yet been devised to really eliminate them ; they constitute a funda-
mental limitation in the application of the chemical methods of
microscopic technique.

In the special case of the localization of enzymes, the sites of
activity may be determined in tissue sections by immersing the
sections in a buffered substrate medium containing a reagent which
will bind one of the products of the enzymatic action in situ by
precipitation. In addition to the four requirements already listed
for determinations of the disposition of tissue constituents, it is also
necessary that the following be included for enzyme methods:

5. A reagent which when added to the buffered substrate will
react with one of the enzymatic products but not with the substrate
or buffer.

6. A reagent which will also have no untoward effect on the
enzyme.

7. If the enzymatic product which reacts with the reagent is a
substance pre-existing in the tissue, either the sites of enzyme action
must be different from those of the pre-existing substance, or the
increase in the amount of the visualized compound resulting from
the enzyme activity must be demonstrable, or, better yet, the sub-
stance must be removed in advance by a method which will not
take out the enzyme.

8. A control experiment in which either the substrate is omitted,
or a highly effective soluble enzyme inhibitor, such as fluoride, is
added (the inhibitor must not react with substrate, buffer, reagent,
or products") — the advantage of the inhibitor is that, in some cases,
naturally occurring substrate may be present with the enzyme and
thus give a false aspect to the nonenzyme control. No such control

CHEMICAL METHODS • 13

experiment is required if all substances pre-existing in the tissue and
capable of giving a positive reaction can be removed without seri-
ously reducing the enzyme activity. This has been accomplished
in certain tissues for the phosphatase test.

Many, if not most, of the tests described in the following pages
leave much to be desired. In some cases they have been developed
for particular tissues and cannot be adapted to others without a
certain amount of additional research. Most of the tests are clearly
not at all good. However, it is the purpose of the writer to present
the published methods for the localization of substances, groups,
and enzymes, even though they may be, and usually are, poor. In
this way the investigator who requires a particular method will
have at hand the procedures already developed, and, if they should
prove inadequate, at least he will have them as a basis from which
to work out improvements.

A word should be added concerning the mounting media employed
for tissue sections. The media given in the procedures that follow
are those used by the original authors. However, newer media are
available and they may be substituted for the balsam or damar
that have been employed in the past. A 60% solution of Clarite in
xylol appears to be superior to neutral balsam for mounting sections
since, according to Lillie (1941), Clarite does not promote the
fading of some stains to the degree that balsam does. Stowell and
Albers ( 1943) showed Clarite absorbs less visible light than balsam.
Tetrachloroethylene may also be used as a solvent for Clarite.
Clarite and Clarite X (also called Nevillite V and Nevillite No. 1,
respectively) are superior in all respects to balsam and damar,
according to Groat ( 1939) . A solution of 60% of the resin in 40%
of toluol by weight is recommended. The resins are clean, cheap,
water-white, inert, high-melting, absolutely neutral, and chemically
homogeneous. Clarite X undergoes a slight yellowing with age and
has a refractive index of 1.567 while Clarite is very color stable and
has a refractive index of 1.544.

The limited availability of certain reagents or enzyme substrates
may make it imperative to employ a considerably smaller volume
than is normally used in staining dishes and Coplin jars. The
hanging-drop technique (Glick and Fischer, 1945a) may be adopted
in these cases. The section on the slide is surrounded by a circle
of vaseline or stopcock grease, a drop of the reagent or substrate

14 MICROSCOPIC TECHNIQUES

solution is placed on the section, and to avoid evaporating when
prolonged contact between the liquid and the section is necessary,
a hanging-drop slide is placed over the section so that the drop is
enclosed by, but does not touch, the walls of the depression. The
two slides are bound together with a rubber band and carefully
inverted so that the section is covered by the hanging-drop.

B. INORGANIC ELEMENTS AND RADICALS

POTASSIUM

Macallum's original method for the histochemical detection of
potassium based on the precipitation of sodium potassium cobalti-
nitrite has been subject to modifications over the past thirty years.
The relatively recent modification of Gersh ( 1938) will be included
in the present work as well as the method of Carere-Comes ( 1938) ,
which depends on the development of an orange color with Siena
orange.

Gersh Modification of Macallum Method for Potassium

SPECIAL REAGENTS

Anhydrous Petroleum Ether freshly distilled over sodium {20-40°

b.p.).
Dried Paraffin {Grilbler, 50-52° ni.p.). Just before use heat at

100° or more in vacuo for about 15 min. or until bubbling stops.

12% Sodium Cobaltinitrite Solution. Dissolve 150 g. sodium nitrite

in 150 ml. hot water, cool to 40° (some crystals appear), add 50

g. cobalt nitrate crystals, while stirring rapidly add 50 ml. 50%

acetic acid in small portions, stopper, and shake well. Pass a

rapid stream of air through the solution and let stand overnight.

Siphon off clear hquid and filter. Add 200 ml. alcohol in small

portions to the filtrate with stirring. After a few hr. filter off

precipitate by suction. Wash precipitate four times with 25 ml,

portions of alcohol followed by two times with ether. Recrystal-

lize by dissolving each 10 g. solid in 15 ml. water and precipitate

with 35 ml. alcohol. Make up the 12% aqueous solution fresh

before use.

POTASSIUM 15

PROCEDURE

1. Subject tissue to freezing-drying (see page 3). ,

2. Transfer to paraffin, infiltrate in vacuo for 10-15 min. at
not more than 60°, and embed. Care should be taken to prevent
condensation of moisture on the dried tissue during the transfer
from the vacuum vessel to the paraffin.

3. Cut 15 fjL sections using no water or ice, mount near edges of
large chemically clean cover slips, press down with finger, melt
paraffin with a tiny flame, and press down again.

4. Remove paraffin by immersing the cover slips with the sections
in dry petroleum ether in a watch glass heated on a warm bar.
(Keep watch glass covered by another at all times except during
actual use. Replace the petroleum ether often.)

5. Remove from petroleum ether and burn off excess quickly.
Allow to cool.

From this point on, the test is carried out entirely in a cold room
the temperature of which should be 0° ± 1° during the manipula-
tions. All instruments and reagents are kept in this room. The
crystals of sodium potassium cobaltinitrite are relatively soluble at
room temperature, hence the precautions to maintain cold.

6. When cover slips with sections are cold, cover each section
with a drop of the sodium cobaltinitrite solution.

7. Drain off the solution and replace with glycerol.

8. Mount on clean slide with section between slide and cover
slip.

9. Examine under microscope with oil immersion lens. (Light
mineral oil is substituted for cedar oil since the latter is too viscous
at 0°.)

Result. Short yellow rods with rounded ends in a diffuse pale
yellow background are the crystals of sodium potassium cobalti-
nitrite.

Carere-Comes Siena Orange Method for Potassium

SPECIAL REAGENTS

Siena Orange Solution. Aqueous sodium p-dipicrylamine (prepared

ready for use by K. Hollborn & Sons) .
10% Hydrochloric acid, p

16 MICROSCOPIC TECHNIQUES

PROCEDURE

1. Fix tissue in neutral formalin and prepare paraffin sections
as usual.

2. Immerse deparaffinized sections in Siena orange soln. for 2
min.

3. Transfer to 10% hydrochloric acid for 3 min.

4. Wash twice in distilled water for 10 min., blot with filter
paper, and dry at 37°.

5. Mount in thickened cedar oil.

Result. Potassium is demonstrated by an orange color on a pale
yellow or colorless background. The author of this method has
failed to consider the effects of potassium diffusion when aqueous
solutions are employed for fixation, etc. Modification of the pro-
cedure to obviate this difficulty would be essential.

CALCIUM

A critical survey of histochemical tests for calcium was pre-
sented by Cameron (1930), who concluded that none of the tests
can be considered wholly specific. In all cases calcium must be
converted to an insoluble salt, if it is not already present as such,
and the insoluble compound is identified directly or it is made more
easily detectable by staining or conversion to a colored compound.
For visualization of calcium in the form of phosphate or carbonate
see page 78. In addition to these tests the Cretin (1924) gallic
acid color test has been extensively used, as has the formation of a
red precipitate by reaction of calcium salts with sodium alizarin
sulfonate (Pollack, 1928). For plant materials it is often sufficient
to produce and identify crystals of the oxalate, carbonate, or sulfate
(Lee's Vade Mecum, pages 293 and 668). The old test of von Kossa
(1901), depending on the reduction of silver salts under bright
light, has been championed by Gomori ( 1945a) . While this method
will demonstrate inorganic deposits in general, it can be considered
specific for calcium in bone or cartilage because the calcium salts
are the only ones present in significant amounts. An adaptation
of the von Kossa test to bone has been described by McLean and
Bloom«( 1940) and Bloom and Bloom ( 1940) .

CALCIUM 17

Calcium Sulfate Test for Calcium in Plant Tissue

SPECIAL REAGENTS

3% Sulfuric Acid.
40% Alcohol.

PROCEDURE

1. Fix tissue in acid-free alcohol or acid-free formalin.

2. Cut sections and bring them down to 40% alcohol.

3. Add 3% sulfuric acid to sections under cover slip.

4. Examine for colorless monoclinic needles of calcium sulfate.

Cretin Color Test for Calcium and Other Minerals

SPECIAL REAGENTS

Gallic Acid Reagent. Grind 0.1 g. trioxymethylene (metaformal-
dehyde) and 0.2 g. gallic acid in a mortar. Dissolve 0.25 g. of
the mixture in 5 ml. boiling distilled water and add 0.5 ml.
ammonium hydroxide (18° Baume, or 14% ammonia) ; stir until
the solution becomes straw colored. When this reagent turns
brown or rose, which it will in a short time, it can no longer be
used.

PROCEDURE

1. Prepare paraffin sections as usual. Remove the paraffin with
xylol and the xylol with chloroform.

2. After excess chloroform has been removed, add gallic acid
reagent. In 10-15 sec, drain off excess reagent and expose slide to
air,

3. Examine when color appears. An eosin counterstain may be
applied (Lison, 1936, page 76).

Result. Calcium gives a blue, barium a bright green, strontium
a water blue-green, cadmium a bluish-gi-een, magnesium a yellowish-
rose, iron a deep violet-brown, zinc and lead a dull yellow, and
silicon a pure yellow color.

Von Kossa Silver Test for Calcium

SPECIAL REAGENTS

5% Silver Nitrate.

18 MICROSCOPIC TECHNIQUES

PROCEDLUE

1. Transfer frozen or parafl5n sections which have been washed
with distilled water to the silver nitrate soln. in the dark for up to 1
hr. ; wash with distilled water in the dark, and expose to bright hght
for 30 min. or longer.

2. Wash well in distilled water, dehydrate, clear, and mount.
Result. Calcium salts are rendered black.

MAGNESIUM

A method for the demonstration of magnesium in plant cells was
developed by Broda ( 1939) . The principle of this method could be
adapted to studies on animal tissue. Most of the tests used for
calcium also give positive results for magnesium.

Broda Method for 3Iagnesiuin

SPECIAL REAGENTS

Quinalizarin Reagent. Triturate 100 mg. quinalizarin and 500 mg.

sodium acetate crystals, and dissolve 500 mg. of the mixture in

100 ml. of 5% sodium hydroxide.
0.2% Titian Yellow.
10% Sodium Hydroxide.
0.1% Azo Blue.

PROCEDURE

1. Prepare paraffin sections as usual.

2. Add 1-2 drops of quinalizarin reagent to a section on the slide,
followed by 1-2 drops of 10% sodium hydroxide.

3. To a different section add 1-2 drops of the Titian yellow
solution followed by 1-2 drops of 10% sodium hydroxide.

4. To another section add a drop or two of the azo blue dye.
Result. In the presence of magnesium the quinalizarin reagent

develops a blue color over several hours, the Titian yellow a brick-
red, and the azo d3'e a violet stain.

ZINC

Very little has been done in regard to the histological localization
of zinc and the procedure of Mendel and Bradley (1905) is still

ZINC AXD IBON 19

the sole method that has been developed. Zinc is precipitated by
nitropmsside and the precipitate is brought out as a deep purple
by treatment with sulfide.

Mendel and Bradley Alethod for Zinc

SPECI.\L REAGENTS

10% Sodium Xitroprusside.

Potassium Sulfide Solution. (Concentration not stated, but 1-5%
should suffice.)

PROCEDLTIE

1. Prepare paraffin sections. (!Mode of fixing tissue not given,
but, as in all other cases, the freezing-drying treatment, see page 3,
would be preferable.)

2. Treat sections with the nitropmsside soln. for 15 min. at 50^.

3. Cool, and wash in a stream of water for about 15 min.

4. Introduce under cover glass placed on section 1 drop of the
sulfide soln.

Result. Zinc elicits an intense purple color.

IRON

The classical histochemical tests for iron are the Prussian and
Turnbtiirs blue reactions and the hematoxylin method of Macallum.
the latter being the least specific (Lee's Vade Mecum, pages 2S9-
292). The Prussian blue test will detect ferric, and Turnbull's blue
ferrous, iron. More recently other methods have been proposed for
which certain advantages have been claimed. The dinitrosore-
sorcinol test of Humphrey (1935) brings out iron as a rich green
of pristine brilliance and the color is much more permanent than
that of Prussian or Tiu-nbull's blue, which fades after a year or two.
Thomas and Lavollay (1935) employed the 8-hydroxj-quinoline
reaction to ^-isualize iron in greenish-black; other metals appearing
in various shades of green and yellow. The strong flourescences of
metallic 8-hydrox^-quinolinates may also be used for identifications
(see page 108).

Iron, like many other metals, occurs in tissues both in the in-
organic or free form, and in the organic or bound form. Before
bound iron can be visualized it must be converted to the free form.

\

c K A R ^ ]

/ >

■^^-rlf

20 MICROSCOPIC TECHNIQUES

Macallum's (1908) technique is still employed for this conversion
and it consists of a treatment of deparaffinized sections with a solu-
tion of either nitric or sulfuric acid in alcohol. The iron liberated
is chiefly in the ferric form. In all tests special care must be taken
to protect tissues and fluids from dust. Iron will also appear in
the tests for lead and copper (see page 22).

Precautions to prevent diffusion of the iron in aqueous solutions
have not been sufficiently exercised in the following procedures.
The investigator should modify them accordingly.

Prussian Blue Test for Ferric Iron and TurnbuU's
Blue for Ferrous Iron

SPECIAL REAGENTS

Prussian Blue Reagent. 2% potassium ferrocyanide (use fresh

soln.).
TurnbuU's Blue Reagent. 2% potassium ferricyanide (use fresh

soln.).
Acid Alcohol. 1% hydrochloric acid in 70% alcohol.
Organic Iron Reagent. Equal vol. of 1.5% potassium ferrocyanide

and 0.5% hydrochloric acid (use fresh soln.).
Organic Iron Conversion Reagent. 3% nitric acid, or 4% sulfuric

acid, in 95% alcohol. (The sulfuric reagent acts more slowly.)

PROCEDURE FOR INORGANIC IRON

1. Fix in 95% alcohol for 24-48 hr.

2. Prepare paraffin sections as usual ( care must be taken to mini-
mize contact with iron — the microtome knife must be free of rust
and not freshly honed and glass needles should be substituted for the
steel ones) .

3. After removing paraffin and passing down to distilled water,
place sections in either the Prussian or TurnbuU's blue reagent for
3-15 min. (If both ferric and ferrous iron are to be visualized, use
a mixture of equal vol. of the two reagents.)

4. Wash in water containing eosin or safranin to counterstain.

5. Dehydrate, clear, and mount in benzol balsam.

PROCEDURE FOR ORGANIC IRON

1-2. Same as inorganic iron.

3. Liberate iron from the bound forms by treating deparaffinized

IRON 21

sections, brought down to water, with the conversion reagent for
24-36 hr. at 35°.

4. Wash in 90% alcohol followed by distilled water.

5. Place in the organic iron reagent for not over 5 min.
6—7. Same as 4-5 for inorganic iron.

Result. The iron appears blue.

Humphrey Dinitrosoresorcinol Test for Iron

SPECIAL REAGENTS

30% Ammonium Sulfide.

Saturated Aqueous Dinitrosoresorcinol or a 3% soln. in 50% alcohol,
(A few days aging improves the reagent; it is stable.)

PROCEDURE

1. Prepare formalin-fixed paraffin sections.

2. Remove paraffin; bring down to water, and place in the am-
monium sulfide solution for 1 min.

3. Rinse in distilled water and place in the dinitrosoresorcinol
reagent for 6-20 hr. depending on the depth of the brown background
desired.

4. Wash in water or dilute alcohol depending on whether an
aqueous or alcoholic reagent was used.

5. Pass through alcohols, carboxylol and xylol, and mount in
balsam.

Result. The iron appears bright dark green against a reddish or
rich brown background. For organic iron introduce steps 3-4 in
Prussian blue procedure ( see page 20) between steps 1 and 2 above.

Thomas and LavoUay Hydroxyquinoline Test for Iron

SPECIAL REAGENTS

Hydroxyquinoline Reagent. Dissolve 2.5 g. 8-hydroxyquinoline in
4 ml. glacial acetic acid with the aid of gentle warming. Quickly
add distilled water to bring volume to 100 ml. and filter the soln.

25% Ammonium Hydroxide.

PROCEDURE

1. Fix tissue in alcohol, neutral formalin, or trichloroacetic acid
soln.

2. Prepare frozen or paraffin sections as usual.

22 MICROSCOPIC TECHNIQUES

3. To the sections washed with neutral distilled water add a
few drops of the hydroxyquinoline reagent, and after 5-15 min. drain
off the liquid.

4. Add 1 drop of 25% ammonium hydroxide to form precipitate.

5. Wash in neutral distilled water. No large crystals should re-
main.

6. A lithium carmine nuclear stain may be applied.

7. Dehydrate with terpinol and mount in petrolatum, or examine
directly in neutral water.

Result. Iron appears as a greenish-black, calcium as a pale
yellow, magnesium as a straw-yellow, aluminum as a yellowish-
green, zinc and manganese as a yellow, and copper as a greenish-
yellow precipitate.

NICKEL

A method has been devised by Cretin and Pouyanne ( 1933) for
the histological demonstration of nickel in bone material by pre-
cipitation of nickel ammonium phosphate.

Cretin and Pouyanne Method for Nickel

SPECIAL REAGENTS

Fixative. Add 30 ml. formalin and 5 drops ammonium hydrosulfide

to 100 ml. physiological saline soln.
10% Ammonium Phosphate.

PROCEDURE

1. Fix tissue in the special fixative soln.

2. Transfer to the ammonium phosphate soln. in order to pre-
cipitate the insoluble nickel ammonium phosphate.

3. Decalcify and section.

4. Stain the nickel compound with alcoholic hematoxylin.

5. Wash, dehydrate, clear, and mount.

Result. Nickel will appear as a lilac deposit, or blue if present
in abundance.

LEAD AND COPPER

For many years the chromate method has been used for the micro-
chemical detection of lead in tissues ( Frankenberger, 1921; Cretin,

LEAD AND COPPER 23

1929) . This procedure depends on the formation of a yellow precipi-
tate of lead chromate when lead-bearing tissue is fixed in Regaud
fluid (20 ml. of 3% potassium dichromate plus 5 ml. formalin) . Lison
( 1936, page 101) has discussed this method as well as the test based
on precipitation of the sulfide, and rather favors the former. Oka-
moto and Utamura (1938) employed 2^-dimethylaminobenzylidene
rhodanine to produce a reddish-violet precipitate with copper in tis-
sues, a reaction given by gold, silver, and other metals (see pages
26, 28, and 29) .

Mallory and Parker (1939) described a method using hema-
toxylin and another employing methylene blue which would visual-
ize both lead and copper. The methylene blue technique was particu-
larly recommended for photomicrography of lead because of the in-
tense blue color developed.

In a study of the histological distribution of copper in the blowfly,
Waterhouse ( 1945) found that the only reagent which could be used,
of those tested, was sodium diethyl dithiocarbamate, which formed a
yellow product with copper. Waterhouse's technique was to drop a
0.1% aqueous solution on the fresh tissue followed by a drop of con-
centrated hydrochloric acid. The acid allowed greater penetration
of the reagent into the cells. Iron can interfere with this test by the
formation of a brown carbamate; however, the reagent can detect 1
part of copper in 100 million and its sensitivity to iron does not
approach this.

Mallory and Parker Hematoxylin Method for
Lead and Copper

SPECIAL REAGENTS * -

Hematoxylin Reagent., Dissolve 5-10 mg. hematoxylin in a few
drops of 95% or absolute alcohol and add 10 ml. freshly filtered
2% dibasic potassium phosphate.

PROCEDURE

1. Fix tissue in 95% or absolute alcohol (formalin may be used
for copper) .

2. Prepare celloidin sections as usual.

3. Stain sections for 2-3 hr. at 54°.

4. Wash in several changes of tap water 10 to 60 min.

24 MICROSCOPIC TECHNIQUES

5. Dehydrate in 95% alcohol, clear in terpinol, and mount in
terpinol balsam.

Result. Lead appears as light to dark grayish-blue and nuclei
as deep blue. Copper or hemofuscin pigment is brought out as an in-
tense blue. Inorganic iron or the pigment, hemosiderin, appears black
provided alcohol was used as the fixative and light to dark brown
if formalin was employed.

Mallory and Parker Methylene Blue
Method for Lead and Copper

SPECIAL REAGENTS

Methylene Blue Reagent. 0.1% of the dye in 20% alcohoL

PROCEDURE

L Fix tissue in Zenker fluid.

2. Prepare paraffin sections as usual, and apply a contrast stain
of phloxine if desired.

3. Treat sections for 10-20 min. with the methylene blue reagent
and decolorize in 95% alcohol for about the same time.

4. Dehydrate, clear, and mount as usual.

Result. Lead is colored intense blue. Copper or hemofuscin ap-
pears pale blue while iron pigment is not colored and hence appears
yellow to light brown. When pigment has both copper and iron it
develops a green color.

MERCURY

Three methods for the visualization of mercury in tissue sections
are given in Lison (1936, page 102). The mercury can be trans-
formed into the black sulfide, reduced by stannous chloride to give
the black free metal, or a violet precipitate can be formed with di-
phenylcarbazide. In addition to these, Okamoto's method for silver
(page 26) using p-dimethylaminobenzylidene rhodanine can be em-
ployed to give a reddish-violet precipitate with mercury.

After trials of the sulfide, diphenylcarbazide, and reduction
methods, Hand et at. ( 1943) favor the latter. They detected mercu-
rous mercury by reducing it to the metal by means of thioglycollic
acid, and the mercuric form was visualized by reducing with stannous
chloride.

MERCURY AND SILVER 25

Method of Hand et al. for Mercurous and Mercuric Mercury

SPECIAL REAGENTS

Mercurous Reagent. Combine 1 ml. thioglycollic acid with 9 ml.

glycerol.
Mercuric Reagent. Combine 5 g. stannous chloride, 5 g. tartaric

acid, and 100 ml. glycerol, and heat until clear. Stabilize by adding

a few grams of metallic tin to the final soln., which should be

stored in a stoppered bottle.
Iodine Reagent. Dissolve 50 g. potassium iodide in 50 ml. distilled

water; add 70 g. iodine and when it has dissolved, add 95% alcohol

to make 1 liter.
1% Chloroauric Acid. Store in a dark bottle.
Control Reagent. Add 5 g. tartaric acid to 100 ml. glycerol. Let
stand overnight to dissolve.

PROCEDURE

1. Prepare fresh frozen sections of tissue 15 /x thick.

2. Place sections on slides and allow to dry.

3. Cover each section with a drop of one of the reagents, depend-
ing on the test to be applied, fit on a cover slip, and blot away excess
reagent.

4. Seal edges of cover slip with commercial gold size (adhesive
used to hold gold foil on glass) .

5. After 10 min. examine under a microscope, comparing sections
with control reagent to those with other reagents. The sections
treated with the mercuric and control reagents remain unchanged
for at least 2 weeks.

Result. The metallic mercury formed in the tissue appears as
minute black spheres which may be dissolved by tincture of iodine,
or made to lose their glossy surface by forming gold amalgam on
treatment with chloroauric acid. In the test for mercurous mercury
characteristic yellowish crystals appear after about 5 min., in addi-
tion to the mercury globules, when mercuric mercury is also present.

SILVER

Particles of reduced silver in tissues may be made more intensely
black by treatment with dilute ammonium sulfide solution.

Okamoto, Utamura, and Akagi (1939) employed the p-dimethyl-

26 MICROSCOPIC TECHNIQUES

aminobenzylidene rhodanine reagent for the precipitation and vis-
ualization of silver in tissue sections. The fact that not only silver,
but also copper, gold, mercury, platinum, and palladium are like-
wise precipitated by this reagent offers little ground for concern
since these elements are not apt to exist in significant amounts in
tissues unless introduced experimentally or perhaps by accidental
poisoning. In these cases only one of the elements at a time is likely
to be present. However, certain differentiations can be made, if it is
assumed that more than one is present, on the basis of the varying
solubility behavior of the precipitated compounds. Thus divalent
copper reacts with the reagent only in neutral solution, whereas
monovalent copper and the other metals will react in either neutral
or acid solution. Furthermore the mercury precipitate is soluble in
dilute hydrochloric acid, the silver compound in potassium bromide
solution, and gold compound in potassium nitrite solution. Neutral
stannous chloride solution reduces the palladium precipitate to the
free metal which can then be converted to the chloride by means of
chlorine gas; this cannot be done with the platinum compound. Based
on these facts, possible separations have been suggested by Okamoto
et al.

Okamoto et al. Procedure for Silver

SPECIAL REAGENTS

Precipitation Reagent I. Add 3-5 ml. of saturated soln. of p-di-
methylaminobenzylidene rhodanine in absolute alcohol to 1-3 ml.
1 N nitric acid and 100 ml. distilled water.

Precipitation Reagent II. Combine 10-20 ml. of the saturated alco-
holic soln. of the rhodanine compound with 1-3 ml. 1 N nitric acid,
5-10 ml. 3% hydrogen peroxide, and 100 ml. distilled water.

Precipitation Reagent III. Add 2-5 ml. of the soln. of the rhoda-
nine derivative to 2 ml. 0.1 N hydrochloric acid, 3-5 ml. 1 N nitric
acid, and 100 ml. distilled water.

PROCEDURE

1. After fixing the tissue in absolute alcohol or neutral formalin,
prepare either celloidin, paraffin, or frozen sections.

2. Place the sections in precipitation reagent I for 24 hr. at 36°,
keeping the vessel closed.

SILVER AND GOLD 27

3. Rinse sections in distilled water and counterstain with hema-
toxylin.

4. Dehydrate, clear, and mount in balsam.

Result. Silver in the tissue is colored reddish-violet. The other
metals will produce shades of the same color.

Monovalent copper may be eliminated from visualization by
substituting precipitation reagent II for I, and mercury may be
similarly eliminated by employing reagent III. The silver precipitate
could be removed from the others by treating the colored sections
with 1% potassium bromide.

GOLD

Several methods have been used for the localization of gold in
tissues and Lison (1936. page 100) has discussed those of Christeller,
of Borchardt, and of Okkels. The first depends on treatment with
stannous chloride; the second employs silver nitrate followed by
nitric acid, and the last merely involves exposure to ultraviolet light
to obtain blackening of the gold granules. The more recent methods
of Roberts (1935), Okamoto, Akagi, and Mikami (1939), and Elft-
man and Elftman ( 1945) follow. The last-named method is probably
the best since it avoids the use of ions that might give rise to arti-
facts, and effects the bleaching of interfering pigments.

Roberts Method for Gold

SPECIAL REAGENTS

Sliver Nitrate Reagent. Just before using dissolve 2 g. pure silver
nitrate in 100 ml. 10% gum arabic soln. in the dark.

Hydroquinone Reagent. The day before using dissolve 1 g. pure
hydroquinone in 100 ml. 10% gum arabic soln.

5% Citric Acid.

5% Sodium Hyposulfite.

PROCEDURE

1. Fix tissue in Bouin fluid or 20% neutral formalin and wash
well with water.

2. Prepare paraffin or frozen sections.

3. Place sections for 5-10 min. in a fresh mixture of 2 ml. silver
nitrate reagent, 2 ml. hydroquinone reagent, and 1-3 drops of 5%

28 MICROSCOPIC TECHNIQUES

citric acid. Shake the mixture for 30 sec. immediately after prepar-
ing it.

4. Transfer sections rapidly to a 5% sodium hyposulfite soln. and
after a few min. wash thoroughly in water.

5. Dehydrate, clear, and mount.

Result. Granules of gold appear black due to a surface deposit
of silver.

Elftman and Elftman Method for Gold

SPECIAL REAGENTS

3 % Hydrogen Peroxide.

PROCEDURE

1. Fix tissue in neutral formalin, prepare paraffin sections, mount
with the aid of egg albumin, and then expose to formaldehyde vapor
for 1 hr. to increase the affixation.

2. After removing the paraffin and running down to water, place
in 3% hydrogen peroxide at 37° for at least 24 hr.; in most cases 3
days or longer gives the best results.

3. Do not stain the sections since staining may mask the gold
deposit. If staining is required, however, the interference is only
slight with light green SF (yellowish) or hematoxylin, and eosin
can be used when the gold is present as a sufficiently dense deposit.

4. Wash the sections in distilled water and dehydrate in the
usual manner.

5. Mount the sections in damar.

Result. Gold is made apparent by its presence in colloidal form.
The color of the deposit depends on the particle size, and accordingly
shades from rose to purple to blue and black are obtained. Usually
rose predominates.

Okamoto et al. Method for Gold

The procedure is the same as that in the Okamoto et al. method
for silver using precipitation reagent II (page 26). The gold ap-
pears as a reddish-violet or brownish-red precipitate. The removal

PLATINUM, PALLADIUM, AND URANIUM 29

of the colored silver precipitate from the sections, if silver was pres-
ent, can be carried out by treating with a saturated soln. of potas-
sium bromide for 1 hr. or more. If the sections are then washed in
distilled water and placed in 1% potassium nitrite for 24 hr. or
longer at 36° (or heated in the nitrite soln. for 1 min.) the gold
precipitate will dissolve leaving those of platinum or palladium,
should either of these be present.

PLATINUM

The Okamoto et al. method for silver may be employed unchanged
for the detection of platinum in tissues (see page 26).

PALLADIUM

The method of Okamoto, Mikami, and Nishida (1939) for the
visualization of palladium in sections of tissue follows the Okamoto
et al. silver method (p. 26) with 1 difference. Between steps 1 and
2 in the procedure, the following is introduced: treat the dry sections
with chlorine gas until the black palladium granules are made
colorless.

URANIUM

Two chemical tests have been presented for the localization of
uranium in tissue sections. Both are founded on the precipitation of
dark brown uranium ferrocyanide. Schneider ( 1903) was the first
to use this technique on the tissues of animals that had been in-
jected with uranium salts. Gerard and Cordier (1932) followed the
Prussian blue method for iron and reported good results. The latter
employed Bouin-Hollande or Carnoy fixatives and their coloring
reagent was 2% potassium ferrocyanide containing 2% hydrochlo-
ride acid. For details of the test, see the Prussian blue procedure for
iron, page 20.

The fluorescent properties of uranium salts subjected to ultra-
violet radiation can be utilized for the detection of these salts in in-
cinerated sections of tissue as indicated by Policard and Okkels
( 1930) ; see page 145.

ft.

30 MICROSCOPIC TECHNIQUES

ARSENIC

Castel ( 1934-1935a, 1936) developed two methods for the histo-
logical localization of arsenic. In one the tissue is fixed in an abso-
lute alcohol-chloroform-hydrochloric acid mixture saturated with
hydrogen sulfide, and the appearance of yellow granules was be-
lieved by Castel to be due to the formation of arsenous sulfide. A
reinvestigation of this technique by Tannenholz and Muir (1933)
led them to conclude that the yellow granules formed were not re-
lated to the presence of arsenic but were more likely composed of a
sulfur-protein complex.

The other method of Castel was based on the precipitation of
either cupric hydrogen arsenite (Scheele's green) or the cupric ace-
tate-cupric arsenite double salt ( Schweinf iirter green), and this pro-
cedure appears to be a reliable one.

Castel Cupric Salt Method for Arsenic

SPECIAL REAGENTS

Formalin-Copper Salt Reagent. Add 2.5 g. cupric sulfate or neutral
cupric acetate to 100 ml. metal-free 10% formalin (hydrogen sul-
fide is used to test for traces of metals in the formalin).

PROCEDURE

1. Place small pieces of tissue in the formalin-copper salt reagent
for 5 days.

2. Wash tissue in running water for 1 day.

3. Prepare paraffin sections as usual and examine after re-
moval of the paraffin.

Result. Green granules are indicative of arsenic.

BISMUTH

The histochemical detection of bismuth is founded on the reaction
of Leger ( 1888) , which is the precipitation of the double iodide of
bismuth and an alkaloid. Komaya (1925) and Christeller (1926)
employed the quinine salt for their tissue studies, and later Castel
( 1936) suggested the use of the brucine salt to avoid the interfer-
ence of iron which plagues the quinine method. He also modified
the earlier procedures by substituting sulfuric for nitric acid in the
reagents, Castel ( 1934-1935b), a change which enables the visual-

BISMUTH 31

ization of the bismuth as a mure reddish, rather than a yellowish-
orange deposit. More recently Wachstein and Zak ( 1946) employed
a modified Castel method in which the black sulfide, the form in
which bismuth appears in tissues, is converted to the white sulfate
by treatment with hydrogen peroxide and the sulfate is then trans-
formed to the brucine iodide salt.

In the procedure of Wachstein and Zak (1946), iron, which may
be present as the black sulfide, is oxidized to golden brown hemosid-
erin, which does not react wath Castel reagent, but which does give
the typical iron reactions. Wachstein and Zak pointed out that lead
sulfide would deposit in tissues in the same fashion as bismuth but
differentiation may be made by the fact that, after the lead sulfide
is converted to the sulfate by the peroxide, it will yield the slightly
yellowish lead iodide on treatment with Castel reagent in contrast
to the brilliant orange-red bismuth product. Similarly silver and
mercury will give yellow, and copper brown, iodides that can be
differentiated from the color of the bismuth precipitate. Melanin in
tissue is not bleached by the short treatment with peroxide and does
not react with Castel reagent. Wachstein and Zak emphasized that
melanin never impregnates capillary walls while bismuth does.

Wachstein and Zak Method for Bismuth

SPECIAL REAGENTS

Modified Castel Reagent. Dissolve 0.25 g. brucine sulfate in 100
ml. distilled water containing 2-3 drops cone, sulfuric acid. After
the brucine salt has dissolved add 2 g. potassium iodide. Store in
a brown bottle and filter before use.

Diluted Castel' s Reagent. Add 3 vol. distilled water to 1 vol.
reagent.

30% Hydrogen Peroxide. (Superoxol, Merck). Store in a refrig-
erator.

Levulose Solution. Dissolve 30 g. levulose in 20 ml. water by warm-
ing at 37'^ for 24 hr. and add a drop of the diluted Castel reagent.

Counterstain Solution. Add 1 ml. 1% aqueous light green SF
{Hartman-Leddon) to 100 ml. undiluted Castel reagent. Filter
before use.

PROCEDURE

1. Prepare either frozen or paraffin sections of formalin-fixed
tissue.

32 MICROSCOPIC TECHNIQUES

2. Treat the tissue on the slide for a few sec. with several drops
of 30% hydrogen peroxide to remove the black sulfide color.

3. Wash well in tap water and place in the Castel reagent for 1 hr.

4. Transfer to the diluted Castel reagent and shake gently to re-
move precipitates.

5. Remove most of the liquid from the slide by careful blotting
and mount in the levulose soln.

Result. Bismuth is indicated by the orange-red deposit. The
color may darken on standing. (If a counterstain is desired, stain
for 4 min. with the counterstain soln.)

Castel Method for Bismuth

SPECIAL REAGENTS

Bismuth Reagent. With the aid of warming, dissolve 1 g. brucine
in 100 ml. distilled water containing 3-4 drops of sulfuric acid
and add 2 g. potassium iodide. As an alternate reagent dissolve
1 g. brucine and 2 g. potassium iodide in 100 ml. of a mixture of
equal vol. alcohol and chloroform,

PROCEDURE

1. Fix the tissue in 10% formalin and prepare paraffin sections.

2. Treat deparaffinized sections for 15 min. with the bismuth
reagent and wash well in distilled water.

3. Mount in syrup of Apathy (heat equal parts of paraffin, m.p.
60°, and Canada balsam).

Result. Red granules are indicative of bismuth.

CHLORIDE AND PHOSPHATE-CARBONATE

Earlier methods for chloride, including Macallum's ( 1908) silver
test, were subject to the difficulty that, in the course of the manipu-
lations, a shift in the topographical distribution of chloride oc-
curred. Distortion due to this cause can be minimized by applying
the freezing-drying process to the tissue before further treatment.
Gersh ( 1938) makes use of this fact in his procedure, which enables
a differentiation between the chloride in the tissue and the phos-
phate and carbonate present. Phosphate and carbonate are visualized
together in this method. Two reagents are used, one permits visuali-
zation of chloride specifically by effecting the maximum precipita-

CHLORIDE AND PHOSPHATE-CARBONATE 33

tion of chloride in the presence of phosphate and carbonate by means
of the phosphoric acid it contains. The acid holds the phosphate in
solution and decomposes the carbonate. The other reagent, with-
out phosphoric acid, precipitates chloride, phosphate, and carbonate.
By comparing sections separately treated with each reagent, chloride
can be differentiated from phosphate and carbonate.

Gersh Method for Chloride and Phosphate-Carbonate

SPECIAL REAGENTS

Anhydrous Petroleum Ether freshly distilled over sodium {b.p.

20-40°).
Dried Paraffin ( GriXhler, m.p. 50-52°). Just before use heat at 100°

or more in vacuo for about 15 min. or until bubbling stops.
Silver Nitrate Reagent 1. To 60% silver nitrate solution add

enough cone, phosphoric acid to prevent precipitation of high

concentrations of phosphate, then saturate with silver chloride.

Filter and add 2-3 drops distilled water to each 10 ml. before

using.
Silver Nitrate Reagent 2. Saturate 60% silver nitrate solution with

silver phosphate and silver chloride. Filter and add water before

using as for reagent 1. Store both reagents in glass-stoppered

brown bottles in the dark.

PROCEDURE

1-5. These steps are identical with those in Gersh method for
potassium (see page 14).

6. Cover sections on one cover slip with reagent 1 and those on
another with reagent 2.

7. Drain off liquid from both cover slips and replace with a
drop of pure glycerol in each case.

8. Mount on clean slides with glycerol-covered sections down.

9. Expose both slides simultaneously to carbon arc radiation at
such distance as to avoid warming .

10. Examine microscopically at once by direct or dark-field il-
lumination. These preparations last only a short time. The highest
power to be used with the dark-field condenser is a 4 mm. high-dry
or 2 mm. oil immersion objective with a numerical aperture of 0.95.

Result. The reduced silver appears yellow to brown with or with-
out black or brown particles when viewed with direct illumination.

34 MICROSCOPIC TECHNIQUES

With the dark field the silver granules appear orange or rust colored.
Reagent 1 gives a test for chloride only. Reagent 2 gives a test for
chloride plus phosphate and carbonate.

IODIDE

A critical study of histochemical methods for the localization of
iodides in tissues was presented by Gersh and Stieglitz ( 1933) . After
a careful examination, these authors conclude that none of the pro-
posed methods is satisfactory. The difficulty is that any precipitat-
ing agent that might be used to fix iodide will also precipitate the
proteins and hence prevent its proper penetration into the tissue.

PHOSPHATE

Microscopic techniques for the detection of phosphorus in tissues
are usually based on reactions designed to visualize the phosphate
ion. Hence phosphate in organic combination must be liberated be-
fore it can be detected. Lison (1936, pages 113-1201 critically re-
viewed the various methods and considered Angeli's procedure to be
highly specific; this is a molybdate method using stannous chloride
as the reducing agent. Serra and Queiroz Lopes (1945) employed a
molybdate reaction with benzidine which they report to give a more
intense color than that developed by stannous chloride, and they
also use a more dilute acid medium, which is less damaging to the
tissue.

Johansen ( 1940, page 198) stated that phosphate may be identi-
fied in plant tissues by treating a section with a drop of solution pre-
pared by adding 25 ml. saturated magnesium sulfate and 2 ml. satu-
rated ammonium chloride to 15 ml. water. Crystals of magnesium
ammonium phosphate should form in the presence of phosphate. This
procedure is doubtlessly less advantageous than those previously
mentioned.

The method for visualization of enzymatically liberated phosphate
(page 78) may also be applied in certain instances.

Serra and Queiroz Lopes Modification of the Molybdate
Method for Phosphate

SPECIAL REAGENTS

Acetic Alcohol-Formalin Fixative. Add a few drops of glacial

<&

PHOSPHATE AND NITRATE 3o

acetic acid to 10 ml. of a mixture of 2 vol. 96% alcohol and 1 vol.

formalin.
Molybdate Solution. Dissolve 0.5 g. ammonium molybdate in 20

ml. distilled water, add 10 ml. cone. (30%) hydrochloric acid, and

dilute to 50 ml. with distilled water.
Acetic Benzidine Solution. Dissolve 25 mg. benzidine in 5 ml,

glacial acetic acid and dilute to 50 ml. with distilled water.
Saturated Sodium Acetate Solution.

PROCEDURE

1. Fix the tissue in the acetic alcohol-formalin mixture and wash
well in water.

2. In order to hydrolyze organic phosphates and precipitate the
free phosphate, treat small pieces of the tissue or frozen sections with
the molybdate reagent at 10-12° for 2-3 weeks, followed by 2-3
days at 20-25°. The temperature is kept low to prevent alteration of
the tissue and the rather long time is required to effect hydrolysis
with the relatively weak acid concentration of the reagent.

3. Cover the tissue with a drop of acetic benzidine soln. for 3
min. and then add 2 drops of the sodium acetate soln.

4. Mount in glycerol which has been stoi-ed with crystals of
sodium acetate in the bottle.

Result. An intense blue coloration characterizes phosphate.

note: Sena and Queiroz Lopez employ a digestion with nuclease to liberate
phosphate from nucleic acid. The visualization of this phosphate then serves
to indicate the nucleic acid.

NITRATE

Cramer ( 1940) developed a method for the histological demon-
stration of nitrate which is based on the doubly refractive property
in polarized light of the insoluble salt formed by the interaction of
nitrate with Nitron (4,5-dihydro-l,4-diphenyl-3,5-phenylimino-l,2,4-
triazole). Busch (1905) originally employed this reaction for the
gravimetric determination of nitric acid.

Cramer Method for Nitrate

SPECIAL REAGENTS

Nitron Reagent. 10% Nitron in 5% acetic acid.

36 MICROSCOPIC TECHNIQUES

PROCEDURE

1. Prepare frozen sections of fresh tissue.

2. Place 1-2 drops of hot Nitron reagent on a cover slip, and
place the cover slip over the section on a glass slide so that the tissue
is bathed in the liquid.

3. Put in a refrigerator for 30 min. to aid in the crystallization of
the nitrate.

4. Examine with polarized light under a microscope immediately
on removal from refrigerator.

Result. The doubly refractive zones are due to the insoluble
nitrate.

SULFHYDRYL AND DISULFIDE GROUPS

The earlier literature dealing with the application to tissue sec-
tions of the nitroprusside reaction for sulfhydryl groups, and the
reduction of disulfide compounds to give this test, has been
reviewed by Lison (1936, pages 133-136). The procedure given by
Pv,apkine and recommended by Lison, as well as the methods of
Bourne (1935) and of Hammett and Chapman (1938-1939), will
be given. The latter investigators critically examined the nitro-
prusside test and concluded that it should not be considered a
quantitative reaction; they established a well-defined procedure
which they believe most likely to yield satisfactory qualitative
results. However, the problem of diffusibility will no doubt limit,
or eliminate, the use of any nitroprusside method.

Nitroprusside Test of Rapkine

SPECIAL REAGENTS

5% Sodium Nitroprusside. For plant tissues.

2% Sodium Nitroprusside. For animal tissues.

Ammonium Sidjate Crystals.

Cone. Ammonium Hydroxide.

10% Potassium Cyanide.

10% Trichloroacetic Acid.

5% Zinc Acetate.

PROCEDURE

A. For free sulfhydryl groups

SULFHYDRYL AND DISULFIDE GROUPS 37

1. Immerse the fresh tissue in the zinc acetate soln. for a few sec.
This will stabilize the red color finally developed, as shown by
Giroud and Bulliard (1933).

2. Add 1 drop of the sodium nitroprusside soln. to a section
on a slide.

3. Add a crystal of ammonium sulfate and a drop of ammonium
hydroxide.

Result. Sulfhydryl compounds such as glutathione produce a
red color.

B. For total sulfhydryl groups

1. Treat sections of fresh tissue with 10% potassium cyanide
for 5-10 min.
2.-4. Proceed with steps 1-3 in A.

C. For protein-bound sulfhydryl groups

1. Treat sections of fresh tissue with 10% trichloroacetic acid
for 15 min. and wash thoroughly in water.
2.-4. Proceed with steps 1-3 in A.

note: The diffusibility of the sulfhydryl compounds formed in B, or liberated
in C, makes for particular unreUability in the localization of the groups in
the sections.

Bourne Nitroprusside Test

SPECIAL REAGENTS

5% Acetic Acid.

5% Sodium Nitroprusside Saturated with Ammonium Sidfate.

Concentrated Ammonium Hydroxide.

PROCEDURE

1. Place fresh frozen sections of tissue in hot 5% acetic acid
for 30-90 sec.

2. Pour off the acid and replace with nitroprusside-ammonium
sulfate soln. for 2 min.

3. Add a few drops of ammonium hydroxide and examine at once.
Result. A purplish-blue color indicates sulfhydryl compounds.

Hammett and Chapman Nitroprusside Test

SPECIAL REAGENTS

27-29% Ammonium Hydroxide.

38 MICROSCOPIC TECHNIQUES

1 % Sodium Nitroprusside.
Ammonium Sulfate Crystals.

PROCEDURE

1. Cover fresh tissue slice with 0.25 niL water.

2. Add 0.05 ml. ammonium hydroxide and then 0.05 ml. of the
nitroprusside soln.

3. Underline the tissue with 0.25 g. ammonium sulfate crystals
and examine at once.

C. ORGANIC SUBSTANCES AND GROUPS

LIPIDS AND CHOLESTEROL*

By means of staining methods, it is impossible to distinguish
with certainty between the various chemical types of the lipids
with the possible exception of cholesterol and its esters. Until
recently, the demonstration of lipids in general was usually carried
out with Sudan dyes which dissolve in the lipids and color them.
However, Jackson (1944) reported an improved method using
acetic-carbol-Sudan III which he claims should supersede all other
Sudan methods since it will bring out lipids that have been con-
sidered refractory to Sudan staining in the past. Jackson's paper
includes an enlightening critical survey of previous work. To cir-
cumvent the loss of small fat globules from the tissue when alcohol
or acetone dye solutions are used, Telford Govan (1944) employed
Sudan dyes suspended in aqueous media. The Kay and Whitehead
( 1935) procedure using Sudan IV, the newer Jackson ( 1944) method
employing acetic-carbol-Sudan III, and the Telford Govan (1944)
technique will be described. The staining of lipids by means of
fluorescent dyes according to Popper (page 105) would appear to
have some advantages, particularly in the use of the water-soluble
dyes such as Phosphine 3R.

Cholesterol and its esters may be visualized by the Liebermann-
Burchardt reaction as adapted for histological use by Schultz ( 1924-
1925) , Romieu ( 1927) , and Yamasaki ( 1931) . The Schultz procedure
has been employed more generally, and hence it will be given in
detail. Lison (1936, page 210) has pointed out that, though the
positive test is specific, a negative result does not necessarily

* See Bibliography Appendix, Ref. 3.

LIPIDS AND CHOLESTEROL 39

exclude the presence of cholesterol or its esters. The Windaus
digitonin test for free cholesterol (Lison, 1936, pages 211-212)
requires further investigation in the opinion of Kay and Whitehead
in Lee's Vade Mecum (1937, page 281). By means of the polarizing
microscope, cholesterol crystals can occasionally be observed in
sections as birefringent rhombic plates. If the temperature is low-
enough, neutral fats and fatty acids can also be observed in some
instances as birefringent crystals.

Kay and Whitehead Procedure for Sudan IV
Stain for Lipids

SPECIAL REAGENTS

Stock Solution of Dye (can be used for at least 6 months). Prepare
a saturated alcoholic solution by boiling 2 g. dye in 1 1. absolute
alcohol; allow to cool.

Staining Solution (good for only about 4 hr. after being mixed).
Add slowly, with stirring, to 7 vol. stock soln., 9 vol. of 45%
alcohol. Filter after standing for 1 hr. The 45% alcohol is prepared
by mixing 4 vol. absolute alcohol with 5 vol. distilled water.

PROCEDURE

1. Place formalin-fixed frozen sections in 50% alcohol for 5 min.
in staining soln. for 30 min. at 37° (turn sections over after 15 min.
for more even staining), in 50% alcohol several sec, and finally in
distilled water a few min.

2. Pass through filtered hemalum and wash in alkaline tap water
for several min.

3. Mount in glycerin jelly.

Result. The lipid will be stained red. -^^

The sections should be stained the day after cutting since they
tend to be sticky for a while just after the cutting. On the other
hand, poor results are often encountered if the staining is delayed
longer than one day after sectioning due, presumably, to crystalli-
zation of lipid material. The stain lasts for only a few months.

Jackson Procedure for Lipids Using Acetic-Carbol-Sudan III

SPECIAL REAGENTS

Sudan III Stock Solution. Cover 2 g. of the finely pulverized dye

40 MICROSCOPIC TECHNIQUES

with 450 ml. of 95% alcohol and heat on a water bath to simmer-
ing. Stir occasionally and then filter while hot. Transfer to a
stoppered bottle and place in a refrigerator overnight. Filter while
cold, and add distilled water dropwise from a burette, while stir-
ring, to reduce the alcohol concentration to 80%. Allow to stand
24 hr. ; filter and keep stoppered.

Acetic-Carbol-Siidan III Reagent. To a given volume of Sudan III
stock soln., slowly add 5% phenol dropwise from a burette, stir-
ring after each addition, to bring the alcohol concentration to 60%
{e.g., add 2 ml. phenol soln. to 6 ml. stock soln.). Let stand for
several hr. keeping the bottle stoppered. Then, in the same drop-
wise manner, add glacial acetic acid in the proportion of 2.5 drops
per ml. carbol-Sudan soln. After standing for 24 hr. in a stoppered
bottle, filter the reagent. Do not use when the soln. is more than
several days old.

5% Glacial Acetic Acid in 50% Alcohol.

PROCEDURE

1. Transfer formalin-fixed frozen sections to 50% alcohol for
1 min.

2. Place in the acetic-carbol-Sudan III reagent for 1.5 hr. or
longer; keep vessel stoppered.

3. Differentiate in the acetic-alcohol soln. for 10-60 sec. and
wash in distilled water for 1 min. In some cases it may be well to
dilute the acetic-alcohol with more 50% alcohol.

4. A counterstain of recently filtered Delafield hematoxylin
diluted with 2 vol. distilled water may be applied for 15 min.,
followed by differentiation in 0.5% hydrochloric acid until reddish
(10-20 sec), and treatment with very dilute ammonium hydroxide
for 5 min. to develop the blue color.

5. Wash in distilled water and mount in glycerin jelly.

Result. The lipid will be stained red.

Telford Govan Technique for Sudan Dye Staining
in Aqueous Media

SPECIAL REAGENTS

Sudan Dye Suspension. Add a saturated soln. of a Sudan dye in
acetone dropwise from a capillary pipette to 1% gelatine soln.

LIPIDS AND CHOLESTEROL 41

containing 1% acetic acid until a deep brick-red color and a
consistency of milk is obtained. Stir well during the addition.
Hold the mixture at 37° for 2 hr. to evaporate the acetone or let
stand in a warm room overnight. Filter off sediment through coarse
paper.
1 % Gelatin Solution.

PROCEDURE

1. Transfer frozen sections from water to 1% gelatin soln. for
2-3 min.

2. Stain for 30 min. at 37° with the suspension.

3. Wash sections in 1 % gelatin soln. for 2-3 min.

4. Wash well in water.

5. Counterstain, if desired, and mount in glycerin jelly or Karo
syrup.

Schultz Cholesterol Test

SPECIAL REAGENTS

2.5% Iron Alum (NH4)2S04.Fe2(S04).-5.24H20.

Concentrated Sulj uric-Glacial Acetic Acid Mixture. Add the sul-
furic acid slowly to an equal volume of glacial acetic acid, stirring
and cooling the while. (Only the purest acids are suitable and
the sulfuric must contain at least 98% sulfuric acid. The reagent
is hydroscopic and must be protected from atmospheric moisture.)

PROCEDURE

1. Place formalin-fixed frozen sections in the iron alum solution
for 3 days at 37°.

2. Rinse in distilled water, mount on slides and blot with filter
paper.

3. Add a few drops of the acid mixture and cover with a cover-
glass.

Result. A positive test for cholesterol, or its esters, is indicated
by the appearance of a blue-green color which reaches its maximum
intensity within a few min. Within 30 min. the sections acquire a
brown discoloration. The appearance of large numbers of bubbles
results from impure acids.

42 MICROSCOPIC TECHNIQUES

CAROTENE, CAROTENOIDS, AND VITAMIN A

The solubility of carotene, carotenoids, and vitamin A in organic
solvents makes it necessary to employ frozen sections of tissue for
histological studies on these substances. Unstained sections show
yellow, orange, or brown regions due to the presence of constituents
of this nature. The blue coloration given by concentrated sulfuric
acid with these compounds has been employed by Steiger (1941)
for the demonstration of carotene in leaves. The deep violet color
developed in the presence of aqueous 1% iodine in 7% potassium
iodide (Lison, 1936, page 245) is also characteristic of these
polyenes, and when treated with oxidizing agents, such as chromic
acid, they are bleached. Bourne (1935) adapted the Carr-Price
reaction to tissue sections by placing frozen sections directly into
a chloroform solution of antimony trichloride. It is well known
that the blue color due to vitamin A fades very rapidly, while that
due to carotene persists. As shown by Raoul and Meunier (1939),
sterols produce a red color in the Carr-Price test. The detection uf
vitamin A in tissue by fluorescence is described on page 104.

Steiger Method for Carotene in Leaves
SPECIAL REAGENTS

Alkali-Alcohol Mixture. Combine 1 vol. saturated potassium hy-
droxide with 2 vol. of 40% alcohol and 3 vol. tap water.
Concentrated Sulfuric Acid.

PROCEDURE

1. Place green leaves in the alkali-alcohol mixture in a wide-
mouth bottle and seal the glass stopper with vaseline.

2. After several days in the dark, when the fluid is green and
the tissue yellow, transfer to distilled water for several hr.

3. Place small pieces of tissue on a slide and dry with filter paper.

4. Add 1 drop cone, sulfuric acid.

Result. Carotene is indicated by the appearance of dark blue
crystals visible under the microscope. Grossly, a green color chang-
ing to blue can be observed.

RIBOFLAVIN AND POLYSACCHARIDES 43

RIBOFLAVIN

The detection of riboflavin in tissue sections, based on reduction
of the vitamin in acid medium to leucoflavin and reoxidation to
bright red granules of rhodoflavin, was employed by Chevremont
and Comhaire ( 1939) . Riboflavin can be recognized in tissue by its
characteristic greenish-yellow fluorescence when irradiated with
ultraviolet (page 104).

Chevremont and Comhaire Method for Rihoflavin

SPECIAL REAGENTS

Fixative. Formol-Nitron or formol-basic lead acetate soln.
Reductant Solution. Add zinc to hydrochloric acid to generate

hydrogen.
Oxidant Solution. Dilute hydrogen peroxide.

PROCEDURE

1. Fix tissue for 5 days and prepare sections.

2. Treat sections for 30 min. with reductant soln.

3. Ptinse in water and treat with oxidant soln.

4. Examine under microscope.

Result. Bright red granules indicate presence of riboflavin. The
localizations are probably unreliable due to the diffusibility of the
riboflavin and its derivatives.

POLYSACCHARIDES IN GENERAL

A general reaction for the microscopic visualization of polysac-
charides has been described by Hotchkiss (1946).* The reaction
involves the oxidation of adjacent hydroxyl groups to aldehydes by
means of periodate, and the coloring of the aldehyde with Feulgen
reagent. The oxidation takes place according to the equation:

— CHOH— CHOH— + HsIOe > — CHOHCO— + HIO3 + 3 H2O

The chief substances in plant tissues that show the stain are
starches, cellulose, hemicelluloses, and pectins and, in animal tissues,
glycogen, mucin, mucoproteins, and presumably hyaluronic acid
and chitin. The pentoses of nucleic acid are so substituted that they

* Subsequent to this writing the author learned of the paper of McManus
(1946) (Bibliography Appendix, Ref. 13) in which the same principal was
independently presented.

44 MICROSCOPIC TECHNIQUES

will not give the reaction and cerebrosides, if present, would be
expected to react.

Method of Hotchkiss for Polysaccharides

SPECIAL REAGENTS

Periodic Acid Solution A. Dissolve 400 mg. periodic acid (H5IO6,
obtainable from G. Frederick Smith Chemical Co.) in 10 ml. dis-
tilled water, add 5 ml. M/5 sodium acetate and 35 ml. alcohol.
^Periodic Acid Solution B. Dissolve 400 mg. periodic acid in 45 ml.
distilled water and add 5 ml. ilf/5 sodium acetate.

lodide-Thiosidfate Solution. Dissolve 1 g. potassium iodide and

1 g. sodium thiosulfate (Na2S203.5H20) in 20 ml. distilled water
and add with stirring 30 ml. alcohol followed by 0.5 ml. 2 N
hydrochloric acid. A sulfur precipitate forms and settles out
slowly although the soln. may be used immediately.

Feulgen Reagent. See page 67.

Sulfite Wash Solution. Add 0.5 ml. cone, hydrochloric acid and

2 ml. 10% potassium metabisulfite to 50 ml. distilled water.

PROCEDURE

1. If water-soluble polysaccharides are to be stained, fix the
tissue in a dehydrating soln. such as Carnoy fluid (page 45),
Otherwise, use any of the usual fixatives. After fixation remove
traces of mercury, if present, with iodine and be sure any formalde-
hyde is completely removed by washing. (70% alcohol may be used
as a washing soln. if it is desired to avoid removal of water-soluble
polysaccharides.)

2. Place section or smear in the periodic acid soln. A or B
(depending on whether an alcoholic or aqueous soln. is desired)
for 5 min.

3. Flood with 70% alcohol (or water) and transfer to the iodide-
thiosulfate soln. for 5 min.

4. Again wash with 70% alcohol (or water) and then transfer
to the Feulgen reagent for 15-45 min.

5. Rinse with the sulfite wash soln., dehydrate, and mount as
usual. ^

^ Result. Polysaccharides are indicated by the violet fuchsin color.

note: If free tissue aldehydes are present it is necessary to remove them
first as on page 93.

ACID POLYSACCHARIDES 45

If the periodate and iodate are not washed out they will give rise to a
brownish coloration in the Feulgen reagent.

Control sections or smears may be made by placing in 70% alcohol (or
water), instead of in the periodic acid soln. A or B, and then carrying
through the remaining steps without change.

Egg albumin adhesive may take a slight stain due to the carbohydrate
content of this material.

If neutral polysaccharides are to be stained, a counterstain with a basic
dye (such as 0.02 mg. malachite green per ml. water) may be used. If
mucin or acid polysaccharides are to be stained, the counterstain should be
an acid dye.

ACID POLYSACCHARIDES — HYALURONIC ACID*

For the demonstration of acid polysaccharides of the hyaluronic
acid type Hale (1946) employed fixation in a dehydrating medium
to prevent solution of the water-soluble acid polysaccharide, com-
bination of the latter with iron, and demonstration of the iron by
means of the Prussian blue reaction. The iron will not combine with
neutral polysaccharides or proteins according to Hale. In order to
differentiate between hyaluronic acid and other substances which
might give the blue stain, Hale suggests that hyaluronidase be
used to digest away the hyaluronic acid in the section on one slide
and a comparison be made to an undigested section.

Hale Method for Acid Polysaccharides

SPECIAL REAGENTS

Acetic-Iron Solution. Mix equal vol. dialyzed ferric hydroxide,
concentration not stated (Hale used the product of the British
Drug Houses Ltd.), and 2 M acetic acid.

0.02 M Potassium Ferrocyanide in 0.14 M Hydrochloric Acid.

PROCEDURE

1. Fix 3-4 mm. pieces of tissue in Carnoy fluid (6 vol. absolute
alcohol, 3 vol. chloroform, and 1 vol. glacial acetic acid) for 0.5 hr.

2. Treat with absolute alcohol; clear, and mount in paraffin.

3. Prepare sections and place on slides without albumin adhesive.

4. Flood the deparaffinized sections with the acetic-iron soln.,
and after 10 min. wash well with distilled water.

5. Treat the sections with the ferrocyanide soln. for 10 min. and

* See page 46 for the staining of mucoproteins.

46 MICROSCOPIC TECHNIQUES

wash with water and counterstain if desired (it is well to use a red
eounterstain such as f uchsin) .

6. Rapidly dehydrate, clear in xylol, and mount in Canada
balsam.

Result. Acid polysaccharide is indicated by the blue color.

MUCOPROTEINS*

Toluidene blue will stain quite a variety of acid substances, but
the metachromatic staining by this dye of mucoid compounds
containing polysaccharide esters of sulfuric acid is specific for
these compovmds, provided that the method of Lison (1935) is
strictly adhered to (Sylven, 1941, 1945). Sylven (1941) has made a
thorough study of the staining and he emphasized that "false"
metachromatic staining can be obviated by the prompt removal, of
water by alcohol after the staining, the alcohol assuring a ''true"
reaction which is the red stain characteristic of, and specific for,
the polysaccharide sulfates in tissue. Holmgren and Wilander ( 1937)
found that basic lead acetate solution was a superior fixative for
tissues to be subjected to the metachromatic toluidene blue stain,
but Sylven now employs a mixture of this fixative with formalin to
reduce the time required for the fixation. The staining time can be
reduced by aging the dye solution, and the greater the alcohol
concentration in the dye solution the paler the resulting stain will
be. In order to bring out mast cell granules properly, the dye is
made up in alcohol of a concentration of 30% or higher (Sylven).

According to the claim of Hempelmann ( 1940) , chondroitin and
mucoitin sulfuric acid proteins can be differentiated from one
another in histological preparations by means of the toluidene blue
stain. In a dilution of 1 : 1,280,000 an aqueous solution of toluidene
blue is supposed to stain the chondroitin material in paraffin sections
a violet-red color, while the mucoitin protein complex remains un-
stained. Differentiation is also claimed when the dye is used in a
1 : 410,000 dilution in a solution of 10 vol. alcohol and 45 vol. water.
The alcohol concentration is stated to be critical, presumably both
mucoproteins will be stained if the proportion of alcohol is less,
and neither if it is greater. No confirmation of these claims has been
made; in fact, to the writer's knowledge several attempts to do so
have failed.

* See Bibliography Appendix, Refs. 8 and 10.

MUCOPROTEINS, GLYCOGEN, AND MUCIN 47

A fundamental study of metachromasy of basic dyes has been
published by Miehaelis and Granick (1945).

See page 45 for another method of staining acid polysaccharides,
and page 50 for the staining of mucin.

Lison Method for Polysaccharide Sulfate Compounds
(after Sylven)

SPECIAL REAGENTS

Fixing Solution. Mix equal vol. of 8% basic lead acetate soln.

and 14-16% formalin.
Toluidene Blue Solution. Prepare separately (a) 0.1% dye in 1%

alcohol and (6) 0.1% dye in 30% alcohol, and let stand for a

number of days to age.

PROCEDURE

1. Fix the tissue for 12-24 hr. in the fixing soln.

2. Prepare paraffin sections in the usual manner.

3. Stain the sections for 30 min. using soln. a on some, and soln.
b on others. Soln. a gives a more intense stain.

4. Wash well in alcohol briefly, immediately after removing from
the dye soln.

5. Mount in natural cedar oil.

GLYCOGEN AND MUCIN*

A critical comparison of the iodine, Best carmine, and Bauer-
Feulgen methods for demonstrating glycogen microscopically was
,made by Bensley (1939), who concluded that the Bauer-Feulgen
method, which depends on the reaction of the aldehyde groups in
the carbohydrate with the reagent, is by far the best if the tissue
is promptly fixed in alcohol-formalin solution. When chrome salts
are present in the fixative, the visualization of intracellular glycogen
was found to require the Best carmine stain since the Bauer-Feulgen
method is not specific in those cases, and the iodine technique is
not suited for high-power studies. A procedure for preparing paraffin
sections for the carmine stain was given by Mullen (1944), who
employed celloidin to hold the deparaffinized sections to the glass
slide.
Mitchell and Wislocki ( 1944) reported that the ammoniacal silver

* See Bibliography Appendix, Refs. 2 and 12.

48 MICROSCOPIC TECHNIQUES

nitrate method which Pap (1929) employed for the staining of
reticulum visualized glycogen more intensely and consistently than
either the Best carmine or Bauer-Feulgen procedures. The admitted
drawback to this method is the fact that since reticulum fibers of
connective tissue are also stained it cannot be applied to this tissue.
However, the authors feel that in other cases the ammoniacal silver
nitrate method has advantages over those previously employed.
Gomori (1946) subsequently modified this procedure and developed
a more selective method which demonstrates glycogen and mucin,
but eliminates possible interference by desoxyribonucleic acid, uric
acid, and granules of enterochromaffin cells, all of which can reduce
silver solutions under certain conditions. However, melanin is stained,
and except for this the method enables the same localizations of the
reducing substances as the Bauer-Feulgen procedure. Should in-
soluble calcium salts be present they too will stain black. A prelim-
inary 10 min. treatment of the sections with citrate buffer of pH
3-4 will remove calcium deposits.

The differentiation of glycogen from other substances that give
positive reactions may be made in some instances by employing
saliva to digest away the glycogen selectively. See page 46 foi* the
staining of mucoproteins.

Bauer-Feulgen Stain for Glycogen (after Bensley)

SPECIAL REAGENTS

Alcohol-Formalin Fixative. 9 vol. absolute alcohol plus 1 vol.
neutral formalin. The alcohol may be first saturated with picric
acid.

Feulgen Reagent* Heat to dissolve 1 g. basic fuchsin in 100 ml.
distilled water. Filter while warm, cool, add 20 ml. 1 A^" hydro-
chloric acid and 1 g. sodium bisulfite. Let stand 24 hr. The soln.
should be straw colored.

1% or 4% Chromic Acid.

Bisulfite Rinsing Solution. 1 vol., 1 M sodium bisulfite plus 19
vol. tap water.

PROCEDURE

1. Fix very small pieces (2-3 mm.) of fresh tissue in the alcohol-

*See pages 65 and 67 for other methods of preparing this reagent.

r

GLYCOGEN AND MUCIN 49

formol solution for 24 hr. (Deane, Nesbett, and Hastings, 1946,
recommend the use of ice-cold alcohol-picric acid-formalin to pre-
serve the glycogen throughout the tissue block.) Wash in absolute
alcohol and embed in paraffin, being careful to prevent overheating.
(It is essential that very fresh tissue be used since glycogen is
rapidly autoiyzed. The Altmann-Gersh freezing and drying tech-
nique for fixation may also be used; in fact it can lead to a truer
picture of the glycogen distribution, as shown by Bensley and
Gersh, 1933a) .

2. Section, mount on slides, and deparaffinize as usual.

3. Place in 4% chromic acid 1 hr. or in the 1% soln. overnight.

4. Wash in running water for 5 min., place in Feulgen reagent
10-15 min., rinse with three changes of bisulfite soln. for 1.5 min. in
each change, and wash in running water for 10 min.

5. Nuclei may be counterstained with hematoxylin.

6. Dehydrate, clear, and mount in balsam.

7. As a negative control, remove the glycogen from some of the
sections, brought down to water, by adding fresh saliva. During a
15-30 min. period, change the saliva several times. Wash with water
at 37° to remove mucus and stain as above beginning with step 3.
Comparison of these sections with those not given the saliva treat-
ment helps to distinguish the glycogen regions.

Result. The glycogen appears deep red-violet, the nuclei laven-
der.

Best Carmine Stain for Glycogen (after Bensley)

SPECIAL REAGENTS

Alcohol-Formalin Fixative. Same as the reagent for Bauer-Feulgen
stain.

Carmine Stain Stock Solution. Gently boil 2 g. carmine, 1 g. potas-
sium carbonate, and 5 g. potassium chloride in 60 ml. distilled
water until color darkens. After cooling, add 20 ml. cone, am-
monia and let stand 24 hr. This solution may deteriorate in a
month in a warm room ; keep well stoppered.

Fresh Carmine Stain. 10 ml. stock soln., 15 ml. cone, ammonia, and
30 ml. methanol (C.P.).

50 ' MICROSCOPIC TECHNIQUES

PROCEDURE

1. Follow steps 1 and 2 for Bauer-Feulgen stain.

2. After bringing down to distilled water, stain nuclei with
hematoxylin.

3. Transfer to fresh carmine stain and after 20 min. wash in
three changes of methanol, dehydrate in acetone, clear in toluol,
and mount in balsam.

4. Run negative control sections as in step 7 for the Bauer-
Feulgen method but apply the carmine stain above.

Result. The glycogen will appear brilliantly red.

Gomori Procedure for Glycogen and Mucin

SPECIAL REAGENTS

Fixative. One of the alcohol fixatives such as alcohol-picric acid-
formalin (page 48) for glycogen. Any routine fixative for mucin.

0.5% Collodion in Alcohol-Ether Solution.

5% Chromic Acid.

1-2% Sodium Bisulfite.

Silver-Methenamine Stock Solution. Add 5 ml. 5% silver nitrate
soln. to 100 ml. 3% methenamine ( hexamethylenetetramine ) soln.
Shake until the initial heavy white precipitate disappears, and
store in refrigerator.

Alkalized Silver-Methenamine Solution. To 25 ml. silver-methena-
mine stock soln. add 25 ml. distilled water and 1-2 ml. 5% borax
(Na2B4O7.10H2O).

0.1 % Gold Chloride.

2—3% Sodium Hyposulflte.

PROCEDURE

1. Fix tissue and prepare paraffin sections as usual.

2. Run sections through xylol, alcohols, and water. (For glyco-
gen, protect sections on slides by dipping into collodion soln. before
transferring to the final alcohol soln.)

3. Place slides in 5% chromic acid for 1 — 1.5 hr.

4. Wash in running tap water for 10 min. and treat with the
bisulfite soln. for 1 min. to remove remaining traces of chromic acid.

5. Wash in running tap water for 5 min., rinse in distilled water,
and incubate at 37-45° in the alkalized silver-methenamine soln.
Examine sections under microscope every 15 min. Staining is com-

GLYCOGEN, MUCIN, AND STARCH 51

pleted when the glycogen 'and mucin appear deep brown or black.
The background will be yellowish. Usually 1-3 hr. is required.

6. Rinse well in repeated changes of distilled water and tone in
the gold chloride soln. for 5 min.

7. Rinse in distilled water and then in the hyposulfite soln. to
remove unreacted silver.

8. Wash in tap water and counterstain if desired.

9. Mount as usual.

Result. Glycogen and mucin will appear in shades from grey-
brown to black on an unstained background. Sometimes the collodion
film becomes stained and it can be removed by acetone or alcohol-
ether.

STARCH

The common practice of employing a dilute iodine solution to
develop a blue color with starch can be applied to sections of plant
material, as can the crystal violet stain followed by washing with
saturated picric acid solution. The use of formaldehyde as a swelling
agent to obtain special effects with safranine 0 and fast green
FCF was described by Bates (1942). Starch granules can also be
recognized by the characteristic black crosses they exhibit due to
their doubly refractive pioperties when viewed under the micro-
scope with polarized light.

The following procedure of Milovidov (1928) is well suited for
the preparation of permanently mounted sections stained for starch.

Milovidov Method for Starch

SPECIAL REAGENTS

Aniline Fuchsin Stain.

5% Alcoholic Aurantia.

2% Tannin.

1 % Toluidine Blue, Gentian Violet, or Methyl Green.

PROCEDURE

1. Fix plant tissue in Regaud fluid and prepare sections as usual.

2. Stain sections with aniline fuchsin for 5 min. and differentiate
in the aurantia soln.

52 MICROSCOPIC TECHNIQUES

3. After washing sections in distilled water, mordant for 20 rain,
in the tannin soln. and again wash.

4. Stain sections in either toluidine blue, gentian violet, or
methyl green for 5-10 min.

5. Differentiate in 95% alcohol, dehydrate in absolute alcohol,
clear in xylol, and mount in balsam.

Result. The starch will appear as either blue, violet, or green
granules depending on which of the stains was used in step 4. The
mitochondria will appear red.

CELLULOSE

Post and Laudermilk (1942) Iodine Stain for Cellulose

SPECIAL REAGENTS

Iodine Solution. 20 ml. 2% iodine in 5% potassium iodide, 180

ml. distilled water, and 0.5 ml. glycerol.
Lithium Chloride Solution. Saturate 15 ml. distilled water at 80°,

cool; use supernatant soln.

PROCEDURE

1. Tease out sections or fibers.

2. Apply 2-3 drops of the iodine soln. and after 10 sec. blot
with filter paper and dry.

3. Add a drop of the lithium chloride soln., cover, and examine.

Result. Cellulose appears in the following colors depending on
its source :

Typical color Fiber

Light blue cotton, soda pulp, bleached sulfite, straw,

esparto

Dark blue pineapple fiber

Greenish blue linen

Green to yellowish green ...sisal, Manila hemp, yucca

Yellow yucca, ground wood, hemp, Manila hemp

Lemon yellow kapok

Brownish yellow \nio

CHITIN*

The horny carbohydrate material, chitin, requires special treat-
ment to soften it sufficiently for the preparation of paraffin sections.

* See Bibliography Appendix, Ref . 16.

CELLULOSE AND CHITIN 53

The most recent method for this treatment is that of Murray ( 1937) ,
but the Diaphanol technique has been widely employed. Once sec-
tions are prepared they may be stained by the procedure of Zander,
Schulze, or Bethe given in Lee ( 1937, page 600j .

Murray Method for Softening Chitin

SPECIAL REAGENTS

Formalin- Saline Fixative. 10% formalin in 0.8% sodium chloride
soln.

Dehydrating Fixative. Equal vol. absolute alcohol, chloroform,
and glacial acetic acid to which mercuric chloride is added to
saturation ( about 4% ) .

Chloral Hydrate-Phenol Reagent. Equal weights of chloral hy-
drate and phenol warmed together until they blend to an oily
liquid that is fluid at room temperature.

PROCEDURE

1. Fix material in the formalin-saline soln.

2. Transfer to the dehydrating fixative.

3. Place specimen in the chloral hydrate-phenol reagent for 12-
24 hr. or longer.

4. Clear with xylol, chloroform, or carbon disufide and imbed
in paraflfin.

Diaphanol Method for Softening Chitin

SPECIAL REAGENTS

Diaphanol Solution. Pass vapors of chlorine dioxide into ice-cold
50% acetic acid. Store in a cool dark place in a glass-stoppered
bottle. (Before the war, Diaphanol was sold by Leitz; and Lee —
1937, page 598 — recommends buying, rather than preparing, the
soln. However, it will probably be impossible to buy for some
time, and there is no reason why the reagent cannot be safely
prepared if the obvious precautions of working in a hood, etc.,
are taken.)

PROCEDURE

1. Fixed material is rinsed in 63% alcohol and placed in Di-
aphanol in a glass-stoppered bottle in diffuse daylight until bleached

54 MICROSCOPIC TECHNIQUES

and softened. The specimen should be pierced or cut to allow escape
of carbon dioxide.

2. If the Diaphanol becomes discolored, transfer to a fresh por-
tion of the soln.

3. Place in 63% alcohol until hardened and then pass through
tetralin into paraffin.

Methods for Staining Chitin

The softened material, or sections of it, may be tested for chitin
by a variety of color reactions. Zander treated for a short time with
a drop of fresh iodine in potassium iodide soln., followed by a drop
of strong zinc chloride soln. Upon removal of the reagents with
water, a violet color is obtained in the presence of chitin. Schulze
divided the material into two portions. One was subjected to the
procedure of Zander and the other was treated with iodine and then
cone, sulfuric acid. The latter test serves to distinguish chitin from
cellulose and tunicin since chitin yields a brown color while the
others give a blue. Bethe employed freshly prepared 10% aniline
hydrochloride containing a drop of cone, hydrochloric acid for each
10 ml. After sections were placed in this soln. for 3-4 min., they
were rinsed with water and the slides were then placed, sections
downward, in a bath of 10% potassium dichromate. Chitin produces
a green coloration which becomes blue in tap water or ammoniacal
alcohol.

ASCORBIC ACID

The stain for ascorbic acid was developed in 1933 by Bourne,
who utilized the fact that reduced silver is deposited when ascorbic
acid in tissue interacts with acid silver nitrate. Bourne (1936)
published a critical survey of this stain and his recommended pro-
cedure is given below with a subsequent modification by Barnett
and Bourne (1941) designed to increase the specificity of the test
by dissolving precipitated silver salts in dilute ammonia. Giroud
and Leblond (1936) also investigated the technique and its appli-
cations, and in reply to criticisms of the specificity of the stain,
these authors ( 1937) point out that the positive test is specific for
ascorbic acid but a negative result does not necessarily mean that

ASCORBIC ACID 55

ascorbic acid is absent. Tonutti (1938) washed the tissue in 5.4%
levulose solution before staining in order to remove blood. The
reliability of the localizations obtained with the silver stain remains
to be proved, according to Danielli ( 1946a) . It would be necessary
to establish that the ascorbic acid is attached to a nondiffusible
body and that the reaction product could not diffuse, or that the
ascorbic acid site has a high affinity for the reaction product.

Bourne Silver Stain for Ascorbic Acid

/. Reduced Ascorbic Acid
SPECIAL REAGENTS

Acid Silver Nitrate. Add 5 ml. glacial acetic acid to 100 ml. 5%

silver nitrate.
5% Ammonium Hydroxide.

PROCEDURE

1. Place frozen sections of fresh tissue in the acid silver nitrate
soln. for a few minutes, and then treat with 5% ammonium
hydroxide.

2. Wash with distilled water. If desired, lipids can be then
stained with a Sudan dye in 90% alcohol.

3. After clearing, mount in glycerin.

Result. Granules containing reduced ascorbic acid appear black.

11. Reduced and Oxidized Ascorbic Acid
PROCEDURE

1. Expose the fresh tissue to the vapor of glacial acetic acid for
several minutes.

2. Cut into thin pieces and subject to an atmosphere of hydrogen
sulfide for 15 min. in order to reduce the oxidized form.

3. Remove hydrogen sulfide by placing in a vacuum for 10-30
min. followed by a good stream of nitrogen gas for 15 min.

4. Treat with acid silver nitrate soln. followed by ammonium
hydroxide as above.

Should glutathione be present in quantities sufficient to inhibit the
test, wash the tissue momentarily after the hydrogen sulfide treat-
ment and immerse at once into a mercuric acetate soln. for a few
minutes. After washing, apply the acid silver nitrate solution and
then the ammonium hydroxide.

56 MICROSCOPIC TECHNIQUES

PROTEIN REACTIONS

Many of the tests for proteins are poorly adapted to histochemical
work because the strong acid or alkaU that they require has too
great a disintegrative eifect on the cellular structure. Tests which
particularly fall into this group are the biuret reaction for com-
ponent peptides, the xanthoproteic reaction for phenolic constitu-
ents, Millon's tyrosine reaction, Romieu's tryptophane test, the
tryptophane reaction of Voisenet-Flirth, and the diazo reaction for
histidine and tyrosine. Serra's arginine test, which is much less
drastic, and Berg's ninhydrin reaction for a-amino acid groups,
which uses no corrosive reagents although heating is required, will
both be described as well as two of the previous group, Millon and
Romieu reactions.

ARGININE AND ARGININE-CONTAINING PROTEINS

In another of those coincidences that occasionally turn up, Serra
at the University of Coimbra, Portugal, and Thomas at the Univer-
sity of Missouri, independently, and without knowledge of the other's
work, adapted the Sakaguchi (1925) reaction for arginine to its
histochemical identification. The reaction is based on the develop-
ment of an orange-red color with arginine when a-naphthol and
hypobromite or hypochlorite react with it in an alkaline medium.

The first description of the method by Serra (1944a,b) was fol-
lowed by a report of Serra and Queiroz Lopes ( 1944) , who empha-
sized the usefulness of the reaction for the visualization of the basic
proteins such as those contained in cell nuclei. Subsequently, Serra
(1946) summarized the work of his group on the arginine reaction
in the course of a more general article dealing with histochemical
tests for proteins and amino acids. Serra pointed out that a positive
reaction is found only with arginine and the rather rare compounds
glycocyamine, gelegine, and agmatine, negative reactions being given
by guanidine, urea, ornithine, creatine, creatinine, asparagine, histi-
dine, and other amino acids. The reaction is specific for guanidine
derivatives in which one hydrogen atom of one or both amino groups
is substituted by an alkyl, fatty acid, or cyano radical. Substitution
of other radicals has not been tested, while guanidine derivatives in
which both hydrogen atoms of one amino group are substituted do
not give the color (Thomas, 1946) .

ARGININE AND ARGININE-CONTAINING PROTEINS 57

The procedures of Serra and Thomas differ in certain details
and, at the date of this writing, no comparison of the two has been
made. An advantage of the method of Thomas is that it does not
employ cooling in an ice bath during the reaction because of the
substitution of hypochlorite for hypobromite. Serra mounts his
sections in glycerol after several transfers through this medium and
he has reported that in this fashion the color, otherwise stable for
only a short time, is stabilized for months.

Serra Method for Arginine and Arginine-Containing Proteins

SPECIAL REAGENTS

Acetic- Alcohol-Formalin Fixative. Add a few drops of glacial

acetic acid to each 10 ml. of a mixture of 2 vol. 96% alcohol and

1 vol. formalin.
1% a-Naphwl in 96% Alcohol. Store in a refrigerator. Dilute

1:10 with 40% alcohol before use.
4% Sodium Hydroxide.
2% Sodium Hypobromite. With stirring and cooling, add 2 g. or

approximately 0.7 ml. bromine to 100 ml. 5% sodium hydroxide.

Store in a refrigerator.
40% Urea.

PROCEDURE

1. Fix the material in the acetic-alcohol-formalin mixture. Wash
well in water.

2. Transfer to a watch glass kept at 0-5° in an ice bath, and
treat for 15 min. at this temperature with a mixture of 0.5 ml. a-
naphthol soln., 0.5 ml. 1 N sodium hydroxide, and 0.2 ml. 40% urea.

3. Add 2 ml. 2% hypobromite, and after 3 mih. stir in 0.2 ml.
urea soln. and then 0.2 ml. of the hypobromite. The maximum color
develops in 3-5 min.; intensify it by a subsequent treatment with
the hypobromite for 3 min.

4. Pass through four glycerol baths, leaving for 2-3 min. in
each. In glycerol the color is stable for months even at room tem-
perature. The fading is inhibited by storage in the cold.

Result. An orange-red color characterizes a positive reaction.

58 MICROSCOPIC TECHNIQUES

Thomas Method for Arginine and Arginine-Containing
Proteins

SPECIAL REAGENTS

0.1% a-Naphthol in 10% (by vol.) Ethyl Alcohol.

0.15 N Sodium Hyp/ochlorite in 0.05 N Sodium Hydroxide. For
preparation of the hypochlorite see page 239; or prepare from
Clorox which is standardized and then stored at 3-5° in a dark
bottle. Dilute the hypochlorite to the proper strength each time
before use. Clorox is approximately 1.6 A^; standardize by adding
1 ml. Clorox to 5 ml. 1 A^ potassium iodide, 8 ml. cone, hydro-
chloric acid (sp. gr. 1.19), and 45 ml. water. Titrate with 0.1 N
sodium thiosulfate using starch indicator.

20% Urea in 0.05 N Sodium Hydroxide.

Tertiary Butyl Alcohol Solution. Add 1 ml. 5 N sodium hydroxide
and 19 ml. water to 80 ml. of the tertiary butyl alcohol. Shake
well and let stand; an aqueous layer collects on the bottom of the
vessel.

Pure Tertiary Butyl Alcohol.

Aniline.

Toluene.

PROCEDURE

1. Fix animal tissues in Bouin fluid (75 ml. saturated picric
acid soln., 25 ml. formalin, 5 ml. glacial acetic acid) . Onion root
tips were treated with medium chrome-acetic fixative.

2. Prepare paraffin sections in the usual manner. Do not remove
paraffin from sections until test is to be applied.

3. Place slide with sections in the a-naphthol soln. for at least
3 min.

4. Transfer to each of the following solns. in succession for the
periods indicated: hypochlorite, 20 sec; urea, 5 sec; 80% tertiary
butyl alcohol, 30 sec; pure tertiary butyl alcohol, 2 min.; aniline,
2 min.; toluene, 5 sec; and finally mount in Clarite. Use a stop watch
to time the immersions in each fluid.

TRYPTOPHANE IN PROTEINS

The red or violet color formed with proteins in the presence of
phosphoric acid is the basis of the Romieu reaction. Blanchetiere

TRYPTOPHANE AND TYROSINE 59

and Romieu (1931) presented evidence that the effect was the
result of tryptophane groups in the protein. As in the other protein
tests, the drastic nature of the reaction seriously interferes with its
use in most instances, as does the diffusibility of the color formed.

Romieu Reaction for Tryptophane in Proteins

SPECIAL REAGENTS

Syrupy Phosphoric Acid.

PROCEDURE

1. Fix tissue in alcohol, formalin, or Bouin fluid.

2. Prepare fairly thick paraffin or celloidin sections and remove
the infiltrating agent.

3. Place a drop of the phosphoric acid on a section and set in
an oven at 56° for a few min.

4. Examine on removal from oven.

Result. A positive test is manifest by the formation of a red or
violet color.

TYROSINE IN PROTEINS

Bensley's histochemical adaptation of well-known Millon reac-
tion for proteins containing tyrosine has been employed in studies
by Bensley and Gersh ( 1933b) .

Millon Reaction for Tyrosine in Proteins
(after Bensley and Gersh)

SPECIAL REAGENTS

Millon Reagent. Add 1 vol. 40% nitric acid (add 600 ml. distilled
water to 400 ml. cone, nitric acid, sp. gr. 1.42; let stand for 48
hr.) to 9 vol. distilled water and saturate with mercuric nitrate
crystals by frequent shaking over several days. Filter, and, to 400
ml. of filtrate, add 3 ml. 40% nitric acid and 1.4 g. sodium nitrite.

1 % Nitric Acid.

PROCEDURE

1. Mount sections on slides without using water. The freezing-
drying technique is preferable.

2. Place in cold Millon reagent. Since the maximum color is
developed in about 3 hr., remove each slide at a different time, dip

_(Ll$ii^ARY

60 MICROSCOPIC TECHNIQUES

immediately in 1% nitric acid and dehydrate rapidly in absolute
alcohol.

3. Clear in xylol and mount in balsam.

Result. A brick-red or rose color develops in the presence of
tyrosine or proteins containing tyrosine.

a-AMINO ACID GROUPS IN PROTEINS

Less soluble peptides and proteins containing a-amino acids may
be demonstrated at their loci in tissue sections by either the alloxan
or ninhydrin reactions. A tendency for the color to diffuse in the
alloxan reaction indicates that caution should be applied in inter-
preting the test, as Giroud (1929) has warned; furthermore the
specificity is not great enough to exclude the need for confirmatory
tests (Romieu, 1925). Hence only the ninhydrin reaction of Berg
( 1926) will be described. A positive reaction is obtained with many
amines, aldehydes, and ammonium compounds as well as with the
amino acids, but the solubility of these compounds enables their
easy removal as a rule.

Berg Ninhydrin Test for a-Aniino Acid Groups

SPECIAL REAGENTS

0.2% Ninhijdrin.

PROCEDURE

1. Fix tissue in 10% formalin.

2. Wash in water and prepare frozen sections.

3. Boil sections in 2 ml. of the ninhydrin soln. for 1 min.

4. Wash in water and mount in glycerin or glycerin jelly.

Result. a-Amino acid groups give rise to an intense blue or violet
color; this should be observed the same day as it fades rapidly.

note: Sena and Quieroz Lopes (1945) emploj^ed a mixture of equal vol.
of 0.4% ninlij'drin in distilled water and phosphate buffer of pH 6.98 (6 ml.
M/15 secondary sodium phosphate — 1L1876 g. Na2HP04.2Hi.O per liter —
and 4 ml. M lib primary potassium phosphate — 9.078 g. KHsPO* per liter).
They heat the sections in the liquid in a watch glass placed over a boiling
water bath. For cementing of the preparations they employ the mixture of
Romeis, which is 80 g. colophonium and 20 g. carefully heated lanolin.

MELANIN 61

MELANIN

Perhaps the most characteristic microchemical test for melanin
is its ability to reduce ammoniacal silver nitrate. Of course many-
other tissue constituents have this property so that the test is of
value only when possible interferences (page 48) are considered.
Dublin (1943) applied the Bodian silver method to the demonstra-
tion of melanin; his procedure follows.

Dublin Application of the Bodian Method to Demonstration
of Melanin

SPECIAL REAGENTS

Protargol Solution. Prepare fresh each time by adding one or more
drops of Protargol (Winthrop) — other brands do not appear to
be satisfactory — to water in a staining jar. The color should be
light amber. Do not stir or mix the solution since this results in
gumming.

1.0% Hydroquinone.

0.5% Auric Chloride.

5.0% Oxalic Acid.

10% Sodium Thiosulfate.

PROCEDURE

1. Fix tissue in 10% formalin.

2. Prepare paraffin sections 8 fx thick.

3. Treat the deparaffinized sections, after passing through graded
alcohols to water, with the Protargol soln. overnight.

4. Rinse with water and place in the hydroquinone soln. for 10
min.

5. Rinse with water and place in the auric chloride soln. for 5
min.

6. Rinse with water and place in the oxalic acid soln. for 5 min.

7. Rinse with water and place in the thiosulfate soln. for 5 min.

8. Wash in running tap water for 10 min.

9. Dehydrate, clear, and mount as usual.

Result. The melanin will appear black and the background a
purplish brown or gray.

62 MICROSCOPIC TECHNIQUES

HEMOGLOBIN

Of the many tests for hemoglobin in tissue, smears and blood cells
the more recent procedures of Ralph ( 1941) , Goulliart ( 1939, 1941) ,
and Dunn ( 1946) will be given. Previously Dunn and Thompson

(1945) had modified the Van Gieson stain, and later these authors

( 1946) adapted the patent blue method of Lison ( 1938) for the
staining of hemoglobin. The cyanol method of Dunn given below is
a simplification of the technique of Fautrez and Lambert ( 1937) .

Ralph Method for Hemoglobin

SPECIAL REAGENTS

Benzidine Reagent. 1% benzidine in absolute methanol.
Peroxide Reagent. 25% Superoxol in 70% ethanol.

PROCEDURE

1. Flood the dried blood or tissue smear on a glass slide with the
benzidine reagent for 1 min.

2. Drain off and flood the smear with the peroxide reagent for 1.5
min.

3. Wash in distilled water for 15 sec.

4. Dry and mount in neutral balsam.

Result. Hemoglobin will appear dark brown.

Goulliart Method for Hemoglobin

SPECIAL REAGENTS

Glacial Acetic Acid Containing a Few Crystals of Potassium Iodide.
Do not use after a week.

PROCEDURE

1. Treat a dried smear or frozen section on a slide with a drop of
reagent.

2. Examine after 30 min. with a polarizing microscope for groups
of very small boat-shaped birefringent crystals of protoiodoheme.
These crystals slowly change into square tabular Teichmann crys-
tals. The reaction may be speeded by warming.

HEMOGLOBIN, BILE PIGMENTS AND ACIDS 63

Dunn Method for Hemoglobin

SPECIAL REAGENTS

Cyanol Stock Solution. Dissolve 1 g. cyanol {National Aniline
Division, Allied Chemical and Dye Corp.) in 100 ml. distilled
water, add 10 g. pure zinc powder, and 2 ml. glacial acetic acid.
Bring mixtm^e to a boil and the blue color will soon fade out. The
soln. is stable for several weeks.

Cyanol Working Solution. Just before use filter 10 ml. of the stock
soln., add 2 ml. glacial acetic acid and 1 ml. commercial 3%
hydrogen peroxide.

PROCEDURE

1. Prepare frozen or paraffin sections of tissue fixed in 4%
formaldehyde buffered to pH 7.0.

2. Bring sections to water and stain in cyanol working solution
3-5 min.

3. Rinse in water and counterstain in safranin (1:1000 in 1%
acetic acid) 1 min.

4. Wash in water, dehydrate, clear, and mount in Clarite.

Result. Hemoglobin stains dark blue to bluish-gray; nuclei,
red; and cytoplasm, light pink.

BILE PIGMENTS AND ACIDS

The well-known Gmelin test has been adapted to the microscopic
detection of bile pigments by simply adding a drop of nitric acid
containing some nitrous acid to the sample on a slide. A positive
test is indicated by the appearance of a green color changing to red
and finally to blue. Stein's test ( 1935) , given below, is probably more
satisfactory. Bile salts and acids may be precipitated by barium and
the precipitate stained with acid fuchsin according to the technique
of Forsgren ( 1928) .

Stein Test for Bile Pigments

SPECIAL REAGENTS

Iodine Reagent. 2 or 3 vol. Lugol solution (6 g. potassium iodide
and 4 g. iodine dissolved in 100 ml. distilled water) plus 1 vol.
tincture of iodine.

5% Sodium Hyposidfite.

64 MICROSCOPIC TECHNIQUES

PROCEDURE

1. Fix for a short period in alcohol or 10% formalin.

2. Prepare paraffin sections and employ egg albumin to hold to
slides.

3. After removal of paraffin and bringing down to water, subject
sections to the iodine reagent for 6-12 hr.

4. Wash in distilled water and decolorize with the sodium hypo-
sulfite for 15-30 sec.

5. Wash in distilled water and stain with alum carmine for 1-3
hr.

6. Wash in distilled water, dehydrate in acetone, clear in xylol,
and mount in balsam.

Result. Bile pigments appear emerald green. Localizations can-
not be considered reliable due to the diffusibility of the reactants and
the final color.

Forsgren Test for Bile Acids

SPECIAL REAGENTS

3% Barium Chloride.
0.1 % Acid Fuchsin.
1 % Phosphomolybdic Acid.

Aniline Blue-Orange G Stain. Dissolve 0.5 g. aniline blue, 2 g.
orange G, and 2 g. oxalic acid in 100 ml. distilled water.

PROCEDURE

1. Treat small pieces of tissue for 6-12 hr. with the barium
chloride soln.

2. Fix in 10% formalin for 12-18 hr.

3. Prepare paraffin sections.

4. Stain sections for 1-3 min. in the acid fuchsin soln., and
wash in distilled water.

5. Place sections in the phosphomolybdic acid soln. for 0.5-1.0
min. and wash in distilled water.

6. Treat sections for 3-5 min. with the aniline blue-orange G
staip and wash in distilled water.

7. Dehydrate, clear, and mount in balsam.

Result. Bile secretory granules appear reddish.

ALDEHYDES, NUCLEIC ACIDS, AND PLASMAL 65

ALDEHYDES, NUCLEIC ACIDS, AND "PLASMAL"

The research of Feulgen and co-workers (1924,1938,1939) and
Imhiiuser ( 1927) led to the demonstration of a loosely bound alde-
hyde, "plasmal," in animal tissues. The bound form, "plasmalogen,"
liberates "plasmal" when treated with mercuric chloride or sub-
jected to prolonged acid hydrolysis. The Feulgen reaction, which
depends on the formation of a purple-colored compound when alde-
hydes react with fuchsin-sulfurous acid, is also given by desoxyri-
bonucleic acid (thymonucleic acid) after its purine bases are re-
moved by acid hydrolysis, but ribonucleic acid does not give the re-
action. The application of the Feulgen reaction to histochemical
studies on animal tissues was elaborated by Cowdry ( 1928) and
Verne (1928). Milovidov (1938) published a complete bibliography
of the 450 papers dealing with the Feulgen reaction up to 1938. Since
then Whitaker (1938) described a technique for plant tissues,
Stowell (1945a) studied the Feulgen reaction for the photometric
measurement of desoxyribonucleic acid (page 126), and Oster and
associates (1942,1944) employed the histochemical approach to
study tissue aldehydes in sections of fresh frozen material. An im-
proved preparation of the Feulgen reagent was reported by Cole-
man (1938). Rafalko (1946) claimed that small and diffuse chro-
matin elements could be detected with greater delicacy when the re-
agent was made by decolorizing a 0.5% solution of the dye by
bubbling sulfur dioxide through it.

The specificity of the Feulgen reaction for aldehydes has been
brought up repeatedly. It has been variously claimed that oleic and
cinnamic acids give a positive reaction, and that ketosteroids can-
not be differentiated from aldehydes by the reaction. Oster and
Oster (1946) have examined the question of specificity and have
found that the "true" reaction is indeed specific for aldehydes, other
carbonyl compounds giving a "pseudo" reaction in certain instances.
The differentiation between the "true" and "pseudo" reactions may
be made according to Oster and Mulinos ( 1944) x)n the basis that
the purple color developed in the former can be decolorized with di-
lute sodium hydroxide and restored to its original intensity with
hydrochloric acid, while the reddish color of the latter cannot be re-
stored by acid after the decolorization.

66 MICROSCOPIC TECHNIQUES

A means for the microscopic demonstration of ribonucleic acid
was developed by Diibos ( 1937) and Brachet ( 1940) , who employed
ribonuclease to break down the compound and thus destroy its
basophilic staining properties. The crystalline ribonuclease prepared
by Kunitz ( 1940) provided a more satisfactory reagent for carrying
out the procedure. Opie and Lavin ( 1946) demonstrated that ribo-
nucleic acid can be protected againct ribonuclease by precipitation
of the acid with lanthanum acetate. The basophilia of the precipitate
was retained even after treatment with ribonuclease.

The danger of an uncritical acceptance of the localizations ob-
tained by the Feulgen reaction has been emphasized by Danielli
(1946a). He pointed out that it remains to be proved whether the
experimental treatment of the nucleic acid has rendered it diffusible
enough for this factor to become significant in the interpretation. In
addition, he stressed the point that the use of an enzyme to digest
away a particular substance is open to some question with reference
to the specificity of the enzyme and the degree to which a clear-cut
removal of the substrate is possible. On the other hand, Stowell
( 1946) reviewed the evidence for and against the specificity of the
Feulgen technique for thymonucleic acid, and he concluded that
with the proper precautions it is one of the most specific histochem-
ical reactions. This does not mean that Stowell considers the tech-
nique beyond all criticism. No doubt he would agree with Danielli
that the interpretation of the results should be tempered with a
healthy awareness of the limitations involved, particularly the
diffusibility factor.*

Turchini and co-workers (1943, 1944, 1945) reported the use of
9-phenyl (or methyl) -2,6,7-trihydroxy-3-fluorone for the differential
staining of ribo- and desoxyribonucleic acids, the former giving rise
to a yellow-pink color, and the latter to a blue-violet. It is necessary
to hydrolyze the nucleic acid, as it is the pentose, thus liberated,
which yields the color. The hexoses formed by the hydrolysis of
tannins produce an orange-yellow color in the staining reaction when
it is applied to plant tissues (Turchini and Gosselin de Beaumont,
1945). -^i

* Other publications which have appeared subsequent to this writing are
given in the Bibliography Appendix, Refs. 15, 18 and 31.

ALDEHYDES, NUCLEIC ACIDS, AND PLASMAL 67

The use of ultraviolet absorption for the localization of nucleic
acids is discussed on page 113.

Coleman Preparation of Feulgen Reagent

Dissolve 1 g. basic fuchsin in 200 ml. boiling water; filter, cool, and
add 2 g. potassium metabisulfite (K2S2O5) and 10 ml. 1 N hydro-
chloric aci^. Let bleach for 24 hr., and then add 0.5 g. activated car-
bon (Norit), shake for about 1 min., and filter through coarse paper.
The filtrate should be colorless.

Whitaker Feulgen Technique for Plant Tissues

SPECIAL REAGENTS

Modified Brenda Fixative. Combine 30 ml. 1% chromic acid with

10 cc. 2% osmic acid.
1 N Hydrochloric Acid.
Feulgen Reagent. See above.
45% Acetic Acid.

PROCEDURE

1. Fix tissue in the modified Brenda fluid for a period depending
on the specimen, e.g., 15-20 min. for root tips, 30-45 min. for whole
anthers.

2. Hydrolyze in 1 .V hydrochloric acid at 50-60° for the same
time used in fixation.

3. Place in stain for 15-20 min. and then transfer to 45% acetic
acid for 10-15 min. or longer.

4. Put specimen in a drop of 45% acetic acid on a glass slide and
perform any dissections at this stage.

5. Place cover slip over the material and heat the slide nearly to
boiling at least three times. Apply pressure to cover slip with each
heating to make the tissue adhere to the slide.

6. Float off the cover slip in a mixture of equal vol. absolute
alcohol and glacial acetic acid.

7. Transfer to 95% alcohol for at least 15 min. and mount in
euperal. The mounting must be done in low humidity and care must
be taken to avoid breathing on the slide since moisture results in
cloudiness. The mounted material keeps well permanently.

Result. A positive reaction is indicated by a purple color.

68 MICROSCOPIC TECHNIQUES

Cowdry Modification of Feulgen Reaction for Paraffin
Sections of Animal Tissues

SPECIAL REAGENTS

Sublimate-Alcohol Fixative. Combine equal vol. saturated mercuric

chloride soln. and absolute alcohol.
1 N Hydrochloric Acid.
Feulgen Reagent. See page 67.
Sodium Bisulfite Solution. Add 30 ml. 1 M sodium bisulfite soln.

to 600 ml. tap water.

PROCEDURE

1. Prepare paraffin sections of tissue fixed in the sublimate-
alcohol fluid.

2. Pass through graded alcohols to water and place in the hydro-
chloric acid for 1 min.

3. Place in another portion of the acid at 60° for 4 min.

4. Treat with the Feulgen reagent for about 1.5 hr. The time may
have to be varied to suit the particular sections used.

5. Pass through three separate portions of the sodium bisulfite
soln. leaving in each for 1.5 min. and agitating frequently.

6. Wash for 5 min. in tap water.

7. Dehydrate, clear, and mount in balsam.

Oster Modification of Feulgen Reaction for Fresh-Frozen
Sections of Animal Tissues

SPECIAL REAGENTS

1 % Mercuric Chloride.

Feulgen Reagent. See page 67.

0.01 N Hydrochloric Acid Containing 1 % Sodium Bisulfite.

PROCEDURE

1. Cut 50 /x sections of fresh tissue on a freezing microtome. (The
sectioning should be carried out within 2-3 hr. after the death of the
animal and removal of the tissue. Until ready for use, keep the tissue
before cutting, and the sections after cutting, in physiological salt
solution.)

2. Place the sections in 1 % mercuric chloride for 5 min. in order
to liberate free aldehyde from "plasmalogen." Wash with water.

WATER-INSOLUBLE CARBONYL COMPOUNDS 69

3. Transfer to the Feulgen reagent for 15 min. and hold the
stained sections in the hydrochloric acid-sodium bisulfite solution.

4. Examine sections immediately after washing in distilled water.
The stain will last for a few days if the sections are kept in sulfurous
acid solution.

Method of Turchini et al. for Nucleic Acids

SPECIAL REAGENTS

Nucleic Acid Reagent. Dissolve 80 mg. of 9-phenyl (or methyl) -
2,6,7-trihydroxy-3-fluorone in 100 ml. 95% alcohol containing 15
drops cone, sulfuric acid.

1 N Hydrochloric Acid. (Or 25% cone, hydrochloric acid in 90%
alcohol.)

1 % Sodium Carbonate.

PROCEDURE

1. Fix the tissue (either plant or animal) in Bouin fluid.

2. Prepare paraffin sections in the usual manner.

3. If the methyltrihydroxyfluorone reagent is used: Hydrolyze
the deparaffinized sections in 1 A^" hydrochloric acid at 60° for 5 min.
wash with water, then alcohol, and treat for 5-10 min. with the re-
agent. Wash with several drops of 90% alcohol, then with 1%
sodium carbonate, rinse with water, and finally mount in balsam.

3a. With the phenyltrihydroxyfluorone reagent: Use the same
procedure as in step 3 but carry out the hydrolysis in the cold in
alcoholic 25% hydrochloric acid for 3-5 min.

WATER-INSOLUBLE CARBONYL COMPOUNDS

While the fuchsin-sulfurous acid test can be used for the localiza-
tion of aldehydes in tissue, other histochemical tests employed by
Bennett (1939, 1940) will react with either aldehydes or ketones.
Bennett concluded that his tests for the carbonyl group were indica-
tive of ketosteroids in the outer layer of the fascicular region of the
adrenal cortex. These carbonyl reactions can only indicate lipid
aldehj'-de or ketone and are in no way specific for ketosteroids as
Gomori (1942) pointed out; however, if other supporting evidence is
at hand, it may be reasonable to ascribe a positive reaction to the
ketosteroids present in a particular tissue. Subsequent work of
Albert and Leblond ( 1946) indicated that '"plasmalogen" rather
than ketosteroids is revealed by the phenylhydrazine reaction.

70 MICROSCOPIC TECHNIQUES

Bennett ( 1940) first removed ascorbic acid from the tissue to pre-
vent its interference with the tests. Albert and Leblond (1946)
substituted 2,4-dinitrophenylhydrazine for the phenylhydrazine of
Bennett. This enabled a more intense staining in thinner sections.

Bennett Use of Phenylhydrazine Reaction for Water-Insoluble
Aldehydes and Ketones

SPECIAL REAGENTS

M/10 Acetate Buffer, pH 6.0 to 6.5.

1 % Iodine in Alcohol.

1% Sodium Thiosulfate Solution.

1% Buffered Phenylhydrazine. Prepare just before use by mixing
equal vol. of 2% phenylhydrazine hydrochloride and the acetate
buffer. Gently bubble carbon dioxide through the solution for 15
min. to remove oxygen.

Control Reagent. Same as the 1 % buffered phenylhydrazine with-
out the phenylhydrazine.

PROCEDURE

1. Transfer frozen sections of fresh tissue from the microtome
directly into acetate buffer. If fixed tissue is employed, transfer to
water.

2. Add the iodine solution dropwise until a faint straw color
persists, and let stand 15 min.

3. Add sodium thiosulfate solution dropwise until the color is
discharged and a little more has been added ; let stand 5 min.

4. Wash the sections several times in distilled water.

5. Place the sections in glass-stoppered bottles containing
buffered phenylhydrazine solution. Fill the bottles to the top so that
no air bubbles are present under the stopper.

6. Run control sections as in previous steps, only use the control
reagent in place of phenylhydrazine.

7. After standing several hr. or overnight, wash all sections
with distilled water a few times.

8. Mount in glycerol or glycerol-gelatin and examine by means
of incident illumination from above.

Result. A yellow color appears in areas giving the positive test.
It is essential that care be taken in conducting this test since the
appearance of a yellow deposit on the walls of the bottle or on top
of the liquid indicates decomposition of the reagent, and when this

WATER-INSOLUBLE CARBONYL COMPOUNDS 71

occurs the yellow color in the sections cannot be relied upon to be
specific for the groups tested.

Albert and Leblond Use of 2,4-Dinitrophenylliydrazine Reaction
for Water-Insoluble Aldehydes and Ketones

SPECIAL REAGENTS

2,4-Dinitrophenylhydrazine Reagent. To a saturated soln. of 2,4-
dinitrophenylhydrazine (No. 1866 Eastman Kodak Co.) in 30%
alcohol, add sufficient 0.2 A^ sodium acetate to bring the pH to
neutrality.

PROCEDURE

1. Fix tissue for 48 hr. in formalin (neutralized with magnesium
carbonate) and wash in nmning water for 24 hr.

2. Prepare frozen sections 10-15 fx and place in 17% alcohol
for 4 hr.

3. Place sections in the 2,4-dinitrophenylhydrazine reagent over-
night and wash in 17% alcohol for 20 min.

4. Carry to distilled water and mount in glycerol gelatin.
Result. A positive reaction is shown by a yellow color.

Bennett Use of Seniicarl>azide Reaction for Water-Insoluble
Aldehydes and Ketones

SPECIAL REAGENTS

Acetate Buffer, Iodine, and Thiosulfate Solutions. Same as in the

preceding phenylhydrazine test.
Semicarbazide Reagent. Grind 10 g. semicarbazide hydrochloride

with 15 g. crystalline sodium acetate, take up the mixture in 100

ml. absolute methanol, and filter.
Control Reagent. Same as the semicarbazide reagent without the

semicarbazide.

PROCEDURE

1.-4. Follow the first four steps in the procedure for the phenyl-
hydrazine test,

5. Place sections in the semicarbazide reagent.

6. Place control sections in the control reagent.

7. After overnight standing, wash all sections several times with
distilled water.

8. Examine at unce with incident light from above.

72 MICROSCOPIC TECHNIQUES

Result. A yellowish deposit of semicarbazones appears in the
areas where the aldehydes and ketones are present.

PURINES

Tests that have been found to give positive results with all of the
purines have the unhappy characteristic of being highly unspecific.
Thus the reduction of silver salts is a reaction much too unspecific
to merit consideration; Saint-Hilaire's method involving precipita-
tion of insoluble copper salts of purines, and the transformation of
the copper into its red ferrocyanide is a reaction also given by
protamines, histones, and other protein products (Lison 1936, pages
183-186) .

The murexide test, which is positive with uric acid, xanthine and
its methyl derivatives, and guanine, is not given by adenine or
hypoxanthine (Lison 1936, pages 186-187). This reaction has the
disadvantage of being too drastic to permit its use for fine structures
and its disintegrating effect on tissue sections presents technical
difficulties. However, it may prove useful in some cases and for this
reason it will be described. Since it is no different from the xantho-
proteic reaction, a yellow-orange color would be indicative of pro-
teins, but it should be kept in mind that the xanthoproteic test is
not specific for proteins since other compounds, such as alkaloids,
benzene derivatives, etc., can also be nitrated in this manner to yield
products having the same color. •

Cowdry (1943, page 196) suggested that the modification of the
Courmont-Andre method by Hollande (1931) be used. It enables a
more reliable localization of urates in tissue.

Murexide Test for Certain Purines

SPECIAL REAGENTS

Concentrated Nitric Acid.
Concentrated Avfimonium Hydroxide.

PROCEDURE

1. Prepare sections by any of the usual methods.

2. Place a drop of nitric acid on a section and warm gently for
30 sec.

PURINES, INDOLE AND RELATED COMPOUNDS 73

3. Drain off the acid by means of blotting paper and add a drop
of water, which is also removed in the same way.

4. Expose the section to ammonia vapors.

Result. A purple-violet color is a positive test for uric acid,
guanine, and xanthine and its methyl derivatives. A yellow-orange
color is usually indicative of protein material. The effect of diffus-
ibility should be considered in the interpretation of localizations.

HoUande Modification of Courmont-Andre Method
for Uric Acid and Urates

SPECIAL REAGENTS

Silver Nitrate-Neutral Formalin Fixative. Equal vol. of 1% silver
nitrate and 4.4% formalin (neutralized with calcium carbonate)
mixed just before using.

0.5% Phosphomolyb die Acid.

PROCEDURE

1. Fix tissue in silver-formalin mixture for 12-24 hr. in the dark.

2. Wash for 24 hr. in several changes of distilled water.

3. Prepare paraffin sections.

4. Stain sections with hemalum for 10 min. and wash in running
tap water for 30-60 min.

5. Place in 1 % aqueous orange G or eosin 30-60 min. and wash
rapidly in distilled water.

6. Treat with the phosphomolybdic acid soln. and wash in dis-
tilled water.

7. Stain with 0.12% aqueous light green for 1-10 min.

8. Differentiate quickly in 95% alcohol, dehydrate in isoamyl
alcohol, clear in xylol, and mount in balsam.

Result. Urates will appear black, chromatin blue, protoplasmic
inclusions red to orange, and collagenic fibers green.

INDOLE AND RELATED COMPOUNDS

Lison (1936, pages 160-162) lists five reactions for the histo-
chemical detection of compounds containing the indole structure.
All of the tests leave much to be desired; their specificity is rather

74 MICROSCOPIC TECHNIQUES

poor. The Ehrlich reaction employing p-dimethylaminobenzal-
dehyde will give a violet color with phenols, aryl amines, and
heterocyclic compounds. Diazotization may indicate the same classes
of substances. The indophenin reaction utilizing isatin and sulfuric
acid gives a reddish-violet color in the presence of five-membered
heterocyclic compounds including indole. The nitrosamino reaction
of Lison converts the imino group in pyrrole or indole to a nitros-
amine by means of nitrous acid, and the nitrosamine is then made
to produce a green color through the Liebermann reagent (5%
phenol and concentrated sulfuric acid). This test is given by imino
groups, phenols, and primary aryl amines. The nitro reaction
enables differentiation between pyrroles and indoles; the sections
are treated with a mixture of equal parts of sulfuric and nitric acids,
and benzene ring compounds including indoles develop a canary
yellow color while pyrroles are not colored.

PHENOLS

Four main staining reactions have been employed for the detection
of phenols in tissue preparations. The azo reaction is based on diaz-
otization to form colored compounds; the indo reaction depends
on the formation of a green or blue indamine when an aromatic
para diamine is oxidized in the presence of tissue phenol; the
"argentaffin" reaction makes use of the reduction of ammoniacal
silver hydroxide and applies to ortho and para polyphenols, poly-
amines, and aminophenols ; and the "chromaffin" reaction, which is
used particularly to indicate adrenaline, gives rise to a brown color
when tissue is fixed with dichromate salts. A discussion of these tests
was given by Lison ( 1936, page 139-160) . The argentaffin test is
quite unspecific since many reducing substances can likewise give
a positive reaction. The chromaffin test is not entirely specific for
adrenaline, but has proved useful for the histochemical localization
of this biologically important substance.

Lison Modification of Chromaffin Reaction

SPECIAL REAGENTS

Formol-Milller Fixative or 5% potassium iodate in 10% formalin.
3% Potassium Dichromate or Potassium Iodate.

PHENOLS, UREA, AND SULFONAMIDES 75

PROCEDURE

1. Fix tissue in one of the solns. indicated. The iodate gives a
less intense reaction but is less prone to pseudoreactions.

2. Prepare sections and treat them with the 3% reagent for a
few hours.

Result. A brownish color indicates a positive reaction.

UREA

Two methods have been proposed for the localization of urea in
tissue sections. The Leschke procedure is based on fixation of the
tissue in a half-saturated mercuric nitrate solution in 1% nitric
acid and subsequent treatment of the sections with a saturated
hydrogen sulfide solution. The mercury urea compound is converted
to black mercuric sulfide, which is easily visualized. The xanthydrol
method depends on fixation of the tissue in a solution of xanthydrol
in acetic acid in order to precipitate dixanthylurea which can be
recognized in sections by its double refraction when examined under
a polarizing microscope. Lison ( 1936, pages 165-170) critically dis-
cussed these methods. As he pointed out, the usefulness of the mer-
cury reaction is entirely vitiated by its extreme lack of specificity,
far too many tissue constituents being capable of precipitation by
mercury salts. The xanthydrol reaction is chemically specific, but
its serious fault lies in a combination of unfortunate factors including
the great diffusibility of urea, the poor penetrability of xanthydrol,
and the slowness of the reaction between the two. The result is that
the position of the crystals formed bears little or no relation to the
regions originally containing urea. A suitable method for the his-
tological localization of urea is not available at present.

SULFONAMIDES

^lacKee et al. ( 1943) described a test for sulfonamides in frozen
tissue sections depending on the formation of a yellow to orange
precipitate of the p-dimethylaminobenzylidene derivative when sul-
fonamides react with p-dimethylaminobenzaldehyde. It should be
borne in mind that procaine, phenacetin, acetanilid, and aromatic
amino compounds in general will also give the reaction.

76 MICROSCOPIC TECHNIQUES

Another method was published by Hackmann (1942), who em-
ployed the freezing-drying technique for the fixation of the tissue,
prior to the preparation of paraffin sections. Colored sulfonamides
were observed directly in the sections, and colorless ones were
visualized by forming a red azo dye in the following manner: The
sections were exposed to nitrous acid vapor for 30 sec. by placing
the slide over a measuring cylinder 20 cm. high containing several
milliliters of 0.1 A^" hydrochloric acid and a few milligrams of sodium
nitrite. The diazotized sulfonamide was coupled with a-naphthyl-
amine by immersing the slide in a 5% solution of the amine in xylol.

The detection of sulfonamides by fluorescence microscopy is
discussed on page 108.

Method of MacKee et ah for Sulfonamides

SPECIAL REAGENTS

Sulfa Reagent. Dissolve 1 g. pure p-dimethylaminobenzaldehyde
in a soln. of 95 ml. absolute alcohol and 5 ml. cone, hydrochloric
acid. Store in a glass-stoppered amber bottle, and do not use after
2-3 weeks or when the soln. becomes yellow.

5% Concentrated Hydrochloric Acid in Absolute Alcohol.

PROCEDURE

1. Fix the tissue for 2-24 hr. with formaldehyde gas by covering
the bottom of a beaker with paraformaldehyde and the top with a
piece of gauze, placing the tissue on the gauze, setting the whole in
a glass jar whose floor has also been covered with paraformaldehyde,
and closing the jar tightly with a glass lid.

2. Cut frozen sections of the fixed tissue 10-20 /^ thick and place
directly on glass slides.

3. Cover each section with a drop or two of the sulfa reagent,
and after 3-5 min. add a drop or two of the alcoholic hydrochloric
acid soln.

4. Dry quickly without heat by absorbing excess fluid on filter
paper and holding in a current of air.

5. Cover at once with a drop of damar resin in xylol (10 g.
resin dissolved in 10 g. xylol) and fit cover slip, taking care to re-
move air bubbles. Seal edges with melted paraffin.

SULFONAMIDES AND UREASE 77

6. Run controls by repeating the above steps but omitting the
treatment with the sulfa reagent, or repeat the complete procedure
on a portion of the same kind of tissue known to be free of sulfona-
mides.

Result. Sulfonamides are indicated by the presence of a pre-
cipitate that ranges in color from lemon-yellow to orange. However,
the color fades rapidly, particularly in the presence of air, making
it necessary to examine the sections as early as possible. Colored
photomicrographs should be taken not later than 3-4 hr. after the
reaction has occurred.

D. ENZYMES

UREASE

Sen ( 1930) elaborated a method for the localization of urease in
tissue sections which he employed for a study on the jack bean. The
carbonic acid formed on decomposition of urea is precipitated as
calcium carbonate, which may be visualized by conversion to silver
carbonate and reduction of the latter to a black deposit of metallic
silver; or the carbonic acid may be converted to cobalt carbonate and
the latter changed to a brown or black precipitate of cobalt sulfide.
The latter method is to be preferred. This principle was later employed
by Gomori for the localization of the phosphoric acid hberated by
phosphatases, pages 78 and 80. However, Sen digested the tissue
in the substrate medium before paraffin infiltration and sectioning,
and only treated the deparaffinized sections with sulfide to convert
the colorless cobalt salt to the black sulfide. This procedure has many
disadvantages; the schedule of Gomori, in which the sections are
prepared prior to digestion, should be used instead, if the enzyme
can stand the dehydration, paraffin embedding, and deparaffinization.

For jack bean tissue. Sen employed a preliminary treatment for
1 hr. with 1% cobalt nitrate in 80% alcohol followed by a 48-60
hr. digestion with a substrate medium consisting of 0.5% urea and
0.5% cobalt nitrate in 80% alcohol. The cobalt carbonate was con-
verted to sulfide by the action of either dilute sodium sulfide or a
saturated solution of hydrogen sulfide. For animal tissues, Sen used
cobalt-urea solns. in graded alcohols from 60 to 80%.

78 MICROSCOPIC TECHNIQUES

ALKALINE PHOSPHATASE*

The same staining technique for the visualization of alkaline phos-
phatase activity was developed independently and simultaneously,
oddly enough, by Gomori ( 1939) in Chicago and Takamatsu ( 1939)
in Japan. Their method was based on the finding that, when sections
of tissue were placed in an alkaline medium containing sodium
glycerophosphate, the sites of the enzymatic liberation of phosphate
could be determined if calcium ions were present to precipitate the
phosphate as it was formed. The deposit of calcium phosphate then
could be converted to a more easily visualized black precipitate of
cobalt sulfide or metallic silver. Gomori ( 1939) , Hepler et al. ( 1940) ,
Takamatsu (1939), and Kabat and Furth (1941) have employed
the von Kossa silver stain; but, as Bourne (1943) has indicated, it
is probably inferior to the cobalt stain used extensively in the latter
work of Gomori ( 1941a, 1943) .

The specificity of the stain for phosphatase has been demonstrated
by Gomori (1939, 1941a) and Kabat and Furth (1941), and in a
critical study later Danielli (1946b) claimed reliability for the
localizations obtained. Preformed insoluble calcium salts will give a
positive test and therefore these should either be removed by treat-
ing the sections, before incubation with substrate, with citrate buffer
of pH 4.5 to 5.0 for 15 min. (Gomori, 1946c), or control sections
stained to demonstrate the preformed salts should be compared to
the sections treated to visualize the enzyme reaction. Of course the
former is preferable.

Tissues too hard to be sectioned without decalcification present
a particular problem since phosphatase is destroyed by the usual
processes of decalcification. Kabat and Furth (1941) circumvented
this difficulty to some degree by the use of 10% diammonium citrate,
which they found could effect certain decalcifications without
damaging phosphatase. Bourne (1943) has proposed that small
pieces of bone tissue be fixed in 80% alcohol, treated with the sub-
strate medium and then with cobalt solution and sulfide to convert
the calcium phosphate precipitate to one of cobalt sulfide, and finally
subjected to decalcification with trichloroacetic acid. Cobalt sulfide
is insoluble in trichloroacetic acid and hence the decalcification can
be performed as a final step. Controls can be made by following the

*See Bibliography Appendix, Ref. 7.

ALKALINE PHOSPHATASE 79

same procedure but oniitting the treatment with substrate. Another
procedure for use with bone has been given by Bourne (1943) in-
volving the addition of 0.01% sodium alizarin sulfonate to Gomori's
substrate medium. The calcium phosphate produced is automatically-
stained red by the alizarin dye.

The use of magnesium ions to activate the phosphatase was intro-
duced by Kabat and Furth (1941) and is employed in the revised
method of Gomori ( 1946c) .

It is of interest to call attention to the different approach to the
staining technique for alkaline phosphatase that was brought for-
ward by Menten, Junge, and Green (1944). These investigators em-
ployed a reaction of the organic, rather than the phosphate, moiety
of the substrate to precipitate a reddish-purple dye at loci of phos-
phatase action. Employing calcium /j-naphthol phosphate as the sub-
strate, |3-naphthol liberated by the enzyme was made to react at
once with diazotized a-naphthylamine present in the substrate solu-
tion. While this procedure can undoubtedly be applied in many in-
stances, it would appear to offer no advantage over the Gomori
method, and, as Menten et al. readily admit, in its present form the
test is intricate and probably not well suited to routine use. Never-
theless, Yin (1945) employed this method for plant tissues in order
to avoid interference by preformed phosphates. Since the preformed
phosphates can be removed with citrate buffer (page 78) it would
appear that interference from this source need not be made a deter-
mining factor in the choice of a method.

Gomori Revised Method for Alkaline Phosphatase

SPECIAL REAGENTS

0% Acetylcellulose (Eastman's No. 4644) in acetone. Optional.

1-2% Cobalt Acetate, Chloride, or Nitrate.

Ammonium Sulfide Solution. A few drops of yellow ammonium

sulfide soln. to a Coplin jar of distilled water.
Substrate Medium, pH 9.4. (Will keep in refrigerator for months.)

Combine 25 ml. 2% sodium glycerophosphate, 25 ml. 2% sodium

barbital, 50 ml. distilled water, 5 ml. 2% calcium chloride, 2 ml.

2% magnesium sulfate, and a few drops of chloroform.

PROCEDURE

1. Place slices of fresh tissue, under 2 mm. thick, in chilled abso-

80 MICROSCOPIC TECHNIQUES

lute acetone and fix for 12-24 hr. in a refrigerator. Dehydrate at
room temperature in two changes of absolute acetone for 6-12 hr.
each time.

2. Optional step. To strengthen sections which tend to break up
when floated on the lukewarm water after sectioning, impregnate the
tissue with the acetone-acetylcellulose soln. 24 hr.

3. Drain off the fluid rapidly and place in two changes of benzol
for 30 min. each.

4. Embed in paraffin not over 56° up to 2 hr. To hasten the proc-
ess use 3 changes of paraffin, each for 20 min., and carry out the
second change in vacuo in a wide-mouth bottle with a one-hole
rubber stopper fitted with a glass tube. Connect the glass tube by
rubber tubing, passed through an air hole in the paraffin oven, to a
water aspirator via a safety bottle.

5. Cut sections 4-8 ju, thick, fioat them on lukewarm water
(30-35°), and mount on slides.

6. Let slides dry, place in the paraffin oven for 10 min. to melt the
paraffin, and run through xylol and alcohols to distilled water. Re-
move preformed mineral deposits as described on page 78.

7. Incubate the sections for 1-2 hr. at 37° in the substrate me-
dium.

8. Rinse with water, immerse in the cobalt soln. for 5 min., and
rinse well with several changes of distilled water.

9. Place in the diluted ammonium sulfide soln. for 1-2 min.

10. Wash well in water, counterstain if desired, dehydrate, and
mount.

Result. Sites of the phosphatase activity appear brown or black.

ACID PHOSPHATASE*

The staining method for alkaline phosphatase cannot be used for
acid phosphatase since calcium phosphate is soluble at a pH around
5, which is optimum for the action of the latter enzyme. Hence,
Gomori (1941b) employed lead ions in the substrate medium at pH
4.7 so that insoluble lead phosphate would be formed at the sites of
enzymatic activity. The lead phosphate was then converted either
to brown or black lead sulfide, or was stained a purplish-red with
acridine red. Wolf, Kabat, and Newman (1943) introduced several

*See Bibliography Appendix, Refs. 1 and 9.

ACID PHOSPHATASE 81

improvements in the procedure, and subsequently Gomori (1946c)
revised his original method. His experience with the technique led
Gomori (1946c) to state: "For some unknown reason, the staining
for acid phosphatase sometimes turns out patchy, occasionally even
negative, when it should be positive. This seems to happen especially
in cases when the pieces have been exposed to the temperature of
the paraffin oven for more than an hour, or when the temperature of
the oven is over 56°C."

The intensification of the acid phosphatase test by manganese ions
has been demonstrated by Moog (1943a), who found that a con-
centration of 0.01 M manganese sulfate gave the most satisfactory
results. This investigator recommends that the activator be added to
a clear portion of substrate medium just before use, and points out
that the incubation period may be approximately halved when the
reaction is accelerated by the manganese. Moog found that 4-5 hr.
was a satisfactory incubation period for tissues of the 6-day chick
embryo. No doubt a certain amount of trial and error must be ap-
plied to determine the proper incubation time for the particular
tissue under investigation. In a later study Moog (1944) found that
0.01 M ascorbic acid activated acid phosphatase and in some respects
appeared to have advantages over the action of manganese sulfate.

The application of the Gomori method to grains and sprouts was
made by Glick and Fischer (1945b). The modification in technique
necessitated for these tissues will be presented.

Gomori Revised Method for Acid Phosphatase in Animal Tissues

SPECIAL REAGENTS

5% Acetylcellulose (Eastman's No. 4644) in acetone. Optional.

£% Acetic Acid.

Ammonium Sulfide Solution. A few drops of yellow ammonium sul-
fide soln. to a Coplin jar of distilled water.

Substrate Medium, pH 5. Combine 30 ml. of 1 M acetate buffer
(100 ml. 13.6% sodium acetate, CHsCOONa.SHsO, -f- 50 ml. 6%
acetic acid), 10 ml. 5% lead nitrate, and 60 ml. distilled water,
and add slowly while stirring 30 ml. 2% sodium glycerophosphate.
Shake the mixture, let stand for a few hr., and store in a refrig-
erator. Before use, filter a small amount and dilute it with 2-3
parts distilled water.

82 MICROSCOPIC TECHNIQUES

PROCEDURE

1-6. Same as for alkaline phosphatase (pages 79 and 80).

7. Incubate the sections for 1-24 hr. at 37° in the substrate
medium.

8. Rinse well in distilled water, followed by 2% acetic acid, and
then in distilled water again.

9-10. Same as for alkaline phosphatase, page 80.

Result. Sites of the phosphatase activity appear brown or black.

Glick and Fischer Adaptation to Grains and Sprouts of Gomori
Method for Acid Phosphatase

SPECIAL REAGENTS

Substrate Medium. Combine the following, shake thoroughly,
centrifuge, and use the clear liquid: 4 ml. 0.1 ilf acetate buffer of
pH 5.1, 1 ml. 0.1 M lead nitrate, 0.6 ml. distilled water, and 0.4 ml.
3.2% sodium-a-glycerophosphate. Booth (1944) showed that the a
compound is hydrolyzed more rapidly than the (3 by wheat phos-
phatase, and Gomori (1941) found that the a compound has the
additional advantage that its lead salt is more soluble than that of
the /? at this pH value. Because it was easily available, the mix-
ture, containing 52% a and 48% fi of Eastman Kodak Co., was
used by Glick and Fischer ( 1945) .

2% Acetic Acid Solution.

Ammonium Sulfide Solution. 1 ml. to a Coplin jar of water.

PROCEDURE

/, Preparation of paraffin sections. A, Kernel sections:

1. Soak kernels in water about 7 hr.

2. For longitudinal sections, cut off a layer from both the crease
side and the opposite side of the kernel. For cross sections, cut o&
kernel just behind germ. This enables more efficient penetration of
liquids.

3. Let kernels stand overnight in absolute alcohol. In the morn-
ing change to a mixture of 1 vol. absolute alcohol -f- 3 vol. n-butyl
alcohol.

4. In the evening transfer to n-butyl alcohol.

ACID PHOSPHATASE 83

5. The following morning place in xylol, and let stand until
evening.

6. Transfer to a xylol-paraffin mixture containing just enough
xylol to keep the paraffin in soln. at room temperature and let stand
overnight. Tissuemat (Fisher Scientific Co.) gives better results than
paraffin. In a warm room the variety melting at 60-62° gives better
sections than the material having a lower melting point.

7. Place in a soln. of 1 vol. xylol + 2 vol. melted paraffin in
a 60° oven for 1-2 hr.

8. Infiltrate with melted paraffin for 2 hr, in the oven, then
change to fresh paraffin for 4 hr., and finally embed.

9. Cut sections 10 /x thick and mount on slides with aid of Mayer
albumin. (Combine 1 vol. filtered fresh egg white with 1 vol. glyc-
erol, and add a bit of camphor as a preservative.) Smear the liquid
in a thin film on a slide and rub with the finger, cover with water,
transfer section to the slide, place in oven for 5 min. at about 55° to
soften the paraffin and allow wrinkles to straighten out, drain off
water with a towel, and allow to dry for 2 hr. in the 55° oven. Store
mounted sections in refrigerator until ready for use.

10. Remove paraffin from sections with two changes of xylol fol-
lowed by two changes of absolute alcohol.

11. Dip slides into 0.5-1.0% collodion in alcohol-ether to cover
section with a protective film; harden film by dipping into 80%
alcohol, and wash with distilled water.

B. Rootlet and leaf section of the sprout:

1. Place in the following solns., for 1 hr. in each case, in the order
given :

a. 70% alcohol

b. 80% alcohol

c. 65 ml. 80% alcohol -^ 35 ml. n-butyl alcohol

d. 45 ml. 95% alcohol ~ 55 ml. n-butyl alcohol

e. 25 ml. absolute alcohol ^ 75 ml. n-butyl alcohol

f. n-butyl alcohol

g. xylol

2. Follow step 6 under A in the preceding part, allowing the
material to stand in the mixture for only i/^ hr.

3. Subject the material to three changes of melted paraffin in the

84 MICROSCOPIC TECHNIQUES

60° oven during the course of 1 hr. If air bubbles are present in the
leaves, apply suction to remove them.

4. Embed in paraffin colored red by stirring a few grains of Sudan
IV in the molten material. In uncolored paraffin it is difficult to see
the tissue in the sections.

5. Cut sections, mount on slides, remove paraffin, and protect
with collodion film just as in steps 9, 10, and 11 under A.

IT, Preparation of frozen sections. A, Kernel sections (rootlet and
leaf sections of the sprout are too fragile to permit satisfactory
frozen-section technique) :

1. Soak kernels 4-6 hr. in water.

2. Mount in a drop of water on freezing head of microtone.

3. Cut sections 15 fi thick, keeping knife cold with Dry Ice, and
transfer, with a needle cooled by Dry Ice, into 80% alcohol. (The
80% alcohol is used rather than water since, in the latter medium,
the starch endosperm disintegrates, and separates from the rest of
the section.)

4. Float the section onto a glass slide immediately. If wrinkled,
straighten section in a drop of 70% alcohol.

5. Dehydrate by covering section with five successive drops of
absolute alcohol, draining off after each drop is added.

6. Cover section with a small drop of 0.5-1.0% collodion soln.
and harden film by dipping slide in 80% alcohol. Wash in distilled
water.

Ill, Demonstration of enzyme activity:

1. Remove preformed mineral deposits by placing sections in dis-
tilled water for 24 hr. at room temperature. Citrate buffer could
probably be used too (page 000) .

2. Place sections in substrate medium at 37°, for the following
digestion periods:

Kernel, paraffin sections — Embryo, 1 hr.; non-embryonic part 30
min.

Kernel, frozen sections — Embryo, 15-30 min.; non-embryonic part,
5-10 min.

Rootlets, paraffin sections — 3 hr.

Epicotyl, paraffin sections — 24 hr.

3. Wash sections with three changes of distilled water, dip into
2% acetic acid, and wash well with distilled water.

ACID AND OTHER PHOSPHATASES 85

4. Place in ammonium sulfide soln. for 2-3 min.

5. Wash with several changes of distilled water, dehydrate in
95% alcohol for 2-3 min., and follow by 5 min. in absolute alcohol.

6. Clear in oil of thyme for 3-4 min., treat with three changes of
xylol. (Treatment with xylol should be brief since the black
precipitate indicating enzyme action is soluble to some degree in
xylol.) Mount in balsam.

Result. The phosphatase activity is visualized as a brown or
black precipitate.

OTHER PHOSPHATASES

Various investigators have studied the histological distribution
of phosphatases capable of hydrolyzing substrates other than those
commonly employed for the acid and alkaline phosphatases. This
work has been accomplished by simply substituting the new
substrates for the glycerophosphate usually used, and employing
the standard phosphatase procedures.

Wolf, Kabat, and Newman ( 1943) used ribonucleic acid and
glucose- 1-phosphate as additional substrates in their work on acid
phosphatase distributions, particularly in the human and guinea
pig nervous systems. Glick and. Fischer (1945b, 1946a) employed
adenosine triphosphate, thiamine pyrophosphate, and glucose- 1-
phosphate in a study of the enzyme distributions in wheat and parts
of the germinated grain. In investigations on the mouse duodenum,
Dempsey and Deane ( 1946) , and in work on the thyroids of various
species, Dempsey and Singer (1946), utilized adenylic acid, ribo-
nucleic acid, glucose- 1-phosphate, fructose diphosphate, and lecithin
as their additional substrates. Krugelis (1946) studied the phos-
phatases in the larval salivary glands of Drosophila and in various
organs of the mouse using adenylic acid, guanylic acid, cytidylic
acid, ribonucleic acid, desoxyribonucleic acid, and a depolymerized
form of the latter, as substrates.

With the staining technique it is difficult at times to define the
specificities of the various phosphatases which act on the different
substrates. For instance, both alkaline phosphatase and adenyl-
pyrophosphatase (adenosinetriphosphatase), which are known to
be two distinct enzymes, can act on adenosine triphosphate, as

86 MICROSCOPIC TECHNIQUES

Moog and Steinbach (1946) have emphasized. Accordingly, differ-
ences in their localizations or in their properties must be exploited
to enable their separate identification in tissue sections (Glick,
1946). From the work of Dempsey and Deane (1946) it would
appear that several phosphatases may coexist in the same cellular
location, and that their differentiation must depend on differences
in pH optima or other properties. When glucose- 1 -phosphate is
employed as the substrate, enzymatic liberation of phosphate might
occur either by phosphatase action or by the phosphorylase action
which converts the substrate to glycogen or starch; accessory
evidence would be required to determine which of these two enzymes
was being visualized by the staining reaction.*

While it does not offer rigorous proof, in some cases differentiation
between enzymes may be based on differences in their localization
as seen in stained sections. The presence in the nucleoli of the cells
of the wheat epicotyl of an enzyme capable of hydrolyzing adenosine
triphosphate, but not thiamine pyrophosphate (Glick and Fisher,
1946a), would suggest that these substrates are acted upon by
different enzymes. Likewise, the fact that a strong enzymatic
activity is found in the cytoplasm of cells in mouse tissues when
ribonucleic acid is used as the substrate, while only a slight reaction
is observed in the nuclei, and the reverse distribution is seen when
a depolymerized form of desoxyribonucleic acid is used, indicates
that separate enzymes are involved in the hydrolysis of these
substrates (Krugelis, 1946). Furthermore, the approximately equal
activities in both cytoplasm and nucleus toward glycerophosphate
as substrate might be taken as an indication of the presence of a
third enzyme in these cells, as Krugelis has pointed out. Another
example is to be found in the differences in the localizations of the
enzymes acting on glucose-1-phosphate and fructose diphosphate
when the mucosa of the mouse duodenum is studied (Dempsey and
Deane, 1946). Other cases might be cited to illustrate the same
general point.

ZYMOHEXASE (ALDOLASE plus ISOMERASE)

Aldolase converts hexose diphosphate to both dihydroxyacetone
phosphate and phosphoglyceraldehyde; isomerase catalyzes equi-

* See Bibliography Appendix, Ref. 19.

ZYMOHEXASE 87

librium between the two. Together these two enzymes are referred
to as zymohexase.

Allen and Bourne ( 1943) adapted the microscopic technique for
phosphatase (page 78) to this enzyme system, whose distribution
they studied in skeletal, heart, and smooth muscle tissue. By
incorporating iodoacetic acid into their substrate media, they
prevented further enzymatic breakdown of the triose phosphates.
The distinct difference in localization of zymohexase and alkaline
phosphatase precluded the possibility of confusing the two; however,
the phosphatase activity could be selectively blocked by fluoride.
Allen and Bourne utilized the fact that the triose phosphates formed
by the zymohexase action will spontaneously liberate inorganic
phosphate at room temperature in alkaline solution. The phosphate
could then be precipitated, and finally visualized in the manner of
Gomori (page 78). It was observed that sections which had been
infiltrated with paraffin lost their enzyme activity and, accordingly,
frozen sections were employed.

Allen and Bourne Method for Zymohexase

SPECIAL REAGENTS

0.1 M Sodium lodoacetate. Neutralize 1.86 g. iodoacetic acid with
1 N sodium hydroxide to bromothymol blue and dilute to 100 ml.

0.1 M Sodium Fluoride.

2% Cobalt Chloride (C0CI2.6H0O).

Ammonium Sulfide Solution. Dilute 1 ml. yellow ammonium sulfide
to 50 ml. with distilled water. Prepare fresh before use.

Magnesia Mixture. Dissolve 5.5 g. magnesium chloride
(MgCl2.6H20) and 7.0 g. ammonium chloride in 35 ml. 5 N am-
monium hydroxide. Filter after 1 hr. and add 60 ml. 4 A^" am-
monium hydroxide to the filtrate.

Purified Sodium Hexose Diphosphate. Formula I: mix 40 ml. 4%
sodium hexose diphosphate ( prepared by treating the calcium salt
with sodium oxalate) and 20 ml. magnesia mixture, and filter off
precipitated phosphate after 30 min. Formula II: mix 20 ml. of
4% sodium hexose diphosphate with 20 ml. of magnesia mixture
and 20 ml. of distilled water; filter after 30 min. as above.

Substrate A. Combine 10 ml. of the purified salt soln. (Formula I)
with 1.7 ml. of 0.1 M sodium iodoacetate and 5 ml. distilled water.

88 MICROSCOPIC TECHNIQUES

Substrate B. Same as A, but use 3.3 ml. water and 1.7 ml. of 0.1 M
sodium fluoride in place of the 5 ml. water. (Substrate B was used
to prevent hydrolysis by phosphatase through the action of the
fluoride. Actually Allen and Bourne found that it was not needed
in their work since no phosphatase action was observed in their
particular experiments.)

Substrate C. Combine 15 ml. of the purified salt soln. (Formula II)
with 2.5 ml. of the sodium iodoacetate and 7.5 ml. water.

PROCEDURE

1. Fix tissue in 80% alcohol for 24 hr.

2. Wash in water for 5-10 min.

3. Prepare frozen sections and place them in either substrate
A or C for 1-2 hr. at 37°. (An extraneous dark precipitate forms
on the surface of some of the sections with substrate A, but not
with C, presumably because of the higher dilution of the substrate
in the latter.)

4. Treat the sections with the cobalt chloride soln. for several
hr. and then with the ammonium sulfide soln. for 10 min.

5. Dehydrate in alcohols, clear in xylol, and mount in balsam.

6. Prepare control sections to demonstrate preformed phosphate
by omitting the treatment with substrate in step 3 and proceeding
with steps 4 and 5.

Result. The formation of a brownish-black precipitate indicates
zymohexase activity.

LIPASE

Gomori (1945b) adapted the principle of his method for the
demonstration of phosphatases in tissue sections to the localization
of lipase. The great difficulty encountered in previous attempts has
been to find a substrate that is water soluble and whose acid split-
product could be precipitated by some ion having no adverse effect
on the enzyme. The ordinary esters of mono- and dicarboxylic acids
do not meet both of these requirements. Gomori was able to circum-
vent the difficulty by the use of some new long-chain fatty acid
esters of hexitans and hexides in which most of the hydroxyl groups
are etherified. These compounds were developed by Atlas Powder

LIPASE 89

Co., and are known as "Tweens." Tween 40 and Tween 60, employed
by Gomori, are described by Atlas Powder Co. as "sorbitan mono-
palmitate polyoxyalkylene derivative" and "sorbitan monostearate
polyoxyalkylene derivative," respectively. Gomori states that these
substrates were found to be hydrolyzed by pancreatic lipase at a
rate about half that of olive oil. In the presence of 0.2% sodium
taurocholate an intensification of the reaction in pancreatic tissue
was noted, while in all other tissues, enzyme inhibition was observed.
In a later publication Gomori (1946a) reported that "Product 81"
of Otiyx Oil and Chemical Co. could also serve as a lipase substrate
for histochemical purposes. This compound is a stearic acid ester
of "comparatively short-chained polyglycols." The only shortcoming
observed by Gomori to the use of these substrates is an occasional
failure to obtain proper counterstaining with hematoxylin, especially
after use of the "Tweens" and more rarely with "Product 81." The
method as finally given by Gomori (1946c) will be described.

Gomori Revised Method for Lipase

SPECIAL REAGENTS

5% Acetylcellulose (Eastman's No. 4644) in acetone.

1-2% Lead Nitrate.

Ammonium Sulfide- Solution. A few drops of yellow ammonium
sulfide soln. to a Coplin jar of distilled water.

Substrate Medium. Stock solution I: combine 150 ml. glycerol,
60 ml. of 10% calcium chloride, 50 ml. M/2 maleate buffer pH
7 to 7.4 (dissolve 5.8 g. maleic acid in 94 ml. of 4% sodium
hydroxide and 6 ml. water), and distilled water to make 1000 ml.
Stock solution II: 5% Tween 40 or 60 or "Product 81." Add
merthiolate to 0.02% in each stock soln. and store in refrigerator;
the solns. may be used for many months. Before use, add 2 ml.
stock soln. II to 50 ml. stock soln. I.

PROCEDURE

1-6. Same as for alkaline phosphatase (pages 79 and 80).

7. Incubate the sections in the substrate medium for 6-12 hr.
at 37°.

8. Rinse with distilled water and transfer to the lead nitrate
soln. for 10 min.

90 MICROSCOPIC TECHNIQUES

9. Rinse in repeated changes of distilled water and immerse in
the diluted ammonium sulfide soln. for 1-2 min.

10. Wash well in water, counterstain lightly with hematoxylin
and eosin, dehydrate in alcohols, clear in gasoline or tetrachloro-
ethylene, and mount in Clarite dissolved in the same liquid. (Toluol
or xylol causes fading of the stain.)

Result. Sites of lipase activity appear golden brown.

PEROXIDASE

Most of the various microchemical methods for the histological
localization of peroxidase activity are based on the oxidation of
benzidine. The methods of McJunkin ( 1922) , designed for use with
human tissues, and Armitage ( 1939), developed for examining blood
and bone marrow smears, have been chosen for presentation here
since they are the most recent and seem to be the best. Peroxidase
actually occurs most abundantly in plants, but the methods appear
to have been worked out for animal tissues or cells exclusively.
However, there appears to be no reason why these methods cannot
be adapted to plant material as well.

McJunkin Method for Peroxidase in Tissue Sections

SPECIAL REAGENTS

Benzidine Reagent. Dissolve 100 mg. benzidine in 25 ml. 80%
methanol and add 2 drops 3% hydrogen peroxide. Dilute with
1-2 vol. distilled water before using. Store in the dark.

PROCEDURE

1. Place formalin-fixed tissue, in pieces 1 mm. thick, in 70%
acetone for 1 hr. followed by pure acetone for 30 min., benzol for
20 min., and melted paraffin 20 min.

2. Cut sections 3.5 to 5.0 [i, fix to slides with albumin, and dry
overnight at room temperature.

3. Remove paraffin by placing in benzol for 20 sec. and acetone
for 10 sec.

4. Plunge in water for a few sec. and remove excess water. Apply
the benzidine reagent for 5 min. and transfer to water for 5 min.

5. The sections may be stained with Harris hematoxylin for 2
min., rinsed in water 1 min., and stained with 0.1% eosin for 20 sec.

PEROXIDASE AND DOPA OXIDASE 91

6. Dehydrate in 95% alcohol for 30 sec, and absolute alcohol
for 5 sec.

7. Clear in xylol and mount in balsam.

8. Run controls in which the benzidine is omitted.

Result. Peroxidase manifests itself by an initial blue color which
changes to brown. Diffusibility, particularly of the color produced,
can be expected to interfere with proper localization of the enzyme.

Armitage Method for Peroxidase in Blood or Bone
Marrow Smears

SPECIAL REAGENTS

Fixing Solution. 10% Formalin in 96% alcohol. Prepare the soln.

just before using.
Benzidine Reagent. Dissolve 750 mg. benzidine in 500 ml. 40%

alcohol, filter, add 0.7 ml. 3% hydrogen peroxide, and shake

before using. If stored in the dark, the reagent will be good for

months.

PROCEDURE

1. Fix the smear in the alcoholic formalin.

2. Cover the material with the benzidine reagent for about 2 min.
if the smear is fresh, and up to 20 min. if it is old.

3. Wash in 40% alcohol until yellow granules appear in the
leucocytes.

4. Dehydrate in absolute alcohol and dry at about 37°.

5. A counterstain of dilute Giemsa or dilute Leishman stain may
be applied for 30 min. followed by washing in water, blotting, and
drying.

Result. The appearance of yellow granules is a positive test for
peroxidase.

DOPA OXIDASE

The enzymatic oxidation of 3,4-dihydroxyphenylalanine — or dopa
— has been applied, histologically, to the identification of melano-
blasts, since these appear to be the seat of the oxidase activity and
the conversion of dopa to melanin results in their becoming black-
ened. Bloch's earlier work has been adapted by Laidlaw (1932) and
Laidlaw and Blackberg ( 1932 ) to the demonstration of dopa oxidase

92 MICROSCOPIC TFX'HNIQUES

activity in histological preparations. Sharlit et al. (1942) have
pointed out that the method for demonstrating dopa oxidase may
of itself, in the absence of substrate, cause an increase in melanin.
This makes it necessary to run suitable control experiments. The
reaction may be hastened by employing a buffer of a little higher
pH, or retarded by shifting toward the acid side.

Laidlaw Melliod for Dopa Oxidase

SPECIAL REAGENTS

Dopa Stock Solution. Dissolve 0.3 g. dopa (Hoffmann-La Roche,
labeled "for Bloch's dopa reaction") in 300 ml. cold distilled
water. Store in a refrigerator and discard when a distinct red color
has developed.

Buffered Dopa Solution (pH 7.4) . Add 2 ml. potassium dihydrogen
phosphate (9 g. KH2PO4/I.) and 6 ml. disodium hydrogen phos-
phate (11 g. Na2HP04.2H20/l.) to 25 ml. dopa stock soln. Filter
through fine paper.

Buffer Solution for Control Experiment. Replace the 25 ml. dopa
soln. with an equal vol. distilled water in the buffered dopa soln.
above.

PROCEDURE

1. Prepare frozen sections of fresh tissue. (It is stated that tissue
hardened in 5% formalin for 2-3 hr. may be used.)

2. Rinse in distilled water for a few sec. and transfer at once to
the buffered dopa soln. At 30-37° the soln. becomes red in about
2 hr. and sepia brown in 3-4 hr. Do not let the sections remain in
the soln. once it becomes sepia colored since overstaining may result.
Examine from time to time under a microscope to determine the
proper intensity of staining. It is good practice to change to a fresh
dopa soln. after the first 30 min.

3. Wash sections in distilled water, dehydrate, and counterstain
w^ith alcoholic cresyl violet or methyl green-pyronine.

4. Clear and mount in balsam.

5. Run controls by treating tissue as above with the substitution
of buffer soln. for the buffered dopa soln. in step 2.

DOPA AND AMINE OXIDASE 93

Result. Dopa oxidase is indicated by blackening in the sections.
Leucocytes and melanoblasts appear grey or black due to their
dopa oxidase content. Melanin maintains its natural yellow-brown
color, and collagen appears colorless or pale grey.

AMINE OXIDASE

Oster and Schlossman ( 1942) developed a histochemical method
for the demonstration of amine oxidase based on the detection of
the aldehyde formed as the product of amine oxidation. The fuchsin-
sulfurous acid reagent of Feulgen was used for the visualization oi
the aldehyde (page 65). Naturally occurring aldehydes and "plas-
mal" are prevented from interfering with the test by binding them
with bisulfite prior to the application of the tyramine substrate
solution. The diffusibility of the color produced subjects the locali-
zations which may be observed to criticism.

Oster and Schlossman Method for Amine Oxidase

SPECIAL REAGENTS

2% Sodium Bisulfite Solution.

Substrate Solution. 0.5% tyramine hydrochloride in M/15 phos-
phate buffer of pH 7.2.
Control Solution. Omit the tyramine in the substrate soln.
Feulgen Reagent. See page 67.

PROCEDURE

1. Place frozen sections of fresh tissue in 2% bisulfite solution
at 37° for 24 hr. Wash thoroughly and test some of the sections
with the fuchsin-sulfurous acid reagent — the test should be negative
(no color) indicating all free aldehyde has been bound,

2. Incubate sections in the substrate solution for 24 hr. at 37°.
Run parallel controls with the control solution.

3. Immerse in fuchsin-sulfurous acid reagent. -

4. Examine sections when the rapidly formed blue color seems
to be maximum.

Result. Regions of enzymatic activity appear blue, offering a
distinct contrast to the reddish-purple given by "plasmal" (see
page 65).

94 MICROSCOPIC TECHNIQUES

CYTOCHROME OXIDASE

Tests for cytochrome oxidases have been adapted to histochemical
work and a discussion of them has been given by Lison ( 1936, pages
269-290). The enzyme has been referred to as "nadi oxidase" and
"indophenol oxidase/' but Keilin and Hartree ( 1938) have made it
clear that it should be called "cytochrome oxidase" since its catalytic
effect applies to the oxidation of reduced cytochrome. In the
presence of cytochrome c, cytochrome oxidase effects the oxidation of
a mixture of p-aminodimethylaniline and a-naphthol (nadi reagent)
to indophenol, or of p-phenylenediamine to the diimine. The diffusi-
bility of the colored compounds produced must be considered with
reference to localizations of the enzyme in tissue.

In order to check whether a nadi reaction is being given by cyto-
chrome oxidase or some other factor, Moog (1943b) exposed fresh
tissue (chick embryo) to a 0.005 M azide solution in acidified
physiological saline (pH 5.8) for 3 min., and then transferred it to
freshly prepared nadi reagent containing 0.005 M azide. Azide
specifically inhibits cytochrome oxidase. As a control of the possi-
bility that indophenol blue might be reduced to the leuco form as
fast as formed, Moog also placed the tissue in 0.003 M phenylurethan
in saline for 3 min. to saturate the reducing systems, and then trans-
ferred to the nadi reagent containing 0.003 M phenylurethan. The
reagent used by Moog was prepared by combining, just before use,
equal parts of 0.01 M /^-aminodimethylaniline in 1 % sodium chloride,
0.01 M a-naphthol in 1% sodium chloride, and 0.066 M phosphate
buffer. The reaction was carried out at 38° for the interval required
to attain a standard coloration (5-14 min.). Under the conditions
employed, identical results were obtained at pH 5.8 and 7.2.

Graff Method for Cytochrome Oxidase in Fixed Tissue
("M. Nadi Oxidase")

SPECIAL REAGENTS

a-Naphthol Solution. Boil 1 g. a-naphthol in 100 ml. distilled water
and add 25% potassium hydroxide dropwise until the melted
a-naphthol is dissolved. Store in the dark; keeps for at least 1
month.

1% p-Aminodimethylaniline or Its Hydrochloride. Boil to dissolve
solid in the water. Store in the dark; may be used for 2-3 weeks.

CYTOCHROME OXIDASE 95

The hydrochloride is favored because it is more stable.
Nadi Reagent. Prepare just before using by combining equal vol.

of the a-naphthol and p-aminodimethylaniline solns. and filtering.
Strong Ammonium Molybdate Solution or Dilute Lugol Solution.

Concentration not stated.
Dilute Lithium Carbonate Solution. Concentration not stated.

PROCEDURE

1. Fix tissue for two hr. in formalin vapor or in a mixture of
10 ml. formalin and 40 ml. 96% alcohol.

2. Prepare frozen sections and place them on slides which are
then laid in a thin layer of nadi reagent in a petri dish. Oxygenation
of the fluid is effected by careful agitation. After 1-5 min., rinse in
water and examine.

3. Make the color more permanent by treating for 2-3 min. with
dilute Lugol soln. The Lugol soln. converts the blue granules to
brown. Washing sections in dilute lithium carbonate restores the
blue. Strong ammonium molybtlate soln. has been used instead of
Lugol soln.

4. Counterstain with Bismark brown, safranine or alum carmine
and mount in glycerin or glycerin jelly.

Result. Cytochrome oxidase is supposed to be indicated by the
blue coloration.

Graff Method for Cytochrome Oxidase in Fresh Tissue
("G. Nadi Oxidase")

The pH of the nadi reagent must be adapted to the requirements
of the particular material under investigation. Lison (1936, page
274) states that the pH range 7.8 to 8.2 is most suitable for animal
tissues and 3.4 to 5.9 for plant material.

SPECIAL REAGENTS

a-Naphthol Solution. Prepare a 10% alcoholic soln. and just before

use dilute 100 times with distilled water.
0.12% p-Aminodimethylaniline Hydrochloride. Store in the dark.
Nadi Reagent. Prepare just before using by combining equal vol.

of the diluted a-naphthol soln. and the p-aminodimethylaniline

soln.
Buffered Nadi Reagent. Mix the reagent with the suitable buffer

96 MICROSCOPIC TECHNIQUES

(acetate, phosphate, glycine, und carbonate buffers have been
employed) in the respective proportion of 50:10 or 5:20.
5% Potassium Acetate Solution.

PROCEDURE

1 . Prepare frozen sections directly from very fresh tissue.

2. Repeat step 2 in the preceding cytochrome oxidase method,
but wash sections with physiological saline instead of water.

3. Nuclei may be stained with lithium carmine.

4. Examine under a microscope with the section covered with
potassium acetate soln. Permanent preparations cannot be made.

Result. The blue or blue-violet color is also produced in this case.

?9

Loele Method for "a-Naphthol Oxidase

SPECIAL REAGENTS

Naphthol Reagent. Add 10% potassium hydroxide dropwise to a
little a-naphthol in a test tube until the a-naphthol is dissolved.
Add 200 ml. distilled water, and after 24 hr. the reagent may be
used for about 3 weeks.

PROCEDURE

1. Prepare frozen sections of formalin-fixed tissue.

2. Treat sections with the a-naphthol reagent and observe effect
under the microscope within a few minutes.

Result. Violet or black granules which soon disappear are sup-
posed to be indicative of a-naphthol oxidase.

SUCCINIC DEHYDROGENASE

Semenoff ( 1935) gave a method for the localization of succinic
dehydrogenase in tissue sections which depends on the reduction of
methylene blue. The diffusibility of the dye should obviate the
possibility of good localizations by this method.

Semenoff Method for Succinic Dehydrogenase

SPECIAL REAGENTS

Substrate Medium. To 2 ml. 0.05% methylene blue add 2 ml.

SUCCINIC DEHYDROGENASE 97

10% sodium succinate and make up to 10 ml. with A//15 phos-
phate buffer, pH 7.6 to 8.0.
Control Medium. Omit the succinate in the substrate medium.

PROCEDURE

1. Prepare fresh frozen sections.

2. Treat sections 10-15 min. with the substrate medium under
a cover slip, taking care to avoid air bubbles. Seal edges of cover
slip with paraffin to exclude air.

3. Observe under microscope and compare with section in control
medium treated in the same fashion.

Result. Fading of dye characterizes the enzyme activity.

///. PHYSICAL METHODS

A. FLUORESCENCE MICROSCOPY

The detection and localization in tissues and cells of certain
substances by virtue of their flourescent properties when subjected
to ultraviolet irradiation is finding increasing application. Sub-
stances investigated in this manner include naturally occurring
compounds, such as vitamin A, and others introduced into organisms
for experimental purposes, such as 20-methylcholanthrene. The
fluorescence exhibited in tissues or cells may be "primary," i.e.,
produced directly by certain compounds, or "secondary," i.e., result-
ing from treatment with so-called "fluorochromes," fluorescent sub-
stances taken up selectively by particular cellular structures having
no fluorescence of their own. For the most part, the use of fluoro-
chromes is limited to purely morphological studies without regard
to chemical nature and hence need not be discussed here, except as
applied to lipids (page 105). However, for those who may be in-
terested, lists of fluorochromes and their properties may be found in
Haitinger (1938), Jenkins (1937), and Metcalf and Patton (1944).
General discussions of fluorescence microscopy are to be found in
Sutro ( 1936) , Jenkins ( 1937) , Ellinger ( 1940) , Simpson (in Cowdry,
1943, pages 76-78) , and Metcalf and Patton ( 1944) . The books of
Haitinger (1938), Radley and Grant (1939), and Pringsheim and
Vogel (1946) are useful for reference.

1. Apparatus

The set-up of the fluorescence microscope is shown in Figure 3.
Ultraviolet radiation having a high intensity in the range 300-400
m/j. is produced by means of a carbon arc or one of the mercury vapor

99

100

MICROSCOPIC TECHNIQUES

lamps (.4) such as those manufactured by Hanovia Chemical Co.
or the H3 or H4 lamps of General Electric Co. or Westinghouse
Electric Co.

The ultraviolet radiation is passed through a filter (F) to screen
out visible light. A variety of filters may be used for this purpose.

The Corning color glass filter No. 584
( new No. 5840) may be used in combina-
tion with a 5-10% copper sulfate solu-
tion, to which a drop or two of sulfuric
acid has been added, contained in a
quartz or other cell transmitting ultra-
violet rays. The copper sulfate solution
may be replaced by a Corning glass filter
No. 428 (4308), but the latter does not
remove the heat rays as well as does the
solution, and cannot be adjusted to com-
pletely absorb red light. Other filters
that may be employed are a combination
of the Shott glass filters UG2 and BG14,
the Corex filter of nickel oxide glass, or
the Uvet glass filters with a copper sul-
fate solution.
The filtered ultraviolet radiation practically free of visible light
is directed into a substage condenser (QC), made of quartz or ultra-
violet-transmitting glass, by means of a reflector (R) consisting of
either a quartz prism, a polished mirror of aluminum-magnesium
alloy, or, if the ultraviolet intensity is great, the usual plane micro-
scope mirror. Of course, w^hen the apparatus is aligned either
vertically or horizontally on a single optical axis, the reflector is
omitted. In those instances in which the fluorescence is generated
by the longer wavelengths of ultraviolet radiation, as is the case for
vitamin A, it is often possible to employ the ordinary substage
condenser, rather than one made of quartz or special glass, since the
usual grade of optical glass does not absorb much of the radiation
in this range. The condenser may be eliminated entirely if radiation
of lesser intensity can be used. Of the three most common forms of
condensers, the aplanatic gives the best results, although the Abbe
is generally quite satisfactory; achromatic condensers reduce the
intensity of the radiation due to the absorption of their many lenses.

Fig. 3. Diagram of apparatus
for fluorescence microscopy.

FLUORESCENCE MICROSCOPY 101

Metcalf and Patton (1944) suggest the use of a drop of water or
petrolatum on the top of the condenser to serve as a connecting
fluid between condenser and slide in order to obtain illumination of
high intensity when objectives of twenty times magnification, or
higher, are used. With low-power objectives, they point out that it
is necessary to remove the top lens or lenses of the condenser so
that the field can be properly illuminated. Other investigators have
employed sandalwood or Shillaber oil between the slide and con-
denser.

The specimen is mounted on a slide (S) made of an ultraviolet-
transparent glass such as the Corex D slide of Corning Glass Co.
However, Metcalf and Patton ( 1944) have found that ordinary
glass slides of 1.2 to 1.5 mm. thickness may be used when the inten-
sity of the ultraviolet radiation is great. A nonfluorescing medium
must be used for moimting the specimen; glycerol or mineral oil
was recommended by Simpson ( in Cowdry, 1943, pages 76-78) ; but
Popper (1944) reported a disturbing fluorescence from glycerol
(although others beside Simpson have found no difficulty with it)
and the use of mineral oil is limited to substances that will not
dissolve in it, e.g., for vitamin A studies Popper ( 1944) used water
as the mounting medium. In some instances, petrolatum serves as
a good temporary mount, and, for permanent mounts, isobutyl
methacrylate (du Pont), suggested by O'Brian and Hance (1940),
is probably the best. With immersion objectives, sandalwood or
Shillaber oil may be employed as the immersion medium. No special
objectives or oculars are required; however, Jenkins (1937) has
pointed out that some of the older objectives contain balsam that
gives rise to its own fluorescence, and in these cases a darkfield
stop must be used in the condenser to prevent the entrance of direct
ultraviolet rays into the objective.

A filter (EF) that excludes ultraviolet, and passes visible rays,
is placed on the ocular. Either the Corning filters No. 3389 or 3060,
the Leitz No. 8547A, the Bausch and Lomb or the Zeiss Euphos
filter, or a circle of Wratten 2A gelatin filter cut to fit inside the
eyepiece may be used. INIetcalf and Patton (1944) recommend a 5%
solution of sodium nitrite contained in a plane-sided glass cell 5-10
mm. in optical depth which may be conveniently placed on the dia-
phragm of the ocular or, better still, on the diaphragm of the
microscope tube.

102 MICROSCOPIC TECHNIQUES

2. Preparation of Tissues

The preparation of sections from frozen dried material has the
advantage that soluble or diffusible constituents will have no chance
of being lost or displaced from their original sites. However, while
this is the method of choice, paraffin sections of formalin-fixed
tissue have been employed with success in certain instances, although
neither celloidin nor gelatin sections can be used since these media
give rise to fluorescence.

When paraffin sections are employed, fixation is usually carried
out in 5-10% formalin for not longer than 24 hr. The sections are
cut 7-8 /x thick; egg albumin has been used to make them adhere to
the slides. The paraffin is removed by immersion in xylol for 30
min. and the sections are dried at room temperature. In this form
they have been kept for months, and may be examined without
further treatment. All reagents and materials should be the purest
obtainable to avoid adventitious fluorescence of contaminants.

3. Photomicrography

Photomicrographs of fluorescing preparations can be made if
certain precautions are taken. Popper and Elsasser (1941) found
that, in the photomicrography of vitamin A fluorescence, it is pref-
erable to use film of maximal daylight sensitivity. The Fluorapid
film of Agfa-Ansco Corp. is particularly well suited for the purpose
(Fig. 4C,D) . Kodachrome film can be used when the tissue is rich in
vitamin A, but the lower sensitivity of this film limits its value. In
general black-white film is preferable to the color variety.

With substances of fading fluorescence, such as vitamin A, the
time employed for focusing must be kept to a minimum even at
the expense of the sharpness of the picture. The exposure itself must
be short (a maximum of 30 sec, regardless of magnification, in the
case of vitamin A) in order to obtain greater contrast between the
fluorescence and the background. Only one exposure can be made
as shown in Figure 4A,B.

When the fluorescence does not fade, Kodachrome film can give
excellent results. The exposure time has to be determined by trial
in each case, usually falling in the range of 1-15 min. with an
average of about 2 min., according to Metcalf and Patton (1944).
The loss in intensity with greater magnifications makes it impracti-

FLUORESCENCE MICROSCOPY

103

cal to employ magnifications exceeding 500 times. Metcalf and
Patton ( 1944 > also report that with black-white film having a
Weston rating of 50, exposures of from 2 sec. to 5 min., with an
average time of about 10 sec, are required with a 35 mm. camera.
In all fluorescence photomicrography the dark-field stop on the

Fig. 4. A, Human liver showing collagenous fibers in the periportal field
and vitamin A fluorescence in the liver cells. B, Second exposure of the same
field; the vitamin A fluorescence has faded. C, Rat liver photographed with
sensitized film (Fluorapid) ; a large amount of vitamin A fluorescence in the
Kupffer and the liver cells is evident. D, Picture of a liver taken with normal
ultraspeed film. From Popper and Elsasser (1941)

condenser and the filter placed on, or in, the eyepiece or microscope
tube to screen out ultraviolet rays must be used. Otherwise fogging
may occur from the stray radiation. Metcalf and Patton ( 1944)
especially recommend the sodium nitrite filter for color photog-
raphy. In order to switch from ultraviolet to visible illumination
a piece of opal or ground glass is substituted for the filter placed in
front of the light source.

104 MICROSCOPIC TECHNIQUES

4. Characterization of Substances

Direct Observation of Fluorescence

Vitamin A. Particularly intensive work has been done on vita-
min A, starting with the work of von Querner (1935), Hirt and
Wimmer (1940), and others, and continuing in greatly expanded
scope and development with the research of Popper (1944) and
associates. The fading green fluorescence of vitamin A in tissue
sections has been found to run parallel with the results of chemical
determination (Popper and Elsasser, 1941). The fading green fluo-
rescence is characteristic of vitamin A], found in salt water fish, and
a slowly fading pale yellow-brown fluorescence characterizes vitamin
A2, found in fresh-water fish. An admirable review of the studies
made on vitamin A distribution in the tissues of animals and man,
both in normal and pathological states, has been presented by Popper
(1944). He pointed out that carotene may easily be differentiated
from the A vitamins by its very slowly fading green fluorescence
which is apparent only in higher concentrations, and that the bio-
logically inactive anhydro ("cyclized") vitamin A may be recog-
nized by its dark brown fluorescence which gradually becomes a dull
green and finally fades out entirely. Volk and Popper (1944a) re-
ported the existence of a factor in biological fluids, particularly
plasma, and in organ emulsions that delays the disappearance of
vitamin A fluorescence in tissue sections.

Riboflavin. EUinger and Koschara (1933), von Euler et al.
( 1935) , Hirt and Wimmer ( 1939a) , and Metcalf and Patton ( 1942)
utilized the yellowish-green fluorescence of riboflavin for its identi-
fication in tissues. According to Ellinger (1938), and confirmed by
Metcalf and Patton (1942), another form of riboflavin exists (prob-
ably bound to another compound) which gives a yellow-orange
fluorescence. INIetcalf (1943> subsequently concluded that in the
American roach, Periplaneta americana L., the bound riboflavin is
converted to the free form in vivo by the injection of pantothenic
acid or thiamine. With the latter compound the conversion proceeds
more slowly.

Other Vitamins.* Attempts have been made to characterize, and
to determine the distribution of, other vitamins by their fluorescent
properties. Hirt and Wimmer ( 1939b) investigated nicotinic acid
and its amide which they claimed gave a stable yellow fluorescence.

* See Bibliography Appendix, Ref . 29.

FLUORESCENCE MICROSCOPY 105

They reported that in the dry state the amide has a weaker fluores-
cence than the acid, but that in a 1% aqueous solution the reverse
is found. EHinger (1940) was not able to observe fluorescence with
purified solutions of these compounds, and he also disagrees with
Hirt and Wimmer (1939b) that ascorbic acid can be detected
microscopically by fluorescence. The histochemical opportunities of
studying vitamin K by means of its well-known fluorescence are
obvious. One may expect that studies of this nature will be made in
the future.

Lipids. During the course of his work on vitamin A, Popper
(1941) also studied the histological detection of lipids by means
of the fluorchromes: methylene blue, thioflavin S, rose bengal
magdala red, and phosphine 3R. The last stain appeared to be the
best. Further examination revealed that fatty acids, soaps, and
cholesterol are not made apparent bj^ phosphine 3R which, however,
does visualize neutral fat as a silver-white fluorescence on a brown
background (Volk and Popper, 1944b; Popper, 1944). The advan-
tage claimed for this method is that, because of the water solubility
of the dye, more and finer droplets of lipid can be detected than
would be possible by the usual stains. Popper (1944) recommends
the use of a 0.1% aqueous solution of phosphine 3R {Pfaltz and
Bauer) for 3 min. on frozen sections of tissue.

Pigments. The fluorescence technique has been applied to studies
of certain biological pigments. Thus the red fluorescence of porphy-
rins has been employed in histological studies of these compounds
(Lison, 1936, page 256; Ellinger, 1940; Dobriner and Rhoads, 1940;
Grafflin, 1942) . Chlorophyll has been localized microscopically in
plant tissues by Tswett (1911) and Wilschke (1914) by means of
its red fluorescence. The fluorescent properties of bile pigments in
the presence of zinc acetate, and of uropterin, might be adapted to
microscopic studies of these pigments.

Carcinogenic Hydrocarbons. Investigations of carcinogenic
hydrocarbons in tissues have made use of the fluorescence of certain
of these compounds. Graffi ( 1939, 1940) investigated the distribu-
tions in normal and tumor cells of pyrene, benzpyrene, anthracene,
dibenzanthracene, and methylcholanthrene, while Gunther (1941)
and Doniach et. al. ( 1943) confined their studies to benzpyrene, and
Simpson and Cramer (1943-1945) explored the histological localiza-
tion of 20-methylcholanthrene in skin.

106

MICROSCOPIC TECHNIQUES

PQ
<

o

d

o

'3

o

CO

OOO

ooooo

oo

oo oo

oo

oo

OOO

!-<' ^'

c

^

Xi

o

^•bi^

'T ^- bb bhi^

1 1

a 44

bb

"? bb

bbi

i; bb C.

lO

O

>> d tJD

bijD-d-d >i

>> >>

'O-^ >.>>

XiO

-3-D

bC^'O

"^

05
1-H

04

fl

&'tjb>;

ti . . . bb

1 t-l t< tH 1

^'^

C u . .

t-T

i

P*^ 1 1 1 ^

1 1 II

1

bb>:,

f'f-'f

4)

t3

n

ti dbb

-d >) bb bb-d

bb >i

t^. bb bb bb

bbo

>> bb

bb bb >>

'3

a

1 1 r>^

. t," C !m" .
bC^nD,^ bC

>>^

bb

^ bb >,>>

>>

bb >>

>>>.^

ffi

o

S

^ — ; '

1 1 1 1 1

1 l_

"^ '. '. '.

l_

1. I_

'. '. t:

lO

-CiX> bb

>>>>>.>>>>

bb >-.

■d >i bb bb

bbo

>^, bb

bO bC42

d

.

,

bb

,

■§

is?:

bbj2 >^ bb
III 1

t-?'

^ bb bb &
P>-1 1 II

bb

i_ 1

ti ''T

o3

CO

-o d >>

>,>,Ui>,>,

& >.

-d >> >. >.

>.o

>i bb

>> bb^

d

. M

g

3

^.^i

^^■z f ^

1 t

bb . •. bb3
1 bC 1 1

^

bb3

_3-?

H

o
O

O

U5

d-Q >,

>i >^ dbb bb

bb bb

>.!X >■, bb

bbo

>i bb

bb ci. bb

1

__;

<v

d

,

3_.

•

1

Q
o

■§

^^^

bb bC k! . •

1 1

^ bb bb^

3

f-3

^3-?

M

dS >-.

& >)3 bb bb

bb bb

>> d d bb

do

bb d

bb d bb

bb

d
1

1

^

r^

03

lO

32 ^

"?|^ '5«>>^

bb .

& • '-3

1 bC bC-T

3

3

m3-?

q;>

ffi

o

d-d >)

rO t>>X5 bC bC

a^

>> &. a bc

do

bb d

d dbb

^

bb

rH

1

bb

03

d

.3 =!^

-3

Ih*

0)

3

C3

bb bb ;^'?
1 1 ?*-> 1^

bb3

T bC ^^

3-?

3

iiirO bb

M
M

2-d >.

3 >^2 bb3

d-d

>. d 3-3

d>>

ti) d

dd-d

H

bb

CO

d

-3

3

o

O

•g

_.2f

^ f'fi

_;

^bb J3

bb^-

^

tH

lO

bb^

^-T

-!;-°

bb3

o

2-6 ti

3 >.3 >^2

d-d

bb-d 3-d

d >»

3 d

d-do

O M

3

bb

i^

S a

<»

1— »

O

0-J3

.3^

CO

f bb '?!' v^l

bb

33

f bD .X3

_;3

33

3-63

^ >,^^i-i

d-d

hC-3 ^-3

^ >-.

3-d

d-do

c

en

i2

CO '3

;3

33 o

o

CO

73

13 a SB

rr, 2

a

CO

-7! S O

s

s

iS »3 ■»

en

?^ M m

1=1 ^

.3.sJ|

u a 3

i2^

•^03 3

g'd;S&

.2§)J

So-o

cSs

S-Sg

o

-t^ 03 +^

bC >> 3

a >^ =* c >>^

J^ >--^

►3

m >- 3 "

rxi

J3^

o

h5 Pi

107

05

-a

CI
o3

c

'-D
o

CO

I— I °
M

c

O

r3
C
o3

tc
bC

3

CO

O

o3
>

O
S

CO

<u

o

3

c 1 .
o^-S in

O ^ '
O

-J o

o o ooooo ooo

ooooooooooooooooooooo
ooooooooooocooooooooo

+ +
+ + +++
++++++++

^^

>1 >-, Crb >! I

-1 CI, >-l >;^^

bC

t.1 ^H fc. .

s

U5

o
o

+
+ ++ + +++
++++++ +++
+++++++++■

+ +4- +

++++++++++

■ r I

hC

^^

bC

bC

'-" o o ^-

>> >^, i; >..Q >; bb >-. >^ J >. >^, >-. L." ^ ^ g.^ >. >; d

u

.»^

d

a
o

d

U

Ui

h

o

to

o

I— 1

U

+++ ++ +++++ +++++++
+++++++++++++++++++++
+++++++++++++++++++++

-. !; he
.-2 •

i'^ bC

bC

u
^

+ + + + + +++++

+ + + ++ +++++++
+++ I ++++++++ I +++++++

hC

bC

b£

,C >, bCJ2 >> C5- >i >>

hr

t-'

a

fl

■*s

o

O

J3

«

O

S

a)

o

O

+
+

+
+
+

bit
I

bC

.3 o

"» 2 !;

0)0.0

2o"

hi S

+

+ + + +

o
O

X! Cr^ bJD

So

2 >' bbS

^H bC bC"Ci X) >i >i

+

+ + + +
+ + + +
+ + + + +

t^ bJC bC-3 .Q

+ +
+ + + +

+ + + +

++++++++

bH

■^ >, C c nxiX^j^

'^ a

IM COCCI 00

I I ■ ■

I 1

F— I CD
■ GO "*

c
I

mo 00(M C5 "-I
(M t^ iC »^ Tf CO
■^ (M ■— I Cj ^ (N

o
oo

1

CO

CD

CO

I

0(M

C: O
-- (N

3

Q

o

•n

'S
o

O

0.3 3 -f^

■73

o

o

<D

rn t_ m M • '-^

L. (A

o -^ o o

O O fc< w

-<<<<:QQ(ilWKPwPMPH

^J . -1 . -H 3

5 P-^^ o

ii S ^ O ^ w

3.g aj o3 (U

Si-

■§1

PhCli

1

^ 1

^

._.

u.

^ — '

ca

d

0)

>^

«

o

o

_o

^

a

i_,

S3

03

H

fl O

O O
o3 03

03

s

3

'3

£S

o3 o3 O 3

5J

c

&.— .
!~ e

a

T3 N
o3 03

0:7:

«
>>

^ o

O M
. o

0

0

II

A

-M

ro

bC

bO

0

tc

bC

+

II

C4-t

0

b<

hr

0

Xi

-^j

f-l

rr

III

3

C

S

0

o-c

0

-M

^

II

0

.^

,

cS

it

0

■^

C

bC

0
0

c

H

■s

0

>

X!

+

II

>v

^

tH

-0

^

^

^

0

0;

J-

03

OJ

s.-^

CO

II

-t-'

,

r/i

a o3

-4—

0)

C

3

n

^

«

OJ

II

-t^

J2

0

r/1

^ ,

en

rn

0)

r!

e

0

0

.^^

r!

f^

>

7J

^

aj

XiA

-<H

108 MICROSCOPIC TECHNIQUES

Chemotherapeutics. Helander ( 1945a) has reported the detec-
tion of chemotherapeutics in paraffin sections prepared from frozen
dried tissue. The blue fluorescence of tissue itself, when illuminated
by ultraviolet, may obscure the fluorescence of compounds present
if they too have a blue fluorescence. This difficulty can be circum-
vented in some cases by heating the sections for a short period in
order to change the color of the fluorescent light emitted by the
drug. In a work of a high order of excellence Helander (1945b)
presented a table (Table I) of the fluorescences of various tissues,
and after intramuscular injection of drugs into mice, he found that
sections of muscle tissue showed the fluorescences indicated in
Table II.

Fischl and Singer (1935) made use of the fluorescences of atebrin
and trypaflavin to show that these compounds are taken up in vivo
by trypanosomes. A table of the fluorescences of compounds, and of
certain parasites after treatment with them, has been given in a
paper by Bock and Oesterlin ( 1939) .

Metals. The detection of uranium salts in incinerated sections
of tissue has been made possible by the fluorescence of these salts,
(page 145).

The colors of the insoluble 8-hydroxyquinoline derivatives of iron,
calcium, magnesium, aluminum, manganese, zinc, and copper have
been used for identifications of these metals in tissue sections (see
page 21). However, no one seems to have exploited the possibility
of utilizing the fluorescences of these derivatives. DeMent (1942)
lists the fluorescent colors of 27 metallic hydroxyquinolinates, and
it would appear that, in some cases at least, a more clear-cut
differentiation between the metals would be possible on this basis.

Spectroscopic Analysis of Fluorescence

The identiflcation of substances in tissues by means of the spec-
troscopic analysis of the fluorescent light they emit has been utilized
to some degree. Dhere ( 1939) presented a thorough and able review
of this technique and gave data relative to various carbohydrates,
lipids, proteins and amino acids, alkaloids, chlorophylls, bile pig-
ments, porphyrins, carotenoids, flavins, thiochrome, and a few other
special compounds. Either direct vision or grating microspectro-
scopes may be employed, and the spectral lines of mercury can be

EMISSION HISTOSPECTROSCOPY 109

used conveniently for calibration after the filters are removed, since
a mercury arc is the source of the radiation.

More recently Sjostrand (1946a,b) has employed this technique
for the localization of riboflavin and thiamine in cells. Sjostrand
used the freezing-drying treatment on this tissue (frog eyes, etc.)
followed by paraffin infiltration. The 3-5 [x sections were subjected
to ultraviolet illumination on the stage of a microscope and the
fluorescent light which resulted was passed from the ocular into a
spectroscope for analysis.

B. EMISSION HISTOSPECTROSCOPY

The technique for the identification of certain elements in selected
histologically defined portions of tissue by means of emission spectra
was developed independently and at about the same time by Gerlach
and Gerlach, and by Policard. Modifications were made later chiefly
by Scott and Williams. The emission spectra are obtained from the
radiation produced by consuming, in high-frequency spark, a chosen
cellular region of a tissue section or a selected bit of biological
material. The amount of tissue destroyed in sections is a disc of
about 1 mm. diameter and 0.1 mm. depth; this would have a weight
of approximately 0.08 mg. and hence elements giving good spectral
lines can be detected in this quantity of fresh or fixed tissue. Ele-
ments that have been identified by this technique include potassium,
sodium, calcium, magnesium, manganese, aluminum, iron, zinc, lead,
copper, mercury, silver, gold, carbon, silicon, phosphorus, and boron.
Attempts at quantitative work have been made, but the technique
is primarily qualitative at present.

There is little essential difference between the apparatus developed
by Gerlach and Gerlach (1933) and Policard (1931-1932, 1933a).
The former employed a high-frequency generator that was superior
to the one used by Policard, and the latter used a metal slide to hold
the tissue section instead of the glass slide used by the Gerlachs.

1. Policard Technique

A frozen section, 50-200 fi thick, cut preferably from fresh tissue
or else from alcohol or formalin-fixed material, is placed on a slide
of pure platinum or gold which is fastened to the microscope stage

no

MICROSCOPIC TECHNIQUES

(Fig. 5). A platinum or gold needle embedded in a cylinder of
insulating material, which is fixed in one of the objective holes of
the microscope's revolving nosepiece, serves as a movable electrode.
The circuit is completed through the frame of the microscope which
is grounded. The cellular region to be examined is chosen with an
objective in position, and then the objective is swung out and the
electrode needle brought into place. The needle should be in line with
the optical axis of the microscope so that it is centered over the

Fig. 5. Arrangement of Policard's
apparatus for emission histospectros-
copy: 1, Microscope stage under re-
volving nosepiece bearing tlie upper
electrode; 2, high-frequency appara-
tus— dermatological type; 3, Hilger
quartz spectrograph; 4, cylindrical
quartz condensing lens; 5, optical
bench; 6, ground lead; 7, current
source; 8, upper electrode lead. From
Policard (1931-1932)

selected spot. The distance between the point of the needle and the
section is fixed between 1-2 mm. When the circuit is closed, the
radiation from the spark produced is condensed by the quartz lens
and directed through a 0.1-0.2 mm. slit into the quartz spectro-
graph. For a strong spark, a period of not more than 1 sec. seems
to be sufficient for the exposure of the photographic plate on the
spectrograph, but for weak sparks 20-30 sec. may be required
according to Policard. The best spectral lines were observed in the
ultraviolet range, 100-300 m/x.

2. Scott and Williams Technique

Scott and Williams (1935) modified the apparatus chiefly by
screening the electrodes, placed 1.4 cm. apart, in order to practically
eliminate electrode lines from the spectra and to obviate the need
of cleaning the electrodes often. Since these workers did not employ
needle electrodes mounted on a microscope, they could not make
as fine a histological selection of the sample as the European investi-
gators. Scott and Williams (1934) minimized this point and claimed
that removal by fine dissection of the bit of material to be studied
is sufficient. In most instances this would be true, but there are

EMISSION HISTOSPECTROSCOPY

111

conditions conceivable in which the needle electrode assembly
might have advantages. In any case the Scott and Williams appara-
tus has a nmnber of significant improvements, as will be apparent
from the following description of it.

The spark generator construction is shown diagrammatically in
Figure 6. Oscillations are produced in the primary circuit by a
spark passing between two thick brass discs rotated by a small
motor to prevent erosion and local heating. The space between the
discs is supplied with photoelectrons by focusing ultraviolet radia-
tion on it from a quartz mercury lamp. The analyzer spark in the
secondary circuit has an improved continuity and steadiness as a

o "

—I i£>

g >

, o
. o

: o
:7

O rsi OJ

o > j: -o

g ™ m e

o fu (/I y^
1^ ^

Quartz

mercury arc

Fig. 6. Wiring diagram of spark
generator assembly. The ionizing beam
of ultraviolet radiation is shown
focused between the rotating discs of
the spark gap. From Scott and Wil-
liams (1935)

Fig. 7. View of box enclosing ana-
lyzer spark gap and electrode screen.
Portion of spectrograph seen at right.
From Scott and Williams (1935)

result. A large open type oscillation transformer is employed having
two heavy primary, and twenty smaller secondary, turns wound
on a wooden frame as close to the edge of a 36 in. square as possible.
A clearance of 0.5 in. allowed between the secondary turns. The
condenser is composed of a stack of glass plates (8 X 8 X % in.)
separating eight sheets of aluminum foil (6 X 6 in.) connected alter-
nately. The current passing over the analyzer spark gap is about
0.45 ampere.

The electrodes of the analyzer spark gap are steel balls (0.25 in.
diameter) placed 1.4 cm. apart which are supported through the
bakelite back of a metal box housing fitted with quartz and glass

112

MICROSCOPIC TECHNIQUES

windows, removable for cleaning. An aspirator tube is fitted into
the top of the box to remove vapors and the box itself is placed on
alignment pegs to maintain fixed optical relations. The disposition
of the spark gap relative to the spectrograph is shown in Figure 8.
As mentioned previously, the horizontal slit in the side of the box
facing the spectrograph serves to screen out electrode lines from
the tissue spectra and to obviate the need of cleaning the electrodes
oftener than once a month. The end of a 2 in. length of Pyrex tubing
(2-3 mm. inside diameter) or rod (2-3 mm. diameter) is centered
in the spark gap. It has been shown that the glass does not affect
the spectra. Bits of cellular material (3-6 [A.) are placed on the
end of the glass so that they will be in the center of the spark. A

To spark generator

Small Pyrex tube carrying bit of tissue
l"

1.4 cm. between faces

Spectrograph collimator

14.0 cm. *^

\*.>\< — 5.3 cm. — »4<

1.3 cm.-^

Fig. 8. Diagram of disposition of the spark gap

relative to the spectrograph.

From Williams and Scott (1935)

film of purified Eastman gelatin may be helpful in effecting the
adherence of dry material. A strip of tissue, 5 mm. or more long,
may be placed in the glass tube and fed into the spark with a push
rod; and in a similar manner liquid taken up on a small strip of
ashless filter pulp can be subjected to test. In the latter case, con-
trol spectra for the pulp alone must be obtained.

A Gaertner L250W quartz prism spectrograph taking 3i/4 X 4l^
in. photographic plates was used. The spectrograph slit was fixed at
0.05 mm., and Eastman "50" plates were employed for high sensi-
tivity and Eastman Process plates for high contrast. Thirty ex-
posures may be made on each plate. Two microscopes with a com-
parison ocular were utilized for comparing the positions and inten-
sities of lines on different plates.

Faint iron lines from the electrodes are apparent in the spectra

VISIBLE AND U.V. ABSORPTION HISTOSPECTROSCOPY 113

SO that, if iron is to be investigated, another electrode metal should
be used. For an adequate exposure (15-30 sec), 2-4 /xl. of tissue
are usually sufficient. When tissue is subjected to fixation, a control
experiment is necessary to test the fixing fluid spectrographically;
the filter pulp method may be employed for this purpose.

The Williams and Scott ( 1935) photoelectric apparatus for dark-
field photometry and densitometry has been used to determine the
intensity of the spectral lines in order to obtain a more quantitative
estimation of the elements. A description of this apparatus is given in
the section dealing with microincineration (page 146) . The blackness
of the photographed spectral lines is measured by placing the photo-
graphic plate on the mechanical stage, adjusting the reflecting prism
so that the light emerging from the ocular is reflected directly down-
ward on the slit of the photocell box, and observing the galvanometer
deflection after the proper focusing has been made. The deflection for
an unexposed portion of the photographic plate is then taken.

C. VISIBLE AND ULTRAVIOLET ABSORPTION
HISTOSPECTROSCOPY*

The application of the quartz microscope to measurements of the
absorption spectra of cellular components in situ, particularly as
developed and applied by Caspersson and co-workers at Karolinska
Institutet, Stockholm, offers a new and promising approach to the
solution of many histo- and cytochemical problems. The ingenious
apparatus of Caspersson (1940), subsequently modified by Gersh
and Baker (1943), has already yielded valuable information con-
cerning the nucleic acids of chromosomes (Mirsky, 1943), the nature
of thyroid colloid (Gersh and Baker, 1943), and chemical character-
istics of the Nissl bodies in nerve material (Gersh and Bodian, 1943-
a,b). (It should be pointed out that the absorption method cannot
differentiate between ribonucleic acids and desoxyribonucleic acids
since their absorption spectra are almost identical, having maxima
at about 260 m/i. However, the differentiation can be made qualita-
tively by staining reactions, (pages 65, 66).

The great advantage of studying cell structures m situ by this
technique is made particularly impressive by the fact that the
spectral measurements can be carried out on quantities of material

*See Bibliography Appendix, Ref. 31.

114 MICROSCOPIC TECHNIQUES

down to 10"^ /tg. Investigations may be made on selected structures
in microtome sections or on mechanically separated cellular com-
ponents. When sections are to be employed, they are best prepared
from tissue embedded in paraffin or celloidin after freezing-drying
treatment. The technical requirements in absorption spectra meas-
urements by means of the ultraviolet microscope have been examined
critically by Cole and Brackett (1940). Laviri' (1943) simplified
the focusing of the ultraviolet microscope by using a willemite
screen which produces a visible image with ultraviolet illumination.

The absorption technique applied in situ has certain drawbacks
that should be considered, elegant though the technique is. Thus,
Danielli ( 1946a) has sounded a warning that hazards exist in as-
cribing to particular substances the absorptions found in different
parts of a cell. Effects of molecular interactions and interferences
by other substances are possibilities that are not to be ignored. Hence
the method will be of greatest value when the results are interpreted
with appropriate regard to these limitations.

A most ingenious microscope arrangement for the colorimetry of
0.5-1.0 ix\. drops of liquid was developed by Norberg (1942), also
at Karolinska Institutet, and applied by him to the measurement of
phosphorus in quantities down to 0.5 n\[xg. This technique requires
the removal of the specimen from the rest of the tissue and its
chemical treatment to yield a solution which can be subjected to
absorption analysis.

Stowell (1942) designed an apparatus for the measurement of the
amounts of stain or pigment in tissue sections. For the measurement
of stained constituents the quantitative significance of the method
depends on the degree of correlation between the amounts of the
stain and the substance for which the stain is specific, a correlation
often poorly defined. Stowell applied his technique to the estimation
of desoxyribonucleic acid by means of the Feulgen stain.

1. Casper sson in Situ Technique

A diagrammatic representation of the apparatus is given in Figure
9. The source of radiation may be either the Philips water-cooled
super-high-pressure mercury lamp (A), a tungsten lamp (B) sup-
plied with current from storage batteries, or Kohler's rotating elec-
trode spark gap (P) . The mercury lamp may be used for radiation in
the visible portion of the spectrum and in the ultraviolet range down

VISIBLE AND U.V. ABSORPTION HISTOSPECTROSCOPY

115

to 230 mix. However, since variations both in voltage and water pres-
sure affect the mercury lamp, the tungsten is employed for the
longerwave ultraviolet range. The spark source is used for wave-
lengths shorter than 235 m/i,.

The radiation, after passing through a monochromator (C) is
concentrated on the object (/) on a quartz slide by the condenser
(//). (The second monochromator slit is at D, lens at E, and a 90°
ciuartz prism at F.) For ultraviolet work a fused quartz condenser
is used, and in the visible range a good achromatic type is employed.
Zeiss objectives designed for the longer ultraviolet (down to about
340 mju) or the fused quartz objectives of Kohler and von Rohr for
the shorter wavelengths are used (K) . For routine work down to

V^

i^ o

Fig. 9. Arrangement of Caspersson's apparatus

for photoelectric absorption histospectroscopy.

From Caspersson (1940)

240 m/A, Caspersson ( 1940) found it convenient to have a glycerin
immersion lens corrected for 257 m/x with an aperture of 1.25, one
corrected for 275 m^a, and a long-wave ultraviolet recorrected
apochromat. Oculars with iris diaphragms (L) were employed. The
lenses in them were of quartz for the ultraviolet work.

A 90° quartz prism (M), adjustable by means of micrometer
screws, is used to deflect the radiation through the opening of an
electrically driven rotating sector (A^) on to a photoelectric cell
(R). The prism can be replaced either by a Kohler focuser (Y) or

116 MICROSCOPIC TECHNIQUES

a photomicrographic camera (X). Every object measured must be
photographed in order to estabhsh its exact position and dimensions.
The photocell is connected to a string electrometer and both of these
instruments are well shielded and also protected from moisture by-
means of phosphorus pentoxide. Various photocells are used for
different wavelengths; gas cells are usually employed. For the
shortest ultraviolet, cadmium; for medium ultraviolet (260-350 m/x),
sodium; for long ultraviolet and visible (350-550 nifi) , potassium;
and for wavelengths over 550 m/x, potassium-cesium cells are used.
The telescope (0) is placed in front of the photocell to control the
optical centering of the system ; this centering must be very exact.

It is necessary to compensate for variations in the source of
intensity of the radiation, and for this purpose a quartz plate (G)
is interposed in the optical path in order to reflect a small percentage
of the radiation on a photocell ( V) . Readings of the changes in the
photocell current can be used to correct the readings of the electrom-
eter (S) . (T and U are leak resistance and four-step potentiometer,
respectively.)

Measurements are made by taking the deflection of the electrome-
ter with the object in position and in focus, and then moving the
object away so that a clear space on the slide lies in the optical
axis. The opening in the rotating sector is reduced until the amount
of radiation striking the photocell is the same as before, i.e., the
same electrometer deflection is produced. The absorption in the
object will then be equal to the decrease effected by the sector, and
extinction coefficients may be calculated.

Gersh and Baker Modification. A somewhat simplified set-up,
with American-made instruments, is employed by these investiga-
tors, as may be seen in the diagram of their apparatus (Fig. 10) .

The source of radiation is a Daniels and Heidt (1932) type of
medium pressure mercury arc in a quartz capillary tube which is
mounted about 1 cm. from a quartz window in a large copper box.
The lamp is water cooled, and since the rate of cooling affects the
radiation output, the water line is equipped with a pressure regulator.
The lamp consumes 500-700 watts from a 220 volt D.C. line; a
ballast resistance is placed in series with the lamp.

The entrance opening of the monochromator is a circular hole of
about 0.8 mm. diameter in a thin sheet of copper fixed 0.5-1.0 mm.
in front of the arc, in the water bath. The two equilateral quartz

VISIBLE AND U.V. ABSORPTION HISTOSPECTROSCOPY

117

prisms of the monochromator are 5 cm. long and 4 cm. high; the
table on which they are mounted can be rotated by means of a
slow-motion screw. The collimating lens has a focal length of about
8 cm. and the telescope lens about 80 cm.; both are held in adjustable
brass mountings and these with the prism table are fastened to an
iron plate on leveling screws. The whole apparatus is enclosed in a
wooden box with the required apertures. A spectrum from the first

Photoelectric cell

Amplifier Lii
Galvanometer ( o

Ultraviolet
light source

^

Quariz
monochromator

CENTRAL

BEAM

k

tf 37

Quartz obiective

Quartz slide
and cover slip

Quartz condenser

^>^ — - — y^

^Quartz ^

reflector

Fig. 10. General arrangement of light source,
monochromator, microscope, and measuring device
for determining the absorption of ultraviolet Hght
by minute volumes of tissue. From Gersh and Baker
(1943)

prism, without entering the second, passes out of one of these
apertures to fall on a calibrated wall scale 2 meters distant. By this
means the prisms can yield any chosen wavelength when manually
adjusted.

A Zeiss quartz microscope of Kohler design is mounted on a
leveling table so that it can receive the radiation reflected by a
quartz right-angle prism. In order to fill the objective field, the
substage condenser is focused on the field of the telescope lens rather

118 MICROSCOPIC TECHNIQUES

than on the image of the entrance opening of the monochromator.
The diameter of the condenser diaphragm is set at 6 mm. to insure
sufficient spectral purity.

Both photoelectric and photographic recording may be employed.
The quartz photoelectric cell is a No. FJ-405 of General Electric Co.,
and it is mounted 56.5 cm. above the ocular in a large brass cylinder.
This cylinder has a quartz window close to which is a frame that
holds a series of circular slits. The cylinder is horizontally adjustable
so that a slit can be brought into the optical axis of the microscope.
The brass cylinder also contains a type D96475 electrometer tube of
Western Electric Co., and a 10^" ohm SS white grid resistor. The
output from the photocell is connected to the grid of the tube which
is included in a Penick amplifier circuit maintained on three storage
batteries of large capacity. The amplified current is measured by a
Leeds and Northrup type R galvanometer with a scale 1.5 m. from
the galvanometer mirror.

In order to bring the photocell into adjustment in the optical
axis of the microscope, the shadow of an ocular cross hair is pro-
jected by means of "white" light on the photocell aperture, which
is then adjusted until its center and the center of the image conin-
cide. The cross hair in a fluorescent finder placed above the ocular
is adjusted similarly with "white" light and ultraviolet radiation.

For the measurement of absorption curves of larger uniform
objects, the object is centered in the cross hair of the finder, the con-
denser is focused on the plane of the telescope lens, the objective is
adjusted to give a sharp image, and the current generated in the
photocell is measured. Then the object is moved away so that only
the clear slide is in the optical path and another measurement is
made. From these data the percentage transmission and the extinc-
tion coefficient can be calculated. The measurements are then re-
peated at each wavelength chosen. For studies of thyroid colloid,
Gersh and Baker (1943) used a 6 mm. objective, lOX ocular, and a
photocell aperture of 11.9 mm. With these optics the light transmis-
sion was measured through a tissue area of 143 /x- and a volume of
2861 /i,^. For measurements on cytoplasm and nucleoli the respective
systems were 6 and 1.7 mm. objectives, 14 and lOX oculars, and
8.73 mm. photocell aperture in both cases.

When the measurements are to be made on smaller and less
homogeneous objects such as Nissl bodies, a different and more

VISIBLE AND U.V. ABSORPTION HISTOSPECTROSCOPY

119

accurate technique is used. A quartz plate 1 mm. thick and 2 cm.
square is placed in the path of the monochromatic beam at an angle
of 45° and at a distance from the telescope lens of 46 cm. (Fig. 10 L
By this means 6-7 % of the incident radiation is reflected to another
photocell mounted at a distance from the quartz plate equal to that
of the aperture of the microscope condenser. The photocell output is
amplified by a Huntoon (1935) direct-current amplifier and passed
through a Leeds and Northrup type P wall galvanometer. Simul-
taneous readings are taken on the same scale, from both this gal-
vanometer and the one used for recording transmission, at each
wavelength without moving the object, but with careful focusing of
condenser and objective at each wavelength. After these readings
are obtained the object is moved away and the readings for the
clear quartz slide are taken. The data are used to calculate the
transmission and extinction coefficients as usual.

l.U

_ 1

' ' L

0.5

I A

/

0.4

-

=j^

0.3

1

/

-

0.2

^/'^^'^

/
/

/

-

0.1

X 2^

— •

- •

/ 1 0
^^ 0.5

0.1

^ B

-^^'

-

U.Ub
0.04
0.03

nn?

-;^Z^'

-'' 2

1

400

300

! 1

1 !

in

ro

o CM

n CO

rt o

ro m

'J- ^

(T> o

00 00

C\J CM

cvj i^ m o

ID on CO o

^ Ln ^ ^

CM CM CM CM

WAVE LENGTH, A

Fig. 11. Absorption curve of colloid in a single
follicle of a thyroid gland in alkaline (^1) and acid
(.42) medium, as compared with the ultraviolet ab-
sorption curves of extracted sheep thyroglobulin
made by Ginsel in alkaline (fil) and acid medium
(B2). From Gersh and Baker (1943)

The reliability of the technique is shown by Figure 11, taken
from Gersh and Baker (1943). The curves in the inset (B) were
obtained by Ginsel (1939) for extracted sheep thyroglobulin in both
acid and alkaline media, while those in (A) were established by
Gersh and Baker by their histospectrographic technique on the
colloid in a single thyroid follicle.

120

MICROSCOPIC TECHNIQUES

2. Norberg Technique

Apparatus

A diagram of the optical system is given in Figure 12. For work in
the visible range, the light source employed is a 100 watt tungsten
band lamp (A) supplied with current from a large capacity (150-
200 amp. hr.) storage battery. Monochromatic light is obtained from
a Winkel-Zeiss monochromator (B) . (C is second monochromator
slit.) A filter (F) may be used between the condenser (E) of the
microscope and the sample slide (G) . The microscope objective is
indicated by H and the ocular by 0. During measurement the ocular
is removed and the light is reflected by the prism (/) , which is mov-

Af L .

/

h!

H

,.G

^7

KyE

Fig. 12. Microphotometer.
From Norberg (1942)

Fig. 13. Photocell amplifying circuit.
From Norberg (1942)

able about both a horizontal and vertical axis, to the photocell (L).
Potassium cells are employed for wavelengths 450-550 m^u, and
potassium-cesium cells for longer wavelength. The current generated
in the photocells is amplified by circuits in M and then conducted to
the galvanometer (A^). Details of the amplifying circuit will be con-
sidered subsequently.

For measurements in the ultraviolet region a high-pressure mer-
cury lamp (P) with movable prism (S) and 90° prism T, of the
Philips Philora H P 300 type, is used with a spectral filter {R) .
Variations in the intensity of the radiation from the mercury lamp

VISIBLE AND U.V. ABSORPTION HISTOSPECTROSCOPY 121

are coaipensated by casting a portion of the radiation on photocell
Lc by means of the semireflecting glass (D). The current generated
in this photocell is amplified and made to oppose that from photo-
rell L.

The galvanometer is employed as a null-point instrument and the
amount of radiation striking the photocell L is controlled by the
rotating sector (K) . The rotating sector with its motor is mounted on
a slide, adjustable by a rack and pinion arrangement. The radiation
passing the sector can be controlled over a greater range (0-50%)
by the use of two discs, each with a 90° segment removed, which are
made to rotate in opposite directions. Kortiim (1934) has described
a simple sector wdiich operates on a similar principle. Norberg finally
employed a sector patterned after the Askania-Werke (Berlin)
model, which enables adjustment and reading while the sector is in
action, and the accuracy obtained in the absorption is 0.025%.

The amplifying circuit for the photocell current is shown in Figure
13. It is a modification of that described by Custers (1933) and it
utilizes two Philips 4060 electrometer tubes. The apparatus can be
used with either the single photocell (L, Fig. 12), employing com-
pensation with the potentiometer, or with both photocells (L and
Lc) . The galvanometer used by Norberg was a Zernike C (Kipp and
Zoonen) instrument having a tension-sensitivity of 10,000 scale divi-
sions per volt and a stability level of 3 X lO^^-'' amp. By means of the
Ayrton shunt ( V) (Fig. 13) the sensitivity can be reduced by 0.1 and
0.01. The galvanometer is shown at iV; the potentiometer for com-
pensation where only one photocell is used at U; Re resistance
= 1.2 X 10^^ ohms; R© leakage resistance = 1.1 X 10^*^ ohms.

Other resistances are marked in ohms. The apparatus must be
mounted within a Faraday cage to avoid electrostatic disturbances
and unsoldered contacts must have large frictional contact surfaces.

Manipulations

The filament current in the amplifier is turned on 30 min. before
measurements are made, and the tungsten band lamp is also turned
on long enough in advance to attain steady illumination. The sample
slide on the microscope stage is adjusted so that the image of the
exit slit of the monchromator ajipears in the middle of the sample

122 MICROSCOPIC TECHNIQUES

drop. The size of the image is 0.12 X 0.12 mm. The slit image is pro-
jected over the opening of the measm-ing photocell (L, Fig. 12) by
the objective and the prism (7). The sector is started; the shutter in
front of the photocell is opened; and the photocurrent is compen-
sated by a potential applied by means of the potentiometer ( U, Fig.
13) to the grid connected to the other photocell so that the zero read-
ing on the galvanometer may be obtained. The galvanometer is
usually constant within 0.5 mm. after 1-2 min. The sample slide is
now shifted so that the light will pass through solvent alone or a
suitable blank. The sector is adjusted until the galvanometer zero
reading is again obtained. It is well to repeat the measurements
several times.

When the mercury lamp is used as the source of radiation, the
photocurrent from the measuring photocell is compensated by the
photocurrent from the other cell which is illuminated by the semi-
reflecting glass as previously mentioned. Thus, with the mercury
lamp the compensation current from the potentiometer is replaced by
the compensating photocurrent.

The Sample Slide for Absorption Measurements

Very clean microscope slides and cover glasses are coated with
hydrophobic films of nitrocellulose in order to prevent the aqueous
drops from spreading. This is accomplished by pouring over the glass
a solution of 1 g. of highly nitrated cellulose (about 13% nitrogen),
0.1 g. diethyl phthalate, and 0.01 g. butyl stearate in 100 ml. butyl
acetate. The glasses are set aside in a tilted position to drain and
dry for at least 3 days in a dust-free place. A drying drum with an
electric fan may be used to reduce the drying time. If plastic plates
are substituted for the glass, no film is required, but care must be
taken that the plastic is optically homogeneous. Two narrow strips
of polished glass having a thickness of about 0.35 mm. ( from a hemo-
cytometer cover glass) are placed on the hydrophobic film parallel
to one another. Between them, sample and blank drops of the order
of 0.5-1.0 ju,l. are pipetted, and paraffin oil is added to fill the area
between the glass strips. A cover glass is set on the glass strips to
complete the cuvette. To determine the layer thickness in the
cuvette:

VISIBLE AND U.V. ABSORPTION HISTOSPECTROSCOPY

123

1. Draw two thin parallel lines 1 cm. apart on the upper surface
of the slide with India ink.

2. In a similar manner, draw two lines 3-5 mm. apart on the
under surface of the cover glass.

3. Place the cover glass so that the lines on the two glass surfaces
intersect.

4. Measure the distance between the upper and lower lines at the
four points of intersection by focusing on the lower line with the
microscope and then using the micrometer screw to focus on the
upper line.

5. Multiply the distance obtained with the micrometer screw by
the refractive index of the paraflBn oil to get the thickness of the
layer.

Accessories for Norberg Technique

Quartz or Supremax Glass (Schott, Jena) Needles. For the

isolation and incineration of the sample, fine needles with slightly
thickened points are used. The needles are cleaned by boiling in 2 A''
nitric acid and rinsing with distilled water.

Fig. 14. Apparatus
for microhydrolysis : A,
Steam mantle ; B, cham-
ber of hydrolysis; C,
holder with quartz
needles; D, water.
From Norberg (194£)

Hydrolysis Chamber. The arrangement shown in Figure 14 is
used for the hydrolysis of certain constituents in the ash after the
sample has been incinerated on the tip of a quartz needle. For the
hydrolysis of pyro- and metaphosphate to orthophosphate, Norberg
placed 1-2 /xl. oi 1 N hydrochloric acid on the tip of each needle and
heated for 1 hr. at 100° in the chamber. Should the acid evaporate
in the chamber, the hydrolysis must be repeated. After hydrolysis
the drops are allowed to evaporate to dryness at room temperature.

Muffle Furnace. An ordinary muffle furnace may be used for
the ashing of the sample on the tip of a quartz needle.

124 MICROSCOPIC TECHNIQUES

Methods
Phosphorus

By means of his microscopic photometric technique (page 120)
Norberg ( 1942 ) developed a method for the estimation of phos-
phorus in quantities down to 0.5 nijug. with an error not greater than
about 20% for single analyses. Naturally the error is less with larger
samples and greater accuracy is obtained by averaging the results
of several determinations. For amounts of phosphorus under 1 m/^g.,
Deniges' stannous chloride method is recommended, while for 1 m/^g.
and more it is preferable to employ Fiske and Subbarow's amino-
naphtholsulfonic acid method, which is technically easier.

Norberg Method for Phosphorus

SPECIAL REAGENTS

0.005 N Calcium Acetate.

1 N Hydrochloric Acid.

Deniges Reagents, (a) 0.01 M sodium molybdate in 0.6 .V sulfuric
acid; (b) 0.2 N stannous chloride in concentrated hydrochloric
acid, approximately 2.5% SnC^-HoO.

Fiske and Subbaroiv Reagents, (a) 0.022 M sodium or ammonium
molybdate in 0.75 N sulfuric acid; (6) dissolve 12 g. sodium meta-
bisulfite in 80 ml. water, stir in 0.2 g. 1,2,4-aminonaphtholsulfonic
acid (some commercial preparations of this compound are not
suitable, that of British Drug Houses Ltd. proved to be good) and
add 2 ml. 20% crystallized sodium sulfite. Let stand overnight
and filter off the undissolved aminonaptholsulfonic acid. Store in
a dark bottle.

PROCEDURE

1. Obtain the sample on the tip of a quartz needle (page 123). If
the sample is liquid, pipette 1 /il. 0.005 N calcium acetate onto the
needle tip with the sample and allow to dry in the air. If the sample
is solid, the calcium acetate is placed on the tip and allowed to dry
before taking on the sample. The excess calcium is required to pre-
vent loss of phosphorus during the incineration.

VISIBLE AND U.V. ABSORPTION HISTOSPECTROSCOPY 125

2. Place the needle in a cold muffle furnace and turn on the heat.
\Mien the temperature reaches 500° turn off the heat and remove the
needle when the furnace is cool. As an alternative, place the needle in
the furnace at 500° and remove it after 20-30 min. at this temper-
ature.

3. Hydrolyze the pyro- and metaphosphate to orthophosphate
by pipetting 1-2 fA. 1 X hydrochloric acid onto the needle tip and
heating for 1 hr. at 100° in the hydrolysis chamber (page 123) . Allow
to dry at room temperature.

4. For the Deniges method, add 0.05 ml. of the stannous chloride
soln. to 10 ml. of the sodium molybdate soln. This mixture nmst be
used within 3 min. after its preparation. Pipette a 0.6 to 1.0 fxl. drop
of the reagent mixture on a prepared slide near one of the inked
hnes (page 123) as a blank. Then pipette a suitable vol. (0.6-1.0 ^,1.)
of the reagent mixture onto the needle tip bearing the ashed sample.
Use the end of the pipette to mix the drop on the needle tip for 10-15
sec. in order to dissolve the sample and obtain a homogeneous soln.
During the next 15 sec, transfer the drop from the needle to the
slide, placing it beside the other iliked line a few mm. from the blank
drop. Surround the drops with paraffin oil and place cover glass as
described on page 122. The entire process from the addition of the
reagent to placing the cover glass can be performed in 35-45 sec.
Standardize the manipulations to maintain a constant evaporation
effect. Carry out the photometry after 5 min. from the beginning of
the color development, and finish within 45 min.

5. For the Fiske and Subbarow method, mix 24 ml. of the molyb-
date soln. with 1 ml. of the aminonaphtholsulfonic acid soln. This
mixture may be used for at least 1 hr. after its preparation, and it
is applied in the same manner as the Deniges reagent. Let the drop
stand for 30 min. before starting the photometry, and finish the
measurement within 2 hr. from the beginning of the color develop-
ment.

3. Stowell Technique

The apparatus consists of a lamp, a microscope, a photocell, and
amplification and recording equipment. A 50 c.p. automobile head-
lamp operated by a .storage battery is used as the source of light.

126 MICROSCOPIC TECHNIQUES

The lamp is housed in a Spencer (No. 367) lamp case which has a
filter holder. A microscope fitted with a mechanical stage, a 44 X
achromatic objective, and a 15X compensating ocular are employed.
The field of observation is limited to an area 50 X 35 /a by inserting
a rectangular diaphragm in the ocular. The light from the microscope
is thrown on a vacuum photocell (RCA 929) enclosed in a light-tight
box having a side tube to extend over the microscope tube. This
side tube contains a movable mirror so mounted that it can be
interposed in the light path to enable inspection of the field, and
then tm'ned out of the path to permit photoelectric measurement.
The photocell current is amplified by a General Electric FP54 tube
in a Barth circuit with a 10^^ ohm grid resistor (Penick, 1935). The
amplified current is measured with a Leeds and Northrup student
type potentiometer and a Leeds and Northrup type R galvanometer.

Measurements are made of the light transmittance through both
stained and unstained sections in order to obtain the absorption
due to the stain itself. In both cases, measurements are also made
of the transmittance through blank portions of the glass slides
adjacent to the sections to correct' for variations in the intensity
of the light source, changes in amplification or potentiometer bat-
teries, and alterations in thickness of cover glasses, slides, or mount-
ing media. When it is possible, fifty adjacent areas on each section
are measured and the mean percent absorption calculated.

Stowell and Albers ( 1943) employed a Coleman Model 10 S double
monochromator spectrophotometer by means of which they meas-
ured absorptions of light bands, having a 5 rap. spectral width, by
stained sections of tissue. Since no microscope is employed in this
apparatus, the absorption of the section as a whole, rather than a
chosen cellular region, is measured.

The photometric procedure was employed by Stowell ( 1942) for
the estimation of desoxyribonucleic acid in tissue sections by means
of the Feulgen stain (page 65). Light from the source was passed
through a heat-absorbing filter (Corning Aklo No. 396) and a green
gelatin filter (Wratten No. 58) before entering the microscope. The
extension of the method to absorption studies with a variety of
stains commonly used in histological examination was included in
the reports of Stowell and Albers (1943) and Stowell (1945b).
Subsequently Stowell (1945a) subjected the Feulgen reaction to
detailed study and described each step in his method of using it.

ROENTGEN ABSORPTION HISTOSPECTROSCOPY 127

D. ROENTGEN ABSORPTION
HISTOSPECTROSCOPY*

One of the most significant advances in histo- and cytochemical
technique has come from the work of Engstrom ( 1946) at Karohnska
Institutet, Stockhohn, who, by employing the roentgen absorption
of tissue sections or of very small volumes of liquid, developed a
procedure whereby quantitative elementary analyses can be directly
]ierformed with an accuracy of about 5-10% on 1 X 10"^ to 1 X
10"^- gram of material, i.e., quantities of the order found in single
mammalian cells. Thus, in specific instances, phosphorus and calcium
can be determined in a 10 /x section of bony tissue within an area
around 10 X 10 ix, and nitrogen and oxygen in a section a couple
of microns thick within an area of 50-100 /x-. A particular advantage
of this technique is that the tissue is not used up, and hence may be
subsequently employed for histological study by the usual methods
so that direct correlation may be made between the chemical
composition and the microanatomical structure. Furthermore, the
analysis is independent of the chemical structure in which the ele-
ment may be bound, and the physical state of the specimen is
unimportant, e.g., fixed tissue, dry powder, and in certain cases,
paraffin-embedded tissue, or solutions, may be used. A number of
elements can be determined on the same sample.

The advantages of being able to determine the total quantity of
a tissue element in situ without regard to its chemical form and state
of valence, or affiliations with other elements, are hardly to be
minimized. The method is confined to the quantitative determination
of elements having an atomic number of 6 (carbon) or greater. This
would include all elements of biological importance with the
exception of hydrogen.

Engstrom (1946) has pointed out that roentgen spectroscopic
methods based on emission analysis are not well suited for elements
with atomic numbers less than 20, and the emission methods cannot
be applied very well to the small surfaces involved in histo- and
cytochemical studies. This applies to the earlier procedure of von
Hamos and Engstrom ( 1944) in which tissue is subjected to roentgen
radiation and the secondary radiation is measured for the quantita-
tion of constituent elements.

* See Bibliography Appendix, Refs. 23, 24, 25, 26, and 27.

128

MICROSCOPIC TECHNIQUES

The use of the roentgen absorption method requires an under-
standing of the theoretical basis of roentgen spectroscopy. All that
can be included here are a few of the more salient features of the
theoretical treatment which Engstrom ( 1946, pages 19-50) applied
to his method, a description of the apparatus, and consideration of
certain other practical aspects. This information can serve merely to
acquaint the reader with the method, the actual use of which will
depend on the understanding of roentgen spectroscopy mentioned
l)reviously and a detailed study of the presentation of Engstrom
(1946).

O

1

I

o
o

z
o

o

h

y

<

CO
CO

<

/

1

^,<

abs

WAVE LENGTH

Fig. 15. Schematic representation of an absorption jump.
From Engstrom (1946)

1. Some Theoretical Aspects

Quantitative analysis based on the roentgen absorption utilizes
the absorption discontinuities which appear at a characteristic
wavelength for every element. The absorption of an element near
the K-absorption edge is indicated in Figure 15. The determination
of the element depends on the measurement of the absorption of
monochromatic radiation wnth wavelengths on each side of, and
close to, the absorption edge of the element to be determined.

The mass (x), in g./cm.^, of the element to be analyzed is given by
the following equation:

In

X =

A

Ml
P

(S)'

In

M2
P

©'

ROENTGEN ABSORPTION HISTOSPECTROSCOPY

129

TABLE III

Wavelength of the Absorption Edges for Certain Elements and Suitable
Analysis Lines (Engstrom, 1946)

Atomic
num-
ber

K-

absorption

edge,

X.U.

Xi

Xj

• Ele-
ment

Line

Wave-
length,
X.U.

Line

Wave-
length,
X.U.

C

X

0

Xa

Mg

P

S

CI

K

Ca

Fe

Cu

Zn

As

Br

Ag

I

6
7
8
11
12
15
16
17
19
20
26
29
30
33
35
47
53

43700

31000

23300

~11500

9496

5775

5009

4384

3431

3063

1740

1377

1280

1043

918

484

373

20 Ca Ka
22 Ti La
24 Cr La
32 Ge Lai
34 Se Lai
41 Xb Lai
44 Ru Lai
46 Pd Lai

20 Ca Kai

21 Sc Kai
70 Yb Lai
77 Ir La,
79 Au La,
81 Tl La,

92 U La,
51 Sb Kai
57 La Kai

36270

27370

21530

10415

8972

5712

4836

4359

3352

3025

1668

1348

1274

1013

909

469

370

6C Ka

21 Sc La
23 V La
11 XaKai
33 As Lai
40 Zr Lai

16 S Kai
45 Rh La,
51 Sb La,
20 Ca Ka,
68 Er La,
76 Os La,

78 Pt La,

79 Au L/3,
37 Rb Ka,
50 Sn Ka,
56 Ba Ka,

44540

31370

24310

11885

9652

6057

5361

4588

3432

3352

1780

1389

1310

1081

924

490

384

Atomic
number

Liii

absorption

edge,

X.U.

X,

X2

Ele-
ment

Line

Wave-
length,
X.U.

Line

Wave-
length,
X.U.

Ca
Cu
Ag

I

Hg
Bi

20
29
47
53
80
83

35630

13150

3691

2714

1008

922

21 Sc La
30 Zn La
50 Sn Lai
57 La La,
34 Se K^i
92 U La,

31370

12250

3592

2660

990

909

20 Ca La
29 Cu La

19 K Ka,

22 Ti Ka,
35 Br Ka,
37 Rb Kai

36270

13600

3734

2743

1038

924

where (mi/p) and {m/p) represent the mass absorption coefficients
for the element at wavelengths Xi and X2, respectively, p, the specific
gravit}^ of the absorbing substance, and /x, the linear absorption coef-
ficient ; ii and h are the intensities of transmitted radiation of wave-
lengths Xi and X2, respectively, and /i and I2, the corresponding in-
tensities of the incident radiation, P is a function of atomic number
and wavelength. When the wavelengths are selected close to the
absorption edge, and the intensities of the incident radiation of both

130

MICROSCOPIC TECHNIQUES

wavelengths are equal, i.e., /i = h, the equation may be simplified
as follows :

ii = 42e-(Mi/p-M2/p)a: = ^26-**

Values for (i) and (/) are determined experimentally, and (m/p) can
be obtained from tables of mass absorption coefficients for different
elements and wavelengths, or it may be calculated.

10-

10'

10-

10'

10'

1
z

K Edge 31
L,,, Edge
Mv Edge

\

1

\

\

*

L,„

My

N

^

X

^

\

\

\

V-

■s.

^

10 ' 20 30 40 50 60 70 80 90 ^

0 ]3,06 1.28 0.69 0.42 0.29 0.20 0.15 0.11 A

9M 5.56 3.15 1.90 1.39 1.01 0.76 A

5.33 3.72 A

10-'

10"

E
^10"'

o 10

<

10"
10"

6C 7N 80 llNa 12 Mg 15P 16 S 17 CI 19 K 20Ca 26re

Fig. 16. Value of k, that is, (mi/p)
— (m2/p), for different absorption edges
and different elements. For 15 P the
wavelength for the L edge is Ljj

1

,

^

>

X

/

/

K i I

^

/

•^i

/

1

\ \

i 1

1

0 10 20 30 40 50 60 70

ATOMIC NUMBER, Z

80 90

Fig. 17. The smallest determinable
amount of an element.
From Engsirom (1946)

-L

III-

Fro7n Engstrom (1946)

From the preceding equation it follows that the smallest deter-
minable quantity of the element per unit of surface is a function of k
or (mi/p) — (m2/p) . Accordingly, an absorption edge should be chosen
at which the difference between the mass absorption coefficients is
as large as possible. The variation of k with elements of different
atomic numbers (Z) is shown in Figure 16, which illustrates that the
K-absorption edge must be used for elements of low atomic number,
the Lni edge for elements in the middle of the periodic table (the
Lni is of greater magnitude than Lj or Ln), and the My edge for the

ROENTGEN ABSORPTION HISTOSPECTROSCOPY

131

TABLE IV
Elementary Composition of Muscle (Engstrom, 1946)

Atomic
number

m. X 10»

20 Ca, Ka

3353 X.U.

51 Sb. L<

« 3432 X.U.

Element

g. X cm. -2 X

p

m. X - X 10'
p

p

m. X ^ X 10'

c

6

11500

50

5750

55

6330

N

7

3600

80

2880

85

3060

0

8

5000

115

5750

120

6000

Na

11

70

265

190

285

200

Mg

12

20

350

70

370

70

P

15

170

635

1080

680

1160

S

16

200

790

1580

840

1680

CI

17

60

885

530

945

570

K

19

370

1215

4500

125

460

Ca

20

2

145

3

150

3

Fe

26

20

310

60

330

70

S2.23910-3

21.960-10-3

heaviest elements (the My is the greatest of Mj-y). The smallest
quantity of an element, in g./mm.^, which can be determined is given
in Figure 17. This is based on the fact that the smallest intensity
difference between ii and i2 which can be measured with certainty is
about 5%.

1.0

0.8

?ti:o.6

0.4

02

:\^^=t:S=^

::^

-^

-i

..

V

V 1
\ 1

N

^ '

u

\

A

^^

^-

N

\

\

\ ^

\\'

I

1 \

s^

\

\

\

\'"

rim.

\

\

i\

100f\

\

k

\

\

\

\

V ^^--I^^

WAVE LENGTH, A

Fig. 18. Absorption of roent-
gen radiation of different wave-
lengths in paraffin of varying
thickness. From Engstrom
(1946)

a. 10'

8 10

6C 7N 80 llNal2Mg 15P 16S 17CI 19K20Ca 26Fe

Fig. 19. Appropriate layer thicknesses
for determinations of different elements
in muscular tissue. The solid columns
indicate the layer thickness when the
K-absorptibn edge is used; the hatched,
the Ljjj edge. From Engstrom (1946)

132

MICROSCOPIC TECH N IQUKS

2. Thickness of Sections

The tissue to be subjected to elementary analysis must be frozen -
dried and infiltrated with paraffin by a procedure such as that of
Packer and Scott (page 5). Removal of the paraffin from sections
of this tissue would involve extractions and displacements of ele-
ments by the action of the solutions reciuired. Therefore, it is pref-
erable to carry out the absori)tion analysis without removing the
paraffin. The absorption of the paraffin itself is demonstrated in
Figure 18. The curves were plotted for C2.iH52 (sp. gr. 0.90) ; they
show that, up to 3 A, the paraffin may be used in a layer up to
50-100 fx, while for 5-6 A the layer should not exceed 10 ju,, etc.

A summary of Engstrom's calculations of appropriate section
thickness for analyses of muscle tissue is given in Figure 19. It is
considered prerequisite that the quantity of an element to be
analyzed be about twice the smallest quantity which can be deter-
mined. Figure 19 was derived from data such as that in Table IV,

2

o

I—
<

<

1 0
0,8

0,6
0.4

S02

z
<
a:

0.1

~ —

1 . 1

^*'*^*s-^

^^^^^

4

^^

::;-.^

^<::^

>^

-^■^^2

1^

0.1

0.2 0.3

THICKNESS, mm.

0.4

0.5

Fig. 20. Analysis of
19 K in muscular tis-
sue: (1) Uh; (2)
i^/h.; (3) ii/U for
/, = /.; (4) ii/io for
wavelengths v e r y
close to the absorp-
tion edge and /, =1-2.
See Table IV.
From Engalrdi)).
(1946)

which gives the elementary composition of muscle and the mass
absorption coefficients and absorption capacity of the elements with
the wavelengths used. The wavelength of the K-absorption edge
for potassium (atomic number 19) is 3427 X.U. (1 X.U. = 0.001 A),
and hence, for the potassium determination analysis lines on each
side, 3353 and 3432 X.U. are used in the table. A specific gravity of
1.0 for the tissue has been used in the calculation, and the layer

ROENTGEN ABSORPTION HISTOSPECTROSCOPY

133

thickness is 1 />.. The data in the table have been used to obtain the
curves in Figure 20, and from these it appears that sections 0.2-0.3
mm. thick are appropriate for potassium analysis. Thus, in a volume
of 0.001 fx\. of muscle tissue having a surface of 0.01 mm.- and a
potassium content of 0.3% the quantity analyzed will be 3 X 10"^ g.

Fig. 21. Schematic picture showing the arrangements for analysi.s in a
microcuvette : S, slit in roentgen tube; C, the crj^stal; K, the microcuvette,
F, the photographic fihn. In place of K a microscopic section can be used.
From Engstrom (1946)

Fig. 22. Roentgen tube for primary excitation.
From Engstrom (1946)

3. Apparatus

The arrangement shown in Figure 21 enables the simultaneous
determination of the incident and transmitted radiation intensity.
The former is proportional to the blackening on the upper part of
line L, and the latter to the lower part. The roentgen tubes employed
by Engstrom to produce the radiation were operated by a direct-
current unit manufactured by G. Schonander Co., Stockholm. This

134

MICROSCOPIC TECHNIQUES

unit was designed to develop 1.5 kilowatts at 50 kilovolts or 4.5
kilowatts at 15 kilovolts. The evacuation of the roentgen tubes was
accomplished by a three-stage mercury diffusion pump connected to
a two-stage mechanical forepump. A cooled trap was placed between
the tubes and the pumps.

Roentgen Tube for Primary Excitation. In order to obtain a
line spectrum of great intensity it is best to solder the element whose
line spectrum is desired to the anode. This cannot always be done,
but for elements which lend themselves to this procedure, the anode
is made with six surfaces having a different element or suitable alloy
of it on each surface. A diagram of the tube is shown in Figure 22.
The forged brass body (A) has four openings and bored channels
for cooling. The water-cooled cathode (F) with filament E is sepa-

Fig. 23. Roentgen tube for secondary excitation.
From Engslrom (1946)

rated from A by rubber packing. The anode (B) is insulated from
A, which is grounded, by the porcelain tube C. A rubber packing
separates the cone (D) from C. The anode may be turned to present
its different surfaces to the cathode without breaking the vacuum
by virtue of an Apiezon grease packing between it and D. The
anode is water-cooled. The roentgen beam is passed through slit G
which is covered with an aluminum foil (9 /a thick) fastened on
with Apiezon grease. The flexible tube H connects to the vacuum
apparatus. The filament (E) consists of a 0.25 mm. platinum wire
winding which is coated with an oxide layer (made by burning off
sealing wax) .

ROENTGEN ABSORPTION HISTOSPECTROSCOPY

135

Roentgen Tube for Secondary Excitation. When the element
whose line radiation is desired cannot be soldered to the anode, the
oxide or the powdered metal must be used. However, the difficulties
of suitably incorporating these powders in the surface of the anode
led Engstrom to the use of secondary excitation outside of the high
vacuum. The resulting reduction in intensity was compensated in
some measure by the use of greater wattage and by obviating the
passage of the radiation through a window or membrane.

The roentgen tube is illustrated diagrammatically in Figure 23.
The brass body (A) is 90 X 90 X 100 mm. The cathode filament
consists of a flat platinum spiral with an oxide coating. An iron
cylinder surrounds the filament to direct the electron stream. The
tube is evacuated through H, which is 42 mm. in diameter. The
surface of the anode {B) forms a 12° angle, and the porcelain tube

Fig. 24. Spectrograph for microanalysis.
From Engstrom (1946)

C insulates the anode. Adjustment of the anode position under
vacuum is obtained by a bellows arrangement (D). Water-cooling
is employed for the body of the tube, the cathode, anode, and slit G.
An aluminum foil window ( 9 fi thick.) is placed over the slit, and a
plug (P) is used to hold the element whose line radiation is desired.
This plug and all of the vacuum connections are sealed with rubber.
The composition of the anode is chosen to give the greatest yield of
secondary radiation from P. Hevesy has shown that the greatest
yield of characteristic radiation results when the incident linp radi-

l.'iC)

MICROSCOPIC techniquj:.s

ation has a wavelength 200-600 X.U. shorter than that for the
absorption edge of the element whose secondary radiation is desired.
Accordingly, it is preferable to use K-radiation from copper ( atomic
number, 29) to excite K-radiation from iron (26). If there are no
suitable lines for the secondary radiation the continuous radiation
from an element of high atomic number such as tungsten (74» or
platinum (78) is used.

Spectrograph for Microanalysis. For wavelengths shorter
than 2.5 A it is not necessary to enclose the spectrograph in a vaccum,
since the absorption due to air becomes appreciable only for wave-
lengths greater than 2.5 A. The spectrograph for microanalysis is

^

■» 1-^ niiTi

+

i

1 ^ \

E

1

1

E

E

CM

F

5 mrr

E

}

d

1 1 1

t

',■,■';';

\ — / \ ^ \

1

Fig. 25. Microcuvette. The hatched area is the volume employed.

From Engstrom (1946)

used with these shorter wavelengths; the diagram of the instrument
is shown in Figure 24. Either photographic or ionization measure-
ments may be made of the radiation intensity with this apparatus.
The roentgen tube (.4) delivers a stream of radiation through the
spectrograph slit (B) to the crystal (C) which is mounted in the
holder (D). The monochromatic beam produced passes both above
and through the microcuvette (E) and onto the photographic film
in the holder (F) . The cuvette can be moved relative to the film by
the micrometer screw (G). An ionization chamber (H) may be
substituted for the photographic film; the chamber is directly con-
nected to an electrometer by its central electrode. (An Edelmann
string electrometer was employed.) A Bakelite bar (7) operates a
cog by which the cuvette may be moved into or out of the path of

ROENTGEN ABSORPTION HISTOSPECTROSCOPY

137

the beam when the ionization chamber is used. Adjustments on the
circular scale can be made to some hundredths of a degree, and stops
on the scale can be set to enable rapid and accurate changes for
different wavelengths.

The construction of the metal microcuvette is shown in Figure 25.
The walls in the direction of the radiation are made of thin aluminum
foil, glass, or cellophane sheets. The cuvette capacity is 0.2 fx\. or
greater. Crystals of calcite (d = 3029 X.U.) and rock salt
(d = 2814 X.U.) were used and the ionization chamber was
charged to 150 A'olts.

10 cm

Fig. 26. Vacuum spectrograph for histochemical analysis.
From Engstrdm (1946)

Vacuum Spectrograph for Tissue Analysis. The construction
of the vacuum spectrograph is shown in Figure 26. The roentgen
tube (yi) is joined to the spectrograph by an air-tight rubber gasket.

138

MICROSCOPIC TECHNIQUES

The radiation enters through the adjustable slit (B) and falls on
the crystal (C), whose angle may be adjusted by the micrometer
screw (D). The monochromatized beam from the crystal passes
through the tissue section in holder E. The holder can be moved in
relation to the photographic film placed behind the tissue by means
of the micrometer screw (/) . The film carriage can be adjusted along
scale F at a chosen distance from C. The angle of the tissue holder
can be set on scale G. The spectrograph whose dimensions are
10 X 20 X 6 cm. is evacuated through H. The lid is sealed to the
chamber with rubber and clamped tight. The films used were
12 X 12 mm. and the crystals employed were gypsum {d = 7578

Fig. 27. Mounting of a preparation.
.A is a part of the preparation holder.
B is the preparation itself. C is a cross
of Wollaston wires (platinum) used
to obtain points of reference.

From Engstrom (1946)

X.U.) and mica (d = 9930 X.U.). The tissue section is mounted
over a hole (0.2-2 mm.) in a sheet of metal, Figure 27. The distance
between tissue and film is 1.5-2 mm.

4. Measurement of Density of Photographic Film

The intensity of the radiation is measured by the degree of the
blackening of the photographic film. The blackening may be deter-
mined by photometric estimation of the proportion of visible light
absorbed by the film or by measurement of the proportion of the
silver that has been reduced.

The photometric estimation has been effected by two methods in
Engstrom's (1946, pages 60-64) work. One procedure utilized a
self-recording microphotometer (Siegbahn type) with a thermo-
element and Moll's microgalvanometer, and the other employed the
Caspersson photoelectric apparatus for the determination of light

ROENTGEN ABSORPTION HISTOSPECTROSCOPY

139

absorption in very small areas. The latter method was previously
described in connection with Caspersson's absorption technique for
cells (page 114) and Norberg's technique for fluids (page 120).

1.00
0.90
0.80
0.70
0.60

0.50
0.40

030

0.20

—I 0.01

0.10
0.09
0.08
0.07
0.06

0.05
0.04

003 -

X =

H

Mi _ t^(h\'

p P \X2/

Ml
P

^lg/mm^

0.02

0.01

//g/mm.^

26 Fe 100 ^g/mm."

20 Ca 40

0.02

0.03

0.04

0.05

0.06

0.07

0.08
0.09 ^

0.10 ■^--

Fig. 28. Nomogram for calculating analytical results by equation

From Engstrom (1946)

0.20

030

040

0.50

0.60
0.70
080
0.90
— ' 1.00

shown.

The light absorption in areas less than 1 fi^ can be accurately
measured with this apparatus; however, the size of the grains,
even in the finest films, makes it necessary to measure the blacken-
ing in areas 10 X 10 fi.

140 MICROSCOPIC TECHNIQUES

The raetluxl for tlie estimation of the j)roportion of reduced silver
is based on counting the silver grains in the photographic emulsion
on the film, following the procedure of Glinther and Wilcke (1926i.
The method is only adapted to low densities (upper limit d = 0.27) .
A microscopic enlargement of 600 X is used for coarse-grained film
and 1350X (immersion objective) for fine-grained film. The counts
are facilitated by the use of a netted ocular having 100 squares.
The number of grains in an unexposed film area is determined in a
region immediately adjacent to that exposed.

In a study of the properties of photographic emulsions, Engstrom
(1946, pages 65-72) pointed out that it is necessary to obtain a
curve of the relationship of photographic density to radiation
intensity for every wavelength, emulsion, and set of development
conditions in order to arrive at a suitable working arrangement.
Engstrom investigated the properties of Agfa, Laue, and Printon
films and Ilford High Resolution plates and presented curves of both
density and number of grains as functions of intensity.

Nomogram

Engstrom ( 1946) has published a nomogram for calculating the
analytical results according to the equation shown in Figure 28:

"In this nomogram are included the most important elements, and the analj^-
sis assumes the employment of the K-absorption edge. The wavelengths for the
analysis lines used are seen [Table III]. The two outer pillars in the nomo-
gram indicate the quotient between the intensity of the transmitted and incident
roentgen radiation in the two wavelengths Xi and X2. The amount sought for
(A') of the respective elements is marked out on the vertical lines in the
centre. The figures after the respective elements indicate the amount of the
element in ciuestion at the end of the scale, e.g., 30 Zn 100 /ug./mm.- The
following example shows how it is employed: In determinations of nitrogen,
it is, e.g., found that ii//i is 0.06 and that i^Hi is also 0.06. The straight line
which joins these two points on the outer pillars cuts the ciu-\e for nitrogen
at the point A'. The end point on the nitrogen scale is 1.5 /xg.N/mm.^, and the
scale is divided up into 15 parts, from which it appears that the amoimt of
nitrogen sought for is 0.49 /xg./mm.^"

E. MICROINCINERATION

Microincineration is a valuable technique for the faithful repre-
sentation of the total mineral distribution in tissue sections. In its
present state of development, its reliability is evidenced by the fact

MICROINCINERATION 141

that motion pictures of incinerating sections of skeletal muscle and
of anterior horn cells at 700-800 X magnification reveal no distor-
tion, Scott (1943). The advantage of microincineration over chemi-
cal tests for the determination of the anatomical disposition of
mineral constituents lies in avoiding inevitable displacements and
losses resulting from the use of solutions. In addition, the danger
of fortuitous adsorption of reagents on colloidal protoplasmic sur-
faces is circumvented. The present limitations of the technique lie
in its essential morphological character, which leaves not only
quantitative but qualitative chemical considerations very largely
in the dark. Only a few elements can be detected in incinerated
preparations, and only a rough estimate of the quantity of ash in a
given location can be made.

The microincineration technique was originally developed by
Policard and co-workers in France and was introduced in America
by Scott. The more recent refinements have resulted chiefly from
the careful and extensive researches of Scott and collaborators.
The earlier reviews by Policard (1931-1932) and Policard and
Okkels (1931) and the later ones by Scott ( 1933a,1937,1943) and
Gage (1938) thoroughly cover the development and applications
of this technique.

Engstrom ( 1944) carried out a very nice study on the localization
of mineral salts in striated muscle fibers by employing ultraviolet
absorption followed by microincineration of the same section. By
correlation of both techniques he was able to conclude that the
intensely absorbing isotropic segments which contain the adenylic
acids yielded the ash, w^hereas practically no ash was derived from
the weakly absorbing anisotropic segments.

1. Preparation for Incineration

Some of the earlier work dealt with the use of various solutions
for the fixation of tissue in preparation for microincineration, and
it was found that absolute alcohol or an alcohol-formalin mixture
w^as best since their use resulted in a smaller loss of mineral matter
than was observed with other fixatives. In Scott's (1937) hands the
intracellular distribution of minerals was preserved remarkably
well when the tissue was fixed for 24 hr. in a solution of 9 vol.
absolute alcohol and 1 vol. neutral formalin followed by treatment

./

142 MICROSCOPIC TECHNIQUES

with absolute alcohol, clearing in xylol, and embedding in paraffin
in the usual fashion. But Scott was quick to point out that the
advantages of freezing-drying the tissue are particularly important
in studies of this nature and hence freezing-drying is the procedure
of choice.

Paraffin sections, 3-5 ^ thick, yield the most satisfactory cyto-
logical details. The sections are placed directly on ordinary glass
slides of good quality and no adhesive is required to make them
adhere to the glass. While the presence of water is scrupulously
avoided, a drop of absolute alcohol or liquid petrolatum may be
employed to flatten the sections if necessary (Policard and Okkels,
1930). If alcohol is used, it is allowed to dry; if petrolatum, it is
drained from the slide before the sections are incinerated. The
greatest care must be exercised at all times to avoid contamination
with dust. Absolutely clean paraffin must be used; and the slides
should be washed in distilled water repeatedly, rinsed with filtered
alcohol, dried with a clean lint-free cloth, and stored in a dustproof
container.

It is good practice to cut serial sections using alternate ones for
incineration and the others for controls to be stained and mounted
in the usual manner. Scott ( 1937) pointed out that the use of the
cold knife for sectioning, as recommended by Schultz-Brauns ( 1931) ,
is undesirable since condensation of moisture on its surface may
result in some wetting of the tissue.

2. The Incineration Furnace

The furnace used for the incineration of tissue sections is simply
a quartz tube electrically heated by windings of resistance wire.
Ordinary laboratory muffie furnaces can be used if their tempera-
tures can be properly regulated and if sufficient care is taken to
protect the slides from possible contamination inside the furnace.

Scott (1937) constructed a very convenient furnace capable of uni-
form and reproducible performance. It consists of a quartz tube 24
in. long that is wound with three separate 600 watt heating units
and the whole covered with asbestos insulation. Each heating unit
is controlled by a 44 ohm, 3.2 amp. rheostat. The slides are slowly
moved through the furnace tube on quartz slabs by means of an
electric motor operating through a speed-reducing worm gear

MICROINCINERATION 143

system. A rack running through the tube and extending from both
ends serves to support the quartz slabs as they are moved along.
Further details of the apparatus have not appeared.

3. Scott Incineration Procedure

Scott (1937) gave the following directions for incineration:

1. Gradually bring to 200° over 10 min.

2. Gradually elevate to 280° over the next 5 min.

3. Gradually elevate to 385° over the next 5 min.

4. Gradually elevate to 480° over the next 5 min.

5. Gradually elevate to 580° over the next 5 min.

6. Gradually elevate to 650° over the next 3-5 min.

7. Shut off furnace and let cool for 5-10 min.

8. Remove slides from furnace and place cover slip over section
as soon as cool enough to handle. Seal edges with a mixture of 1 part
paraffin, 1 part beeswax, and 1 part resin (by weight). The use of a
cover slip permits observation with an oil immersion objective, and
it prevents absorption of moisture and efflorescence of the ash.
Greatest care is advised to avoid any air current between the time
of removal from the furnace and sealing the cover slip, since the
ash is easily disarranged.

The practice of covering the ash with collodion or Canada balsam
is undesirable, since it involves the danger of disarrangement and
disturbance of optical properties.

Variation in the above procedure may be necessary for particular
tissues. The greatest shrinkage occurs between 60-70°, and especially
in tissues rich in elastic and fibrous material such as blood vessels.
In order to produce practically all of the shrinkage in advance,
Policard and Ravaut ( 1927) place fixed tissues in absolute alcohol
and bring slowly to the boiling point. However, this procedure is
not advised by Scott for cytological studies because there is the
possibility of dissolving salts.

The passage of a stream of nitrogen through the tube during
incineration was recommended by Schultz-Brauns (1929), and
Tschopp (1929) suggested a similar use of oxygen. Policard (1933b)
employed nitrogen containing a small concentration of oxygen, which
he claimed effects more rapid oxidation. Although satisfactory
results have been obtained with these methods, and they seem to

144 MICROSCOPIC TECHNIQUES

permit lower incineration temperatures which result in less volatili-
zation of chlorides, nevertheless it is sufficient as a rule to carry out
the treatment in air.

4. Microscopic Exainiiiation and Interpretation

Observations of incinerated sections should be made under the
microscope with dark-field illumination provided by a cardioid
condenser. A proper light source is an important factor and Scott
(1937) pointed out that a carbon arc seems to produce excessive
longitudinal aberration, while a Spencer illuminator fitted with a
500 watt projection lamp or a Zeiss Point-0-Light lamp, with proper
centering of the condenser and adjustment of the mirror, are suitable.
Unscreened light is best for observation of minute particles, but
the use of a daylight filter or ground glass is more restful to the eye.
Mercury vapor lamps enable high resolution of small particles but
make recognition of colors difficult.

The use of a comparing ocular with two microscopes is recom-
mended for simultaneous observation of both an incinerated section
and its stained control. Of course, the stained section is illuminated
in the ordinary bright field.

If the incineration has been carried out properly, there will be no
black or brownish carbon deposits. The topographic disposition of
the ash no doubt fairly represents the distribution of mineral con-
stituents in the fixed tissue. That this distribution is exactly main-
tained in the living tissue cannot be said with certainty; however,
Scott ( 1932 ) has found the parallelism that histological sites in
living tissues that absorb ultraviolet radiation (275 m/x) are those
which yield large amounts of ash on incineration.

The only elements that can be identified in the ashed sections
with any measure of certainty are considered in the following
sections.

Sodium and Potassium. It has been assumed that sodium and
potassium yield a fine-grained, faintly bluish-white ash. Policard
and Pillet (1926) attempted the identification of sodium and
potassium as the sulfates by exposing sections, before incinerati(^n,
to the fumes of sulfuric anhydride in order to convert the chlorides
to sulfates. The sulfates are resistant to volatilization during the
ashing, while the chlorides are apt to be lost.

MICROINCINERATION 145

Calcium and Magnesium. The dense white ash seen in the
dark field is due chiefly to calcium with a smaller amount of mag-
nesium. Unless spectrographic means are employed, magnesium
cannot be identified in the presence of calcium in incinerated
sections. As a test for calcium, Moreau (1931) suggested dissolving
ash in a ''microdrop" of 0.1 N hydrochloric acid followed by tlie
addition of a "microdrop" of 0.1 A^ sulfuric acid in order to form
the needle-shaped crystals of calcium sulfate.

Silicon. The identification of silicon can be made with assur-
ance since silica retains its typical crystalline structure during
incineration, and its double refraction when examined with polarized
light serves as an additional means of characterization (Policard
and Alartin, 1933) . The tendency of certain constituents in the ash
to combine with the silica in the glass slide during the heating may
give rise to a misleading appearance. The probability that silica and
calcium salts combine in the incineration process must also be
kept in mind.

Iron. The oxidation of iron that occurs produces a color in the
ash which may vary from yellow to deep red making the identifica-
tion of this element relatively simple. Scott ( 1937) cautions that
care must be taken to avoid contamination with iron from the
microtome knife. A newly sharpened knife will be apt to cause the
most trouble. After 40-50 sections have been cut the number of
particles of iron left in the subsequent sections is practically negli-
gible.

Lead. Exposing incinerated sections to hydrogen sulfide gas
has been employed by Tada (1926) and Okkels (1927) for the
identification of lead as its black sulfide. However, sulfides of other
metals are also black and the possibility of an interference of this
nature should be kept in mind. It is necessary to make sure that
carbon particles due to faulty incineration are not present before
the ash is subjected to the gas, since these particles and the black
sulfide can be easily confused.

Uranium. Policard and Okkels ( 1930) claimed to have detected
uranium, by its fluorescence under ultraviolet radiation, in ashed
sections from animals poisoned by this element. Fluorescences may
be produced by impurities too, and hence this criterion is not a very
rigorous one.

146 MICROSCOPIC TECHNIQUES

5. Quantitative Estimation of Ash

Attempts at the quantitation of the relative amounts of ash left
by various structures were made by Schultz-Brauns (1931) based
on a standardized development of photomicrographs. However, this
method has many inherent difficulties and can yield little.

Scott (1933b), and Williams and Scott (1935) developed a photo-
electric apparatus to measure the intensity of light reflected from
the ash. This light intensity is roughly proportional to, and serves
as an approximation of, the quantity of the mineral residue. While
the method obviously leaves much to be desired, as Scott would
no doubt be the first to admit, it is capable of furnishing some
information that at present can be obtained in no other way.

An idea of the sensitivity of the apparatus, as assembled and used
by AVilliams and Scott, may be gained from the fact that, with a
magnification of 700X, the ash produced in a 5 ju, section by a single
hepatic cell nucleus results in a galvanometer deflection of about
half-scale (25 cm.).

Williams and Scott Photoelectric Apparatus. The microscope
illumination is furnished by a 6 volt, 108 watt ribbon filament
projection lamp enclosed in a ventilated housing. This lamp is
supplied through a constant-voltage regulator and a step-down trans-
former. The slide is held by a special mechanical stage which has
rack and pinion adjustment laterally and vertically and a fine screw
adjustment axially; and the microscope, mounted horizontally, is
fitted with a Zeiss aplanatic 1.2 condenser, a Leitz No. 3 objective,
various oculars, and a clamped-on 90° reflecting prism. A fixed
diaphragm made of a disc slightly larger than the aperture of the
objective is fitted into the ring on the condenser lens. The light
emerging from the ocular is reflected by the prism to a mirror which
in turn reflects it to the photocell.

The gas-filled photocell is mounted in a light-tight copper box
which serves as an electrostatic shield as well. The cover of this
box carries rotatable and interchangeable 3 in. white cardboard discs
with various size openings to determine the illuminated area on the
photocell. A shutter under the disc permits exposure of the photocell
when desired. Within the copper box containing the photocell, a
rD54 Pliotron tube is mounted with its 10^ ohm high-resistance
shunt. The photocell is connected directly to this tube, and the

ANALYTICAL ELECTRON MICROSCOPY 147

connections to the DuBridge and Brown (1933) amplifier and the
photocell B battery are made thi'ough a shielded flexible cable, which
also carries the galvanometer leads. The amplifier is powered by-
storage cells ( 12 volts) . A Leeds and Northrup type R galvanometer
is used. The entire apparatus is mounted in a dark room, and black
felt is used to prevent stray light of the apparatus from reaching
the photocell.

Procedure. The image of the lamp filament is sharply focused
on the center of the cardboard screen with the dark-field diaphragm
removed and the slide in place. It is convenient to place a 12 power
convex lens in front of the lamp housing to enlarge the image to
about 4 in. The dark-field diaphragm is inserted, and, with the room
almost completely dark and the felt curtains in place to eliminate
stray light, the image of the ash is focused on the screen with the
axial adjustment of the mechanical stage. With the other adjust-
ments of the stage, the desired area is brought to the center of the
screen and the photocell box is moved so that the aperture is
properly set. Then the shutter is pulled and the galvanometer deflec-
tion is observed. The reading obtained when the clear glass of the
slide is placed in the optical field is substracted from the first
deflection, and the resulting figure is proportional to the quantity of
ash on the area taken when this quantity is small. Similar data
obtained for other areas enable a comparison of relative amounts
of ash without regard to the absolute quantities involved.

F. ANALYTICAL ELECTRON MICROSCOPY

The higher resolving power of the electron microscope has
enabled finer morphological differentiations in biological material
than were hitherto possible with the best optical microscopes. How-
ever important this advantage in other fields, as used today its
purely morphological value limits its contributions to cyto- and
histochemistry. However, a beginning has been made toward the
use of an analytical electron microscope, not merely as a means of
obtaining high magnifications, but as a tool for the identification
and localization of certain metallic elements in biological prepara-
tions. To date only calcium and/or magnesium can be identified and
localized. These elements can be detected with a sensitivity of about
1 X 10"^^ g- per kilogram wet weight of tissue (muscle). For the

I*.

148 MICROSCOPIC TECHNIQUES

conception of this possibility and the ingenuity to carry through to
fruition, credit is due to Gordon H. Scott and his associates, J. H.
McMillen and D. JVI. Parker, wlio carried out their investigations
at Washington University.

Scott and co-workers utiHzed the well-known fact that when
metals or certain of their compounds are heated in vacuo they emit
a number of electrons depending in part upon both the nature of
the metal and the temperature. Hence identification of the metals
might be possible on the basis of their differential emission of
thermally excited electrons. The localization of these metals in tissue
sections would be possible since the electrons emitted could be
focused by the magnetic lenses of the electron microscope to yield an
image, on a fluorescent screen, of the topographical disposition of
the emitting substances.

The first apparatus developed for this purpose was that of
McMillen and Scott ( 1937) but, since a number of changes have
been made, only the later apparatus of Scott and Packer ( 1939a)
will be described.

The Scotl-Packer Analytical Electron Microscope

A diagram of the instrument is given in Figure 29. Basically it is
an electron microscope of relatively low magnifying power fitted
at one end with a chamber (C), lined with a material capable of
fluorescence by electron streams for the visualization of the electron
image, and at the other end with a special cathode (B), carrying
a tissue support (.4) to hold the paraffin sections that are employed.
The microscope tube is made of brass, and is 1 m. long and 63 mm.
in diameter. Two magnetic lenses (Lj and Lo), swung on gimbals,
surround the tube and are free to move along it as well as rotate
around it to some degree. Each lens is composed of 1550 turns of
No. 22 enameled, single-cotton-covered copper wire wound on a
copper inner ring 75 mm. in diameter. The coils are enclosed in
sheaths of soft iron having a wall thickness of 3 mm. The lenses have
an axial width of about 43 mm. It is apparent that the object-image
distance is fixed; therefore focusing for any magnification must be
accomplished by altering the power of the lenses. This is done by
varying the current passing through the lens coils. The power for
these coils is supplied from two 30 volt banks of storage cells in
parallel.

ANALYTICAL ELECTRON MICROSCOPY

149

The tissue holder assembly is made of a 25 mm. glass tube fitted
snugly with a brass sleeve soldered to a heavy brass plate. A
shoulder on the plate fits into the microscope tube and the joint is
rendered vacuum-tight with Apiezon Q sealing compound. Proper
centering alignment is maintained by brass cylinders {A and ^i)
which, fit over the glass tube. The flat top of the nickel cathode
cylinder (B) serves as support for the tissue sections and the
cylinder itself is held by a wire fixed to its wall. The cathode is
heated by a coiled filament (H), which consists of 10 mil tungsten
wire supported by a ceramic core not indicated in the diagram. The

Fig. 29. Diagrammatic sketch of the essential features of the electron
microscope. A, tissue holder (cathode) support; Ai, inner shell of same; B,
nickel cathode; C, fluorescent screen; D, diaphragm; H, heating filament for
cathode; Li, objective magnetic lens; L2, ocular magnetic lens; P, pumping
port; T, transformer. Other letters and symbols are standard usage in vacuum
tube technique. From Scott and Packer (1939a)

cathode filament is supplied with current from two 6 volt storage
batteries in series which are insulated from ground for 15.000 volts.
The insulation is required since the tissue holder assembly and
batteries are at a potential of 6000 volts with respect to the grounded
microscope tube.

The fluorescent screen (C), which is sealed to the microscope
tube with wax, is a commercial oscillograph type. The bare portions
of the inner walls are coated with colloidal graphite (Aquadag) to
furnish electrical conductivity from the screen to the ground. This
prevents the accumulation of charge, which would distort the image
on the screen.

150

MICROSCOPIC TECHNIQUES

The diaphragm (D) has an aperture of 25 mm. over which are
placed a few fine wires to serve as fiduciary marks in focusing the
lens Lo. The image is formed with lens Li (Fig. 30) on the aperture
of the diaphragm in order to reduce spherical aberration. The half-
wave rectifier and filter system (Fig. 29) supplies the potential
difference required for accelerating the electrons. The current pass-
ing through the 10^ ohms resistance placed across the high-voltage
leads is measured by the microammeter from which the magnitude
of the accelerating voltage can be obtained. The rectifier is supplied
by the secondary of a transformer that is insulated from the primary
for 15,000 volts. A variable resistance in the primary circuit of this
transformer permits the choice of the high D.C. voltage employed.

Fig. 30. Diagrammatic representation of the electron path and consequent
image formation. A, cathode and support; C, fluorescent screen; D, diaphragm;
E, batteries for heating filament of cathode; F, schema of magnetic field of
the lenses Li and La; V, high-voltage source. From Scott and Packer (1939a)

Constancy is maintained in the accelerating voltage in order to
prevent a distortion, similar to chromatic aberration in optical
systems, by the use of a voltage regulator, of the saturated-core
transformer type, in the primary circuit of the transformer. Another
precaution taken to avoid distortion of the image is the placing of
the batteries, high-voltage supply, and all iron objects at quite a
distance from the microscope.

In order to compensate, at the magnification used (< 150x),
for the deflection of the electron stream by the earth's magnetic
field, the first lens (Li) is tilted.

Water-cooling coils of copper tubing are employed to remove heat
from the microscope tube. One coil is wound around the tube on the
image side of lens Li and another at the junction of the tube and the

ANALYTICAL ELECTRON MICROSCOPY 151

object holder. The latter is essential to absorb the 50-60 watts of
power given off by the heated filament, which would tend to soften
the sealing compound and weaken the vacuum.

The microscope is evacuated through a port (P, Fig. 29) by
means of two double-stage mercury vapor pumps in parallel employ-
ing a Cenco Hyvac oil forepump. A vapor trap ccioled with carbon
dioxide in butyl alcohol is placed between the microscope and the
pumps, and the glass-to-metal connection is sealed with black
vacuum wax. Pressures are measured with the ionization gauge of
Montgomery and Montgomery ( 1938) employing a No. 47 radio
tube of Radio Corporation of America. A portable power pack is
used so that it can be employed with several vacuum systems to
obviate duplication of expensive meters. A pressure of 10"^ mm.
mercury or less is sufficient for the electron microscope, and the
sensitivity of the pressure gauge is about 7 X 10"^ mm. mercury per
microampere ion current.

Manipulation

Preliminary experiments with salts of sodium, potassium, calcium,
magnesium, and iron demonstrated that very bright images were
formed on the screen by calcium and magnesium, weak ones by
sodium and potassium, and none by iron. By maintaining a cathode
temperature of 700-800° for an hour the sodium and potassium
were volatilized so that the bright image could be safely assigned
to calcium and magnesium. For these experiments, ashless gelatin
impregnated with the chlorides was hardened in 10% formalin,
dehydrated in alcohols and embedded in paraffin. Paraffin sections
( 10 fi.) were placed directly on the prepared cathode.

The nickel cathode is prepared for use by polishing with optical
rouge and washing with water and nitric acid. It is then coated
with a mixture of 40% barium carbonate and 60% strontium
carbonate in a 2% solution of nitrocellulose in amyl acetate. The
barium-strontium mixture serves to increase the emission of the
calcium and magnesium in the tissue by about 1000%, by activa-
tion, and at the same time the mixture emits electrons itself to give a
contrast background on the screen. When completely dry, a 10 /x
section of tissue embedded in paraffin, prepared by the freezing-
drying technique (see page 3), is placed on this surface and
smoothed down with a steel needle. The cathode is then inserted into

152 MICROSCOPIC TECHNIQUES

the microscope tube and the vacuum pumps are started. When the
pressure falls to 10 •'' mm. mercury, or less, the cathode-heating
filament is turned on and the temperature is gradually elevated
over an hour or more to the operating level. The slow heating is
essential to avoid distortions in structure, to minimize or prevent
curling on the cathode, and to approximate microincineration condi-
tions in order to permit a comparison. The practice is followed of
heating the cathode higher than operating temperature for a short
time to volatilize the sodium and potassium and to initiate active
emission of electrons. When this activation period is over, and the
operating temperature obtained, the high-voltage and lens currents
are turned on. The electron image formed on the screen is compared
with stained or incinerated control sections. When the tissue curls
away from the cathode and is improperly oxidized, dark areas appear
in the image and carbonization is evident by optical examination.
The magnification is altered by changing the position of the mag-
netic lenses on the tube and then bringing to focus by adjustment
of the lens current. Electron-accelerating voltages of 5000-6000 were
employed. Various portions of the section are focused on the center
of the screen by virtue of the mobility of the lenses.

Photographs of the image are made with a high speed camera
(/ = 2.9) at a camera magnification of 0.73 on Eastman Kodak
Superspeed Ortho Portrait or Panchro-Press film. Exposures of 1-5
sec. are usually employed. The films are developed with Eastman
Kodak D-72 developer. Some photographs obtained in this fashion
are shown in Figures 31 and 32.

G. RADIOAUTOGRAPHY

The novel and thus far very limited technique of radioautography
has been employed in a few instances for the localization of radio-
active elements in tissue sections. The use of these isotoi)es as tracers
in biochemical investigations, particularly as the result of the
pioneering work of Hevesy, is a well-established device; however
histological distribution cannot be determined quantitatively, as yet,
on the basis of radioactivity, by any very satisfactory means,
since the order of the intensity of the radiation produced in the
quantities of tissue commonly employed for histological examina-
tions is far too small to permit suitable measurements M'ith the

RADIOAUTOGRAPHY

153

Geiger-Miiller counter or the electroscope. Radioautography, based
on the ability of emanations from radioactive elements to affect the
photographic plate, is an attempt toward the solution of this difficult
problem. Tissue sections containing radioactive elements leave their
"autographs" on photographic plates when placed in contact with

Fig. 31. Emission electron micro-
graph (X300) showing calcium and
magnesium distribution in rectus ab-
dominus muscle of cat. Note strong
cross-bandings in muscle fibers. From
Scott (1943)

Fig. 32. Emission electron micro-
graph of cat gastric mucosa (fundus)
showing calcium and magnesium dis-
tribution (left), compared with a
microincinerated section from the
same animal (right). Magnification
about X75. From Scott (1943)

them for a sufficient period. When developed, these "autographs"
indicate to some degree the relative distribution of the substances
responsible for the radioactivity.

Historically, the first use of radioautography was made by
Lacassagne and Lattes ( 1924) for the demonstration of polonium in
tissue. Since that time the usefulness of this technique, as well as all
others employing radioactive tracers, has been greatly expanded by
the recent revolutionary developments which have made possible
the preparation of radioactive isotopes of elements that occur in
living systems. Limiting factors in regard to the suitability of a
radioactive isotope for studies by radioautography are the nature
and intensity of its radiation and its half-life period. The duration
of the photographic exposure will depend on these factors as well as
on the concentration of the isotope in the tissue. The half-life
periods of the principal artificial radioactive elements that might be
used in tracer studies are given in Table V.

Perhaps, the greatest deficiency of the technique of radioautogra-

154

MICROSCOPIC TECHNIQUES

TABLE V. Principal Artific^ial Radioactive Isotopes Used as Trace Elements
as Compiled by Pool and Kurbatov (1943)

Radioactive

Atomic

Intensity of

Type of

element

Half-life

weight

activity

radiation

0

2 . 1 min.

15

Strong

+ 0

N

9.93

13

Strong

+ 07

Mg

10.0

27

Strong

-0y

Co

11 0

60

Strong

-07

C

21.0

11

Strong

+0

Ag

24.5

106

Strong

+ 0

I

25.0

128

Strong

-0y

CI

37.5

38

Strong

-0y

In

54

116

Strong

-0y

Zn

57

69

Strong

-0

Ba

1.42 hr.

139

Strong

-0y

F

1.8

18

Strong

+ 0

Se

1.81

75

Weak

+ 0

Y

2.0

88

Strong

+0

Cr

2.27

55

Weak

-0

Mn

2.59

56

Strong

-0y

Si

2.60

31

Strong

-0

Ni

2.6

63

Strong

-0y

Ti

3.1

45

Strong

+ 0

Sc

4.0

43

Strong

+ 0y

Br

4.45

80

Weak

y

Ab

7.5

211

Weak

(xy

K

12.4

42

Strong

-0y

I

12 6

130

Strong

-0y

Cu

12.8

64

Strong

-0, +0

Au

13.0

196

Weak

-0

Pd

13.0

109

Weak

-0

Zn

13.8

69

Weak

y

Na

14.8

24

Strong

-0y

Pt

18

197

Weak

-0

Co

18.2

55

Strong

-\-0y

W

1.01 day

187

Weak

-0y

Sn

1.05

121

Weak

-0

As

1.11

76

Strong

-0,+0y

La

1.70

140

Strong

-0y

Ni

1.5

57

Weak

+ 0

Br

1.66

82

Strong

-0y

R A DIO A U TOG R AP H Y

155

TABLE V (Concluded)

Radioactive

Atomic

Intensity of

Type of

element

Half-life

weight

activity

radiation

Cd

2.3 days

115

Strong

-0y

Au

2.7

198

Weak

-Py

Mo

2.8

99

Weak

-/3

Ag

7.5

111

Strong

-0

I

7.8

131

Strong

-M

Ca

8

41

Weak

y

Ag

8.2

106

Strong

y

Sn

10.0

123

Weak

-/3

P

14.5

32

Strong

-0

V

16

48

Weak

+ 0y

As

16

74

Strong

-^, + l3y

Rb

19.5

86

Strong

-/3

Cr

26.5

51

Weak

y

Be

43

7

Weak

y

Fe

47

59

Weak

-fiy

Sr

55

89

Strong

-0

Sb

60

124

Strong

-0y

Zr

63

93

Weak

-0

Ti

72

51

Weak

-0y

W

74.5

185

Weak

-0

Sc

85

46

Strong

-0y

S

88

35

Weak

-0

Ta

97

182

Weak

-0y

Y

105

86

Strong

y

Ca

180

45

Weak

-0y

Zn

250

65

Weak

+ 0y

Mn

310

54

Weak

y

Cs

1.7 yr.

134

Weak

-/3->

V

1.71

47

Weak

7

Na

3.0

22

Weak

+ ^7

Co

5.3

60

Weak

-0y

H

31

3

Weak

■ -&

Ra

1590

226

Strong

a

C

10000

14

Weak

-0

U

7.1 X IQs

235

Weak

a.

U

4.5 X 109

238

Weak

a.

Rb

1 X 10"

87

Weak

-0

156 MICROSCOPIC TECHNIQUES

phy lies in its inability to reveal distribution in the finer structures,
and to this lack of resolving power must be added the further draw-
back that, quantitatively, only a rough approximation is possible.
However, there is the considerable advantage, inherent in the use of
radioactive elements regardless of whether a histological or gross
tissue study is involved, that very small quantities of an element
introduced into a biological system can be followed without reference
to, or interference from, the large stores normally present. The
amount of a radioactive element that can be detected is fortunately,
very minute. According to Hamilton (1941) a total of 2 X 10^
(i particles, with an average energy of at least 150 Kev., are
required to strike each cm.^ of photosensitive surface to yield a
satisfactory image.

Reviews dealing with radioautography have been presented by
Hamilton (1941-1942) and Simpson (1943), and important physical
data have been furnished in a review by Kurbatov and Pool ( 1943) .

Preparation of Radioautographs*

Both fresh-frozen and paraffin sections of tissue have been used
to obtain radioautographs. In general the paraffin sections give the
hest results since they can be cut thinner and are less subject to
distortion. It is essential that the sections be of uniform thickness
and free of wrinkles. There would be a particular advantage in the
use of freezing dehydration (page 3) for the preparation of the
paraffin-infiltrated tissue. The diffusion of the radioactive substances
during fixation and dehydration in solutions would be eliminated
and a more authentic "autograph" could be obtained.

As examples of procedures which have been used the following
may be cited: Hamilton, Soley, and Eichorn (1940), in a study of
radioactive iodine in thyroid tissue, removed the paraffin from 3-5 /x
sections with xylol, dipped the slide containing the sections in dilute
celloidin, allowed it to dry, and obtained a celloidin film over the
sections about 1 fx. thick. The sensitive surface of the photographic
film was placed in contact with the celloidin surface. Harrison,
Thomas, and Hill ( 1944) in an investigation of the distribution of

* See Bibliography Appendix, Refs. 20, 21, 22, 2S, and 30.

RADIOAUTOGRAPHY 157

radioactive sulfur in the wheat kernel, employed paraffin sections
25-50 p. thick which were covered directly by a layer of aluminum
foil 0.8 jx thick. The sensitive photographic surface was placed in
contact with the foil.

The greater the distance between the tissue and the photosensitive
surface, the poorer the resolution in the radioautograph due to
scattering of the radiation. It is preferable that this distance be kept
under 1 mm. Ultraspeed x-ray film has been extensively used, but
it has the disadvantage of producing grainy enlargements. Harrison,
Thomas, and Hill (1944) recommend a fine grained panatomic film
when it is possible to have longer exposures. The photographic film
over the sections on a glass slide is covered with another slide and
the whole bound firmly together with cellulose tape. All of these
operations are carried out in a dark room, of course. After wrapping
the slide in light-tight black paper, it may be placed in a cold place
for the duration of the exposure. It is advisable to keep the sections
cold to inhibit any tendency toward diffusion of the radioactive
element. After exposure, the sections are stained in the usual manner
to bring out their morphology, and compared to the developed
"autographs" with the aid of a dissecting microscope.

More recently, Belanger and Leblond (1946) extended the useful-
ness of radioautography by the ingenious expedient of spreading a
photographic emulsion directly on the sections. This not only permits
a more intimate contact between the tissue and the photographic
surface, but it obviates the matching of the "autograph" to the cor-
responding histological detail, which is particularly difficult at higher
magnifications. The possibilities of this technique merit a more
detailed description of the procedure.

Belanger and Leblond Technique

Preparation of Photographic Emulsion. Soak lantern slide
plates (medium contrast, Eastman) in distilled water at room
temperature. When the gelatin swells, remove from the water. With
a glass knife scrape off the gelatin, and melt it in a beaker placed
in a 35-40° bath. Carry out this procedure and all others in which
the emulsion is used in a dark room. A Wratten "Safelight — No. 1"
(Eastman) may be used at a distance of about 3 ft.

158

MICROSCOPIC TECHNIQUES

m. int

h.int.

X""-

mol. tub

Nj. ^m. int.

O

i

">!.

2A

3 A

V4 «»^

3B

■n js.

4 A

Fig. 33. Radioautographs (A) and corresponding stained sections (B) (X8).
White areas in radioautographs are exposed parts of film. (1) Thorax of adult
mealworm, transverse section, paraffin; (2) abdomen of adult mealworm, trans-
verse section, paraffin; (3) abdomen of wax moth larva, longitudinal section,
frozen; (4) abdomen of wax moth larva, transveise section, frozen; (g)
ganglion; (gon.) gonad; (hy.) Iwpodermis; (mal. tub.) malpighian tubule;
(m. int.) midintestine; (mus.) muscle; (h. int.) hind intestine; (rep. org.)
reproductive organs; (s. g.) silk gland. Fi(ii)( Lindsay and Craig (1943)

1 !

Mk

L .''

■•i«

W

I^:' .

^

'^^:v^

"/.*'■'

- ii

■M^

0^ ■

• ■

*

't.

_.i-^

J

Fig. 34. Radioautographs of adult rat tissues. (1) Cross section through the
lower limb (X50). The radiophosphorus is in the diaphysis of the tibia and
the fibula. Arrow A points to heavy periosteal layer in the tibia, (i?) Para-
median longitudinal section of thoracic vertebra (X50). Arrow B indicates
phosphorus deposit in the ossifying neural arch. (5) Cross section of the lower
jaw (Xl7). The deposition of radiophosphorus clearly outlines the mandible.
In the right portion of the bone an incisor tooth is developing and is also
impregnated with the radioactive element. (4) Section of the thyroid from an
adult rat (X70), treated with radioiodine. The tracheal cartilage is visible at
the bottom of the figure. The thyroid follicles show a reaction due to radio-
iodine. Friim Belnnger and Leblond (19.'/i)

160 MICROSCOPIC TKCHNIQUES

PROCEDURE

1. Prepare 10 /x })araffin sections and attach to slides with egg
albumin.

2. Dry, deparaffinize with xylol, and carry through absolute
alcohol to 1% celloidin in alcohol-ether.

3. Drain off excess celloidin by standing the slides in an empty
Coplin jar.

4. Place in 70% alcohol for about 1 min. to harden the celloidin,
and dry at room temperature.

5. In the dark room, pipette 5 drops of the melted photographic
emulsion on to each slide and spread evenly with a camel's hair brush.
Carry out this operation a little below 40° on a hot plate to prevent
premature gelling of the emulsion. (The temperature should be held
below 40° with this emulsion to reduce the fogging.)

6. Allow to cool and dry, and place the slides horizontally in a
dustjiroof, light-tight box for the duration of the exposure.

7. x\fter the exposure develop for 3-4 min. in Kodak developer
D72 at 18-20°. Wash rapidly in water and fix for about 10 min. in
5% thiosulfate at the same temperature. Wash finally in cold run-
ning water for about 30 min. The black silver deposit indicates the
site of the radioactive element.

8. Counterstain in Coplin jars cooled in running water. (Methyl-
ene blue and Harris hematoxylin may be used for radiophosphorus
"autographs" and Harris hematoxylin for those of radioiodine.)
With methylene blue place slides in a 1% alkaline soln. for about
30 min. and rinse in running water until the stain is removed from
the gelatin coating. With Harris hematoxylin, stain lightly to avoid
the need of differentiating. (Artifacts caused by gelatin swelling and
disengagement of the sections disappear when the slides are thor-
oughly dried after each operation.)

9. Pass through several changes of 95% alcohol, absolute alcohol,
and xylol, and mount in Canada balsam. (Clearing in oil of origanum
before mounting will also give good results.)

Discussion

Points on various procedures required for particular studies may
be obtained by referring to some of the applications already made.
To date, most of the investigations employing radioautography

RADIOAUTOGRAPHY 161

deal with the isotope P^-. Thus its distribution was studied in bones
by Dols et al. (1938) and Belanger and Leblond (1946), in tomato
fruits by Arnon et al. (1940), in squash plants by Colwell (1942),
and in insects by Lindsay and Craig (1942). "Autographs" have
been obtained in bone studies with radioactive calcium and strontium
by Pecher (1941) and Treadwell et al. (1942). The distribution of
radioactive lead in the animal body was investigated by Behrens
and Baumann (1933a,b), and thyroid studies were carried out with
radioactive iodine by Hamilton et al. ( 1940) , Gorbman and Evans
(1941), and Belanger and Leblond (1946). Harrison, Thomas, and
Hill ( 1944) employed radioactive sulfur for a radioautograph survey
of the distribution of this element in wheat. Many new applications
are constantly appearing.

CHEMICAL TECHNIQUES

"By calling attention to the cell I desired to provoke
investigators to inquire into the processes within the
cell, to define that which happens within these small-
est elementary organisms. And it was self-evident that
an exact definition could be nothing else than to find
the chemical and physical foundations upon which

vital phenomena and the activity of the cell are
based."

ViRCHOW

as quoted by Paul Klemperer in Some Recent Biologic
Investigations and Their Significance for Pathology,
J. Mt. Sinai Hosp. N. Y. 14: 442 (1947/48).

INTRODUCTION

The chemical techniques to be described are all of the quantitative
variety and they differ from their macro counterparts primarily as
regards the volumes employed and the mode of handling them. In
general, the same reactions and concentrations of reagents are used
in both. The degree to which the localization of the chemical constit-
uents in tissues and cells is limited, in these techniques, largely
depends upon the degree to which the anatomical parts can be
isolated mechanically in preparation for their separate analysis. The
precedures most commoly used are: (a) the preparation of serial
microtome sections of tissue and analysis on each of selected sec-
tions, (fc>) isolation of cells or cellular particulates by centrifugation
for their separate analyses, or (c) use of micro dissection to obtain
the part to be analyzed. It is considerably more of a problem, as a
rule, to obtain a satisfactory sample for analysis than to perform
the analysis itself. While the ultimate goal of being able to apply
quantitative procedures in situ to biological material is still essen-
tially beyond the present horizon, the use of these chemical
techniques can lead to the acquisition of knowledge which can now
be obtained by no other means.

It should be pointed out that in the interests of simplicity and
accuracy certain well-established procedures of macroquantitative
analysis are best avoided in work on the level considered here. The
procedures to be given are those of the original authors, but the
laboratory worker should introduce his own simplifications of the
following type at every opportunity : (1) Avoid quantitative trans-
fers— rather remove an aliquot. (2) Avoid dilution to a given volume
in a vessel; this necessitates the calibration and marking of the
vessel — rather dilute by adding a known volume of liquid with
a pipette. (5) Employ pipettes calibrated to deliver rather than to
contain — this obviates the necessity of rinsing the pipette. (4) Avoid
filtration when it is possible to separate a precipitate by centri-
fugation.

165

i. GENERAL APPARATUS AND
MANIPULATION

A. VESSELS, STOPPERS, HOLDERS, ETC.

Vessels. Most of the reactions employed in the various chemical
techniques are carried out in simple glass vessels. The tube shown
in Figure 35 is especially useful; it is nothing more than a small test
tube having a total capacity of 0.25 ml. (available from A. H.
Thomas Co. and E. Petersen, Carlsberg Laboratory, Copenhagen
Denmark). Norberg (1937) employed the tubes (Fig. 36) for use
in centrifugation and the apparatus shown in Figure 37 for removal
of supernatant fluid from centrifuged precipitates. By applying
suction at A the fluid is drawn into the reservoir; the low-power
microscope is used to enable careful control of the operation. As
indicated in Figure 37, the tube may be surrounded by a larger
vessel filled with a clear liquid, such as alcohol, to permit better
observation, particularly when the vessel B (Fig. 36) is used, since
the bottom part of the tube has a dark zone due to its form.

The tubes may be cleaned conveniently by immersing in the clean-
ing liquid, heating to drive the air out of the tubes, cooling to let
them fill up with the liquid, and shaking out the liquid from each
one. Usually the process is repeated two or three times. To place
films of liquid across the upper portion of a reaction tube, as in
iodometric titrations, Holter and Doyle ( 1938) employed the vessel
shown in Figure 38, which has a total volume of 0.20 ml. These
vessels must be given an inner hydrophobic coating to prevent the
liquid from spreading on the glass surface. The method of doing
this is described on page 169.

Levy (1936) used the 2.5 ml. tube illustrated in Figure 39 for the
Kjeldahl digestions and Linderstr0m-Lang, Weil, and Holter (1935)
employed the two-piece unit shown in Figure 40 for ammonia dis-

166

«Id>

^

u

Fig. 35. Reaction
vessel in holder.
From Linderstr0m-Lang
and Holler (1933a)

W V

10

'-20 mm

Fig. 36. Centrifuge

reaction vessels.
From Norberg (1937)

rO

"-10 cm.

Fig. 37. Arrangement for removing

supernatant fluid over a precipitate.

From Norberg (1937)

■ 5 mm

// w

^WA

Fig. 38.

(r\

Ml

-^ -^-S mm.

Fig. 39.

\y

Fig. 40.

Fig. 38. Reaction vessel for iodometric titration. In neck, lower film is
sulfuric acid, upper film starch solution. From Holler and Doyle (1938)

Fig. 39. Kjeldahl digestion tube. From Levy (1936)

Fig. 40. Vessel for ammonia distillations (no longer used). From Ldnder-
str0m-Lang, Weil, and Holler (1935)

168

CHEMICAL TECHNIQUES

tillations, e.g., in arginase measurements. The ammonia was distilled
from vessel A into cap B, which was coated internally with paraffin
and charged with standard acid. Ramsay grease (thick) was used
to seal the parts together. Later it was found preferable to abandon
the use of this form of vessel for ammonia distillation (Briiel et al.,
1946) (see page 283).

Glass diffusion cells for the distillation of ammonia were described
first by Conway and Byrne (1933), and later by others (Figs. 41-
43) . Ammonia diffuses from the outer well into standard acid con-

F

67 mm.-
61 mm.-
40 mm.-
35 mm-

^ "R^J

^^- ~^

"' "

Ie

in

Fig. 42, Gibbs and
Kirk (1934) diffusion
cell (cross section, one
half actual size).

D

\it ill UJ lU ll< OJ

Fig. 41. Conway and Byrne (1933)
diffusion cell. Above, top view;
below, vertical, section on line AB.

Fig. 43. Kinsey and Robinson
(1946) diffusion cells: upper, top
view; lower, side view.

tained in the center well in the types shown in Figures 41 and 42
(available from Microchemical Specialties Co.). The Kinsey and
Robison (1946) apparatus (Fig. 43) consists of a Lucite plate 0.5
in. thick with rings 1 mm. deep having inner and outer diameters of
14 and 18 mm., respectively, reamed out of the plastic for one form
of cell (A); for the other form (B), rings of the same diameter but
8 mm. deep are reamed out and a center hole 4 mm. deep and 8 mm.
in diameter is drilled. In the A form cell, two glass vials are used
alone. Ammonia diffuses into the receiving solution placed in the

GENERAL APPARATUS AND MANIPULATION 169

bottom of the outer or larger vral. When the vial is inverted and
set on the plate this solution forms a hanging drop over the liquid
which is liberating ammonia.

A small open porcelain dish {Micro chemical Specialties Co.) (Fig.
44 heknv) was used by Kirk and associates as a titration vessel.

P^ig. 44. Titration dish, actual size.
Frotn Kirk and Bentley (1936)

Coating Vessels with a Hydrophobic Layer. At times it is
desirable to coat reaction vessels with a hydrophobic layer to pre-
vent aqueous liquids from spreading on the glass surface, as in
iodometric titrations where liciuid films are placed across the lumen
of the neck of the titration tube. Linderstr0m-Lang and Holter
( 1933a) used paraffin and Holter and Doyle ( 1938) employed
ceresine. The procedure followed by the latter was to boil about 50
vessels for 5-10 min. in 75 ml. of water to which 0.1 g. ceresine was
added. After the water had cooled, the vessels were emptied and
dried for at least 3 hr. at 100-110°.

The procedure finally employed at the Carlsberg Laboratory for
paraffin coating was described by Brliel et al. (1946). The clean,
dry glass tubes are immersed in melted paraffin at 150-200° (the
synthetic paraffin used had a melting point of 82°), picked out one
at a time with forceps, quickly emptied and rotated in a clean towel
between the fingers until the paraffin solidifies. A heavy layer of
paraffin on the bottom of the tube and a thinner one on the upper
part is desirable. The outside of each of the tubes is wiped free of
paraffin and they are stored protected from dust and fumes. After
the vessels have been used, they are cleaned by rinsing first with
water, then with acetone, hot toluene, acetone, and water in the order
given.

Stoppers. For most work it is sufficient to stopper reaction tubes
with a short piece of rubber tubing one end of which is plugged with
a glass bead or short piece of glass rod. A stopper consisting of a
cap with a small hole (Fig. 45) is useful in some cases as in the
addition of alkali in the method of Linderstr0m-Lang and Holter
(1933b I for ammonia. In this same method a stopper was used

170 CHEMICAL TECHNIQUES

having a drawn-out piece of glass tubing to plug one end (Fig. 46)
so that the larger air space would prevent the displacement of the
liquid film, which was across the tube, when the stopper was fitted
on. To protect solutions from atmospheric carbon dioxide, Linder-
str0m-Lang, Weil, and Holter (1935) employed stoppers containing
soda lime tubes (Fig. 47).

Tube Holders. Perhaps the simplest holder for a small reaction
tube is a short length of thick- walled rubber tubing into which the
bottom of the tube may be placed, as in Figure 50. It is more con-
venient to use a small wooden or metal block with three flexible
metal prongs to hold the tube. For titration, where the color of the
solution is to be matched with a color standard, a single block with
prongs to hold two tubes is used (see Fig. 64, page 180; A. H. Thomas
Co. and E. Petersen, Carlsberg Laboratory).

Rediiclor. Kirk and Bentley (1936) devised a small glass volu-
metric flask of either 0.1 or 0.2 ml. capacity for use as a reductor
in their method for the estimation of iron (Fig. 48). The reductor
is made from heavy-walled 2 mm. bore capillary tubing. In the iron
method (page 277) cadmium amalgam is employed to reduce the
iron.

Tube and Pestle. For the grinding of bits of tissue, Glick ( 1937)
used a small pestle with a 250 /xl. tube having the inner bottom sur-
face ground as shown in Figure 49.

B. MICROLITER PIPETTES

Pipettes of various designs have been employed for measuring
microliter volumes. The chief among these will be described.

Fixed Pipettes. One of the pipettes developed by Linderstr0m-
Lang and Holter (1931) is shown in Figure 50 (A. H. Thomas Co.
and E. Petersen, Carlsberg Laboratory). It consists of a capillary
tube drawn out to a tip which is slightly bent so that contact can
be made with the wall of a vessel. The pipette is calibrated by first
weighing it, and then filling it with water to a little more than the
i-equired volume. The i)ipette is again placed on the pan of the
balance and water is removed by touching a piece of filter paper to
the tip. When the desired weight of water remains in the pipette, it
is removed from the balance and a mark is placed at the meniscus,
either by etching with hydrofluoric acid or using a piece of gummed

GENERAL APPARATUS AND MANIPULATION

171

Fig. 45. Reaction tube cap with hole.
From Linderstr0m-Lang and Holier (1933b)

Fig. 46. Stopper
for reaction tube with
cap of drawn-out glass
tubing. From Linder-
str0m-Lang and Holler
(1933b)

Ky

Ground
inner
surface S^^/

I

Ground
surface

Fig. 47.

Fig. 48.

Fig. 49.

Fig. 47. Soda lime tube for stoppering reaction vessel. From Linderslr0m-Lang ,

Weil, and Holler (1935)
Fig. 48. Reductor. actual size. From Kirk and Bentley (1936)
Fig. 49. Tube and pestle for grinding tissue. From GUck (1937)

172 CHEMICAL TECHNIQUES

paper. In the assembly shown in Figure 50, the pipette is filled by-
applying gentle suction through the tube S with H closed and K
open. When the liquid is a little above the mark, K is closed and the
slowly falling meniscus is observed through the low-power micro-
scope (M). The moment the meniscus reaches the mark, the vessel
of liquid is quickly lowered away from the tip and the capillary
forces will prevent the liquid from running out of the pipette. The
vessel into which the liquid is to be delivered is brought up so that
the pipette tip touches the vessel wall near the bottom and H is
opened. P leads to a source of compressed air, and the pressure
regulator ( T) enables the hquid to be forced out of the pipette under
constant pressure. Usually a 20 cm. column of water gives the
required pressure; the emptying time should not be less than 5 sec.
H is not to be closed until the delivered liquid has been lowered away
from the pipette. With pipettes having a capacity of 7 ix\. the error
of pipetting was found to be less than 0.3%.

Hand Pipettes. A hand pipette (A. H. Thomas Co. and E. Peter-
sen, Carlsberg Laboratory), Figure 51, having an accuracy of about
1 % was also used by the Carlsberg group. The instrument is filled or
emptied by sucking or blowing through the attached rubber tubing.
The tip of the pipette is fine enough to prevent liquid from running
out unless a slight pressure is applied through the rubber tubing.
Hand pipettes, in which the suction or pressure is applied by a glass
syringe, have been used by Kirk's group, Kirk and Craig (1932),
Sisco, Cunningham, and Kirk (1941) (Figure 52) {Microchemical
Specialties Co.). A rubber gasket fixed to the end of the syringe
barrel receives the large end of the pipette, or the metal syringe
fitting of a hypodermic needle is cemented to the pipette in order to
permit easy attachment to, and separation from, the syringe.

Constriction Pipettes. The preceding types of pipette have been
displaced very largely by the constriction pipette (Levy, 1936;
Linderstr0m-Lang and Holter, 1940) shown in Figure 53 (A. H.
Thomas Co. and E. Petersen, Carlsberg Laboratory) . In this pipette
the calibration mark is replaced by a constriction in the lumen of
the capillary. Liquid is first sucked up over the constriction and a
slight pressure is then applied which causes the liquid to fall down
to the constriction but not past it. To deliver the charge, a momen-
tary greater pressure is applied to force the meniscus through the
constriction and then gentle pressure can be employed to empty the

173

H

-r- -

"&

c=a=^

[M]

M

Fig. 50. Fixed pipette.
From Linderstr0ni-Lang and Holler (1931)

Fig. 52. Capillary pipette and syringe
control. From Sisco, Cunningham, and
Kirk (1941)

n

Fig. 51. Hand pipette.
From Linderstr0m-Lang
and Holler (1933a)

-o-

-n

=TJ

<]

Fig. 53. Constriction pipette:

(A) proper form; (B) faulty form.

From Linderslr0m-Lang and Holler (1940)

}

^^

Fig. 54. Constriction pipette

with water jacket.
From Holler and Doyle (1938)

174 CHEMICAL TECHNIQUES

pipette. If employed in the assembly shown in Figure 50, a quick
squeezing of the rubber tubing over K will be sufficient to initiate
the emptying process. Greater accuracy is obtained by adjusting the
dimensions so that the pipette delivers automatically without apply-
ing excess pressure w^hen the tip touches the vessel or the liquid.
The tip and the constriction should be constructed as in A (Fig. 53) ,
and not as in B. Holter and Doyle ( 1938) employed a constriction
pipette surrounded by a water jacket to control the temperature of
the liquid being pipetted (Fig. 54).

In the procedure for the determination of total nitrogen (pages
234 and 283) the pipettes used must meet certain dimensional re-
quirements as defined by Briicl et al. (1946). Thus, the pipette used
to transfer the digested sample must have a tip, the opening of
which is not so narrow as to become blocked by small crystals or
other particles; but neither must it be so wide as to make it difficult
to empty the pipette without blowing air through the tip, which
might cause spattering of the liquid delivered. Furthermore, the
pipette stem must be thin enough for use in a narrow tube without
causing the liquid to be drawn up between the tube and pipette, and
yet it must be thick enough to have mechanical strength. A suitable
pipette is illustrated in Figure 55, and the allowable variation in the
dimensions is shown in Figure 56, which gives the dimensions of a
rather thin and a rather thick pipette, either of which may be used.

The dimensions of a suitable constriction pipette for pipetting the
acid used to absorb ammonia are given in Figure 57.

Placing a water seal of known volume across the lumen of a reac-
tion tube is best performed with the type of constriction pipette
shown in Figure 58. Water is drawn up to the point X; the entire
amount is blown out to form the seal, and then the excess water is
sucked back into the pipette to Y. The amount left in the seal is
then the volume between X and Y in the pipette. Acid-selenium mix-
ture is pipetted with the horizontal pipette illustrated in Figure 59.
Each division corresponds to 1 //.I. These pipettes are available from
E. Petersen, Carlsberg Laboratory.

Automatic Pipettes. An automatic pipette was designed by
Linderstr0m-Lang and Holter (1931) and it was used by them for
the accurate delivery of 20—40 /xl. alcoholic acid to stop enzyme ac-
tion (Fig. 60) {A. H. Thomas Co. and E. Petersen, Carlsberg Labo-
ratory). The pipette consists of a narrow glass tube drawn out to a

175

0.4

1 1 1 1 ,

1.0

1

0.2

0

0.2

ry////////^

0.8
0.6

\'////////Z^.

0.4

1 0.6

-

io.2

<■

UJ 0.4

J UJ

D < 0.2

^^"

DIAMETER,
o

E

E

0

j^^_

0.4

o

0.2

\^^/7/77777y>^

0.6

0.4

_^^^^^^^^22^%

0.8

0.6

(

. , , , "

1 0

3 10 20 30 40
LENGTH, mm.

30

10 20 30 40 50 60 70 80
LENGTH, mm

Fig. 55.

Fig. 56.

Fig. 57.

Fig. 55. Pipette for transfer of digested sample in nitrogen determination.
From Brilel et al. (1946)

Fig. 56. Allowable variation in dimensions of pipette in Fig. 55. From
Brilel et al. (1946)

Fig. 57. Dimensions of constriction pipette for pipetting acid used to absorb
ammonia in nitrogen determination. From Brilel et al. (1946)

y

-X

■ 45 n\.

-Y

-A/il.

Fig. 59. Horizontal mouth-operated

pipette with vertical delivery tip.

From Brilel et al. (1946)

Inner diameter 0.35 mm.
Outer diameter 0.70 mm.

Fig. 58. Constriction pipette
for placing liquid seal across

neck of reaction tubes.

From Brilel et al. (1946)

Fig. 60. Auto-
matic pipette.
From Linderstr0m-
Lang and Holler
(1931)

176 CHEMICAL TECHNIQUES

capillary at both ends so that, with one atmosphere pressure, the
fineness of the tips will prevent liquid from running out. 22 is a
siphon arm connecting to a reservoir of the liquid to be pipetted
which is placed about 50 cm. above the instrument. The pipette is
filled as follows: Close L and open K and H to fill the outer cham-
ber. When the level is a few mm. over the upper tip of the pipette,
close H; the pressure from the reservoir will fill the pipette. Then
close K and open // and L to bring the level of the liquid in the outer
chamber below the upper tip. Deliver the pipette charge by closing
H and L and opening K, which compresses the air in the chamber and
forces the liquid out. A pipette of this type having a capacity of 30
^il. was found to have an error of measurement of less than 0.1%.

Accurate syringe pipettes have been employed which use a screw
(Krogh and Keys, 1931; Krogh, 1935) or a micrometer spindle
(Trevan, 1925) to move the plunger of a small hypodermic syringe.
The micrometer syringe pipettes are essentially the same as the
micrometer burettes of Dean and Fetcher (1942) and Hadfield
(1942) (page 255). The Krogh-Keys instrument is manufactured
by Macalister Bicknell Co. It has a delivery precision of 0.1 ix\.

Devices for Drawing Finer Pipettes. Finer pipettes which are
used under a microscope may be drawn by hand, but mechanical
devices for making them are considerably more efficient. DuBois
(1931) described an automatic device for drawing very fine micro-
pipettes and microneedles which has been made available commer-
cially (Leitz). A capillary tube is clamped in two arms of the device,
and between the arms the tube passes through a small electric heater.
When the glass softens in the heater the spring tension on the arms
pulls them back, thus drawing out the tube into a pair of pipettes.
Rachele's device, described by Benedetti-Pichler and Rachele ( 1940) ,
operates in a similar manner except that gravity is used to pull out
and lower the arms when the glass is softened by the electric heater.

C. FILTERS

Sintered-glass filters for small volumes of liquid were used by
Kirk and co-workers for the quantitative collection of precipitates
for various determinations. The filter described by Cunningham,
Kirk, and Brooks (1941b) is made of capillary tubing (2 mm. in-

GENERAL APPARATUS AND MANIPULATION

177

ternal diameter, 6 mm. external diameter) the end of which is
tapered and contains a fused-in 4 mm. section of sintered glass at
the tip. Kirk ( 1935) discussed the preparation of sintered-glass
filters for those who cannot obtain them commercially. A layer of
fine asbestos 1 mm. thick is sucked onto the tip of the filter to form
a pad; after pressing this pad down with the fingernail, a layer of
asbestos is deposited on the pad to form a cone about 2 mm. deep
The filter is used as shown in Figure 61. Only the tip of the cone ia
allowed to be in contact with the liquid in order that tlie precipitate
may be collected entirely on this part of the asbestos. The precipitate
can be transferred quantitatively by disengaging the pad at its base.
These filters are available from Microchemical Specialties Co.

Fig. 61. Filtration detail. A represents

an asbestos filtering cone; B, an asbestos

base pad; and C, a sintered-glass plug.

From Cunningham, Kirk, and

Brooks (1941b)

Fig. 62. Ultrafilter for small

volumes of liquids.
From Johnson and Kirk (19IiO)

Bott ( 1943) employed a capillary tube filter with paper pulp (Fig.
76) in the determination of sodium. A description of the preparation
of filter and its use is given on pages 204-207.

Ultrafilters. Various devices have been employed for the ultra-
filtration of small volumes of liquids, and only that of Johnson and
Kirk (1940), designed for about 0.1 ml., will be described, since
it is one of the simpler and more efficient types. The ultrafilter shown
in Figure 62 consists of two brass tubes, A and B, drilled to fit snugly
the glass capillary tubes having a bore of 2 mm. or less and an out-
side diameter of 7-8 mm. Kronig cement ( 1 part white wax and 4

178

CHEMICAL TECHNIQUES

parts rosin melted together) is used to bind the glass to the metal.
A brass collar, C, engages B and is threaded to A. For visibility, the
center part of the collar is cut away on two sides. The ends of the
capillary tubes are flared and ground fiat, and a piece of collodion
membrane is held tightly between the ground surfaces at D. The
ultrafilter is used with positive pressure.

D. STIRRING DEVICES

Many investigators have used a stream of carbon dioxide-free
air bubbles ejected from a fine glass tip to obtain stirring in manipu-
lations of a submacro order. However, when very small volumes of
liquid are to be stirred, recourse must be had to other means. When
dealing with a drop of liquid in a capillary tube, a practical method
of agitation appears to be the simple expedient of moving the drop
back and forth in the tube by means of controlled air pressure.
Schmidt-Nielsen (1942) devised a centrifuge for sealed capillary
tubes which rotates them so that liquid contained within will be
thrown from one end to the other to effect mixing. The apparatus,

, rt R ^

■ Ampullae

63. Centrifuge apparatus for mixing and extracting small amounts
of liquid in ampules. Length of the apparatus about 25 cm.
From Schmidt-Nielsen (1942)

shown diagrammatically in Figure 63, is attached to the motor
shaft and revolves with it. During the revolutions the plate with
the ampules is slowly turned, being connected by means of a
rubber band to a small rubber wheel which in turn is being driven
by its frictional contact with the motor housing. The rubber wheel

GENERAL APPARATUS AND MANIPULATION 179

is suspended in such a way that it is held against the motor housing
by means of the rubber band. The tubes are turned once for about
each five revohitions.

Agitation of a liquid in a capillary tube was effected by Bessey
et al. (]946) by touching the side of the tube to a rapidly rotating
nail head (page 250).

For stirring small volumes of liquid in an open shallow vessel,
Kirk ( 1933) employed the tip of a glass needle drawn from the end
of a tube in which a piece of iron was sealed. The core of an electric
buzzer placed in proximity to the iron made the glass needle vibrate
and stirring was thus produced. This type of stirrer is manufactured
by Micro chemical Specialties Co.

A particularly effective and convenient stirring device for small
volumes in open or closed vessels is the electromagnetic "flea" of
Linderstr0m-Lang and Holter ( 1931) . The "flea" consists of a sealed
glass spherical shell, about 1-2 mm. in diameter, filled with ferrum
reductum; stirring is effected by an electromagnet repeatedly turned
off and on by means of an interrupter. The core of the magnet is
placed near the outer wall of the vessel, and the lifting and dropping
of the "flea" provides the agitation. The arrangement employed in
titration is shown in Figure 64. The interrupter is not shown; it is a
small glass-enclosed mercury switch mounted on a pivot which is
connected to a movable strip of iron in the field of the magnet. When
the current is turned on, the magnetized core tips the iron strip,
which tilts the mercury switch and thus breaks the current. The iron
strip falls back when the core is no longer magnetized, and in so
doing it brings the mercury switch back to its original position,
which again completes the circuit, magnetizes the core, and starts the
process over again. "Fleas" are made by blowing a small bulb in the
end of a drawn-out piece of glass tubing, tapping a little ferrum
reductum down into the bulb, and sealing off the neck with a micro-
flame. The "fleas" may be cleaned by rinsing with water and cover-
ing them with fuming nitric acid. After washing well with distilled
water, the "fleas" are allowed to dry on a piece of filter paper. The
development of a brown stain of iron oxide on the filter paper under
a "flea" indicates that it has an incomplete seal and it should be
discarded. A method of testing the "fleas" suggested by Linderstr0m-
Lang and Holter ( 1940) is to place a drop of neutral bromothymol
blue solution on each one on a glass plate. Those not properly sealed

180

CHEMICAL TECHNIQUES

will become apparent since the acid that seeped into them during
the cleaning will cause the indicator to turn yellow. The complete
stirring equipment is available from A. H. Thomas Co. and E.
Petersen.

istiH_3> -=C_M}

nT

1

(qM§)

100

98

96

Fig. 64. Microtitration arrangement

for use with magnetic "flea" stirrer.

From Linderstr0m-Lang and Holier (1940)

Heatley, Berenblum, and Chain (1939) employed steel ball bear-
ings, ^/i6 in. in diameter, given several coats of Bakelite varnish No.
V-5209/2. Each coat was polymerized by stoving before applying the
next, and then the balls were given a layer of paraffin by heating
them to 100° in a paraffin bath. After excess paraffin was removed
by rolling the bearings on hot filter paper, they were rolled in the
clean, dry palm of the hand with some well-washed kaolin to enable
them to be wetted by aqueous solutions. Of course these balls should
not be used with liquids that might attack the coating.

E. HEATING DEVICES

A simple micro muffle furnace was described by Kirk and Bentley
( 1936) which, when employed with the proper rheostat, can be
used for temperatures up to 1000°. The furnace is made by winding
Chromel A resistance wire around a porcelain cup (2 in. inside di-

GENERAL APPARATUS AND MANIPULATION

181

ameter, 4 in. deep), embedding in insulating material, and surround-
ing with iron or brass pipe.

Linderstr0m-Lang (1936) employed an incineration oven for the
ashing of samples in small tubes. The oven (Fig. 65) is made of a
solid copper block containing holes 18 mm. deep and about 7 mm.
in diameter. Two electric heaters placed at the sides of the block
enable a temperature of 440-460° to be maintained. A rheostat is
used to obtain lower temperatures and to regulate the heating. The
sides and bottom of the oven are insulated with asbestos. The tubes
used with this oven were of quartz and had an inner diameter of
3.8 mm., an outer diameter of 6 mm., and a length of 20 mm. A
solid copper block with holes drilled to accept small tubes for
Kjeldahl digestions was used with gas heat by Borsook and Dubnoff
(1939), as shown in Figure 66.

Copp'

Fig. 65. Incineration oven

to accommodate small tubes.

From Linderstr0m-Lang (1936)

8 mm. H h-

Fig. 66. Digestion rack and Kjeldahl tubes.
From Borsook and Dubnoff (1939)

Naturally, modifications in the micro furnaces and ovens may be
made, and commercially available types such as that of Micro-
chemical Specialties Co. are also often suitable.

F. MOIST CHAMBERS

When working with small volumes of liquid it is necessary in
certain instances to maintain a moist atmosphere around the liquid
to prevent evaporation. The various forms of the moist chambers
employed on the stage of a microscope have been described by
Chambers and Kopac (1937). In general these are partially en-

182

CHEMICAL TECHNIQUES

closed chambers containing strips of wet filter paper, or a layer of
water on the floor.

Holter (1945) described a large moist chamber into which the
hands may be placed for various operations, and into the top of
which a binocular dissecting microscope is fitted to enable observa-
tion of the material being manipulated. The humidity is kept con-
stant through the regulation afforded by an electrically fitted hy-
grometer connected to a circulation pump which supplies an adjust-
able proportion of wet and dry air.

Holter Moist Chamber. The chamber is illustrated in Figure
67. It is made of varnished plywood ; the dimensions of the box are
64 X 35 X 30 cm., not considering the bevelled surfaces on the ends
of the front which contain doors 12 X 12 cm. The hands may be
inserted through tight-fitting rubber cuffs attached in the openings
of the doors. A slanting glass window (20 X 20 cm.) is fitted into
the front of the box and is hinged so that larger objects can be

f^\-^

Fig. 67. Air-conditioning chamber.
From Holter (1945)

Fig. 68. Arrangement of

air-conditioning apparatus.

From Holter (1945)

placed inside. The edges of a sheet of rubber are sealed into a hole
( 15 X 15 cm.) cut in the top of the box; holes in the rubber fit tightly
around the tubes of a binocular dissecting microscope. The sheet of
rubber is rather limp and bulging so that vertical movements of the
microscope will not cause it to stretch unduly. Illumination of the
interior is supplied through a window in the back wall which, for
some purposes, should be screened with a heat-absorbing device.
A moist atmosphere is maintained in the box by means of the

GENERAL APPARATUS AND MANIPULATION 183

arrangement shown in Figure 68. An electric circulation pump (P)
sends a current of air, divided by the T-tube (a), into the chamber.
The water in the large bottle (B) is warmed by a lamp (about 40
watt) to a temperature 5° higher than that of the room, and the
copper coil attached to the arm (6) is surrounded with water cooled
5° below the room temperature. The air passing into the water in
(B) is dispersed into fine bubbles. The currents of warm and cool
air enter the chamber at the same corner and the baffle (d) permits
them to mix without allowing water droplets to be carried into the
center of the box. The box contains a thermometer (/) and a hair
hygrometer (e) fitted with an electrical contact at the percentage of
moisture desired. By means of pinch cocks on tubes g and b, the
warm air stream is first regulated so that the temperature rise in
the chamber is about 1° in 15 min. when the cool air is shut off, and
then sufficient cool air is admitted to compensate for this tempera-
ture rise. The contact on the hygrometer is connected through a relay
to the pump which automatically stops when the desired humidity
is attained and starts when it begins to fall. A thermoelectric control
of the air flow through a or 6 could be used to achieve finer regula-
tion, but Holter found it unnecessary for his work.

G. ELECTRODES

Linderstr^m-Lang, Palmer, and Holter Silver Electrode. A

simple electrode arrangement (Fig. 69) was used by Linderstr0m-
Lang, Palmer, and Holter (1935) for the micro determination of
chloride by electrometric titration. The silver wire electrodes (A)
and (^4') are employed as shown. A is fixed with a bit of picein in
the side tube of the tip of the burette (B). Before sealing it in the
side tube the wire is cleaned with a little cold dilute nitric acid, and
afterward it is kept in contact with the acid silver nitrate titration
solution. In this manner it need not be changed for months. A^
requires occasional cleaning with a little nitric acid and, if neces-
sary, fine emery cloth may be used. After titration this electrode is
dried with filter paper, taking care not to touch it with the fingers.
The tip of the burette may be protected from contact with A' by
the glass cap shown on the right of Figure 69.

Sisco, Cunningham, and Kirk Glass Electrode. An open-cup
glass electrode was used by Sisco, Cunningham, and Kirk (1941)

184

CHEMICAL TECHNIQUES

for formol titrations in the manner shown in Figure 70. The cup is
made by blowing a bulb and then sucking in a depression. The

\^

Fig. 69.

Fig. 70.

Fig. 69. Silver electrodes (A and A') arranged for chloride titration with
burette (B) and electromagnet on the left to enable stirring with the "flea" in
the vessel (see page 282). Tip of burette may be protected from contact with
A' by the glass sleeve shown on the right. From Linderstrdm-Lang , Palmer,
and Holler (1935)

Fig. 70. Cross section of the glass electrode titration vessel. B represents an
inverted glass electrode, C a burette, D a reference calomel cell, E a glass
tube. A is a cup assembly, consisting of: a, a central block; b, a lower cup;
c, an upper inverted cup. From Sisco, Cunningham, and Kirk (1941)

outside of the cup is coated with paraffin and the electrode is filled
with 0.1 A'" hydrochloric acid saturated with quinhydrone. The
chamber (A) is made of Lucite. Stirring is effected by blowing a
streari of nitrogen through the tube (E) in such a fashion as to
whirl the sample drop in the cup.

Claff and Swenson Glass Capillary Electrode. Glass electrodes
employed for measurements of the hydrogen ion concentration of
small volumes of solutions have been numerous. One of the more

GENERAL APPARATUS AND MANIPULATION

185

Lucite wafer
(insulator)

Picein
plug

Lucite wafer
(insulator)

Heated glass
rod

Lucite wafer

Paraffin

Corning 015
glass capillary

Fig. 71. Capillary glass electrode assembly.
From Claff and Swenson (1944)

186 CHEMICAL TECHNIQUES

recent of these is tluit described by Claff and Swenson ( 1944) , which
can be used for voliunes as little as 5 ix\. and is capable of repro-
ducibility in measurement of ±0.02 pH units. The apparatus is
indicated in Figure 71. The glass electrode is a capillary tube of
Corning No. 015 glass 75 mm. long, attached to two Lucite wafers
with picein as shown. The capillary assembly is fitted in a jig so that
the wafers will always be a fixed distance apart. Starting 4 mm. from
each end, the capillary is brushed with hot paraffin up to the wafers.
One end of the capillary tube is plugged with picein and the open
end dips into a drop of the saturated potassium chloride solution
from the calomel half cell. The portion of the capillary tube between
the wafers is immersed in 0.1 A^ hydrochloric acid contained in a
flattened Pyrex funnel. The funnel is filled to capacity, surface
tension preventing the solution from overflowing. The stem on the
funnel is bent upward to form a silver-silver chloride electrode, and
the funnel unit is mounted on an insulated standard so that it can
be raised and lowered or moved horizontally. The entire assembly
is electrically shielded, and shielded leads connecting to the pH meter
are used. The capillary tubes are cleaned by sucking through them in
the following order: Keego cleaner {J. B. Ford Co.) 0.1 N hydro-
chloric acid, alcohol, distilled water, and, if blood is to be used,
0.2% potassium oxalate. The tubes are then dried by drawing air
through them. The tubes are stored in 0.1 N hydrochloric acid when
not in use.

Pickford Sealed-In Capillary Glass Electrode. A permanently
mounted capillary glass electrode was described by Pickford ( 1937) .
The capillary, made of Corning No. 015 glass, is sealed into the
apparatus as shown in Figure 72. The three-way stopcock is of the
type developed by Stadie, O'Brien, and Lang (1931). Of course it
must be made of the same kind of glass as that used in the electrode
jacket. A fine bore ( 1 mm. diameter) in the stopcock plug is required
for filling in order to keep the volume of the sample as small as
possible. The bore of the filling and connecting tube is about 0.5
mm. A 1 ml. syringe (preferably of the short insulin type) is em-
ployed to fill the electrode, using the assembly shown. The syringe
may be used to obtain the sample, in which case the needle would be
removed after the sample was taken. A short piece of hemocytom-
eter tubing (3 mm. inside and 5 mm. outside diameter) is then
slipped over the nozzle of the syringe and this is connected to the

GENERAL APPARATUS AND MANIPULATION

187

One way
stopcock

Bulb CaO. 1 n-^l -

Stdndard buffer
lOf 0 1 A' HCI/

Capillary glass
membrane

Ag-AgCI
electrode

To calomel

cell via

KCI flusti

To calomel

cell via

KCI flusfi

Diagonal bore

for filling,

diameter 1 mm

Three way
stopcock

Rubber
i.ubing

/

iMose ol Luer syringe

Fig. 72. Sealed-in capillary-
glass electrode assembly.
From Pickjord (1937)

D

O

O O

D

O
o o

B

D D

Fig. 73. Conductivity cell. The
upper section shows the faces of the
two blocks which make the cell (actual
size), the center section, the construc-
tion, and below is an enlarged dia-
gram of the recess which holds the
fluid. Fro7n Bayliss and Walker (1930)

cup attached to the filling inlet. The electrode is calibrated with
standard buffer, washed, and then dried with alcohol and ether.
After the sample has been introduced the lower stopcock is rotated
clockwise, first to flush saturated potassium chloride through the
groove and right- angle bore (the position shown in Fig. 72) and

188 CHEMICAL TECHNIQUES

then to make liquid junction between the sample and the calomel
half cell. After use the electrode is washed with dilute salt solution
and kept filled with distilled water. The outer jacket is filled with
0.1 A^ hydrochloric acid. Pickford used a Pliotron tube amplifier
with the apparatus, and the electrodes were made by Macalister
Bicknell Co.

H. CONDUCTIVITY APPARATUS

The Bayliss-Walker Cell. A conductivity cell was designed by
Bayliss and Walker (1930) for measurements on as little as 0.5
ix\. of liquid (Fig. 73). Two blocks of vulcanite (A, B) are shaped
and drilled as shown. Two small holes are drilled near the upper
edges and in one block an enlargement is made to form a recess (C)
0.5 mm. deep, 0.75 mm. wide, and 1 mm. long in the face of the
block. In each of the small holes platinum wire (0.3 mm. diameter)
is sealed with sealing wax, taking care that the end of the wire is
flush with the bottom of the recess in the one case and with the face
of the block in the other. Glass tubes {E) are sealed into the blocks
with sealing wax, as shown, and when filled with mercury enable
electrical contact to be established. The blocks faced to fit perfectly
against each other are located by two pins and held firmly together
by a thumb screw so that a small cavity having a platinum electrode
in each face is formed. A conical hole (D) is made in the face of the
block nearest the cavity. The recess is lined with sealing wax, which
has to be replaced from time to time as the surface deteriorates.
Each electrode is coated with platinum black by covering with a
drop of 2% platinum chloride, dipping a wire into the drop, applying
3 volts to the circuit, and reversing the polarity every 10 sec. for
2-3 min. It is necessary to keep the cell in distilled water, drying it
only before use, in order to prevent a drift in the resistance during
measurements. It is also necessary to reblack the electrodes every
week or so.

The cell is filled through the conical opening with a fine pipette,
using a low-power microscope to aid in the operation. The pipettes
are made of small-bore glass tubing drawn out for about 10 cm. to
a diameter of around 0.2 mm. or a little less. Each is cut at points
20-30 mm. from the beginning of the constriction where the diameter
begins to be uniform, and the center capillary is discarded. The ends

GENERAL APPARATUS AND MANIPULATION 189

of the pipettes thus formed are bent at right angles about 5 mm.
from the small end. Care must be taken that the ends are cleanly
and squarely cut. A mercury leveling bulb connected to the pipette
with rubber tubing may be used for filling and emptying. Small
air bubbles sometimes cling to the cell walls or to the electrodes when
the cell is filled. This difficulty is overcome as a rule by withdrawing
the fluid into the pipette and refilling the cell more slowly.

The circuit used by Bayliss and Walker was the standard Kohl-
rausch bridge fed from a 1000 cycle audiofrequency generator. The
null point was determined with head phones in the usual way. Be-
cause of the small size of the electrodes and the necessity of drying
them, the null point tends to be flat. A 0.01 /xF. condenser placed
across the standard resistance makes the null point sharp. The
practice is to adjust the resistance until minimum sound intensity
is obtained with the slide wire at midpoint, or until a sharp increase
in intensity occurs symmetrically on each side of it.

I. BALANCES

The development of the instrumentation for the weighing of very
small amounts has been thoroughly review-ed by Gorbach (1936).
The commercial balances, including the torsion balances of Roller
Smith Co., which are sensitive down to about 2 fig., require no
comment here. The quartz fiber balances, which are considerably
more sensitive, are particularly useful in histochemical work and
these will be considered in detail.*

Quartz Fiber Balance. Lowry (1941) designed a simple and
serviceable quartz fiber balance that can handle a maximum load
of 200-300 fig., and that has a sensitivity of about 0.03 fig. and a
reproducibility of 0.1 jug. The functioning of the instrument (Fig.
74) depends on the measurement of the bending of a horizontal
hollow quartz fiber when a weight is attached to its free end. The
fiber (A), about 20 cm. long, is drawn from narrow quartz tubing.
One end of the fiber is fused at B to a low tripod (C) made of 1-2
mm. quartz rod. Instead of fusing the fiber to a quartz tripod, the
end can be inserted into a short Pyrex sleeve (having a lumen just

* Since this writing a new quartz fiber balance has been described by Kirk
et al.; a "Cartesian-diver" balance has been announced by Zeuthen (Bibliog-
raphy Appendix, Refs. 39, 49, and 54.)

190

CHEMICAL TECHNIQUES

large enough to hold it) fused to a Pyrex tripod. This enables easier
replacement of fibers. The Pyrex sleeve should lean toward the
front of the instrument at an angle of a little less than 90° to the
plane of the tripod. The free end of the fiber (D) is bent into a tiny
V in a plane at right angles to the fiber axis. Without a load, the
free end of the fiber should be 12-15 cm. above the tripod. The
tripod is mounted inside a metal cylinder (E), a gallon tin can will
do or a smaller instrument can be made to fit into a smaller can. The
open front of the cylinder is fitted with a removable glass plate {F).

tz^

_^.w

•10 cm-

N

Fig. 74. Quartz-fiber balance.
From Lowry (194V

The tripod is fixed in place with DeKhotinsky cement, and the
cylinder is mounted rigidly on a heavy wooden block. A cathetometer
(Q) , reading to 0.01 mm., is used to observe the positions of an
arbitrary point on the fiber tip when weights are applied. Illumina-
tion of the interior of the cylinder can be enhanced by removing
the back end of the can and replacing it with a plate of glass.
Electrostatic shielding can be increased by lining the inside surfaces
of the glass plates with metal foil from the center of which strips
have been cut to act as windows. For less accurate measurements
without a cathetometer, a narrow ribbon of graph paper running
down the center of the front window can serve for the indication of
the displacement of the end of the fiber.

GENERAL APPARATIS AND MAXIPlLATIOISr 191

The samples to be weighed are held in quartz fiber hooks (G)
1 cm. long with loops at each end 2 mm. in diameter. These are
made from 3-4 cm. lengths of solid fiber weighing about 30 /xg./cm.
One end of a piece of the fiber is held in a small oxygen flame so
that the force of the flame bends the tip as it softens. By proper
manipulation a complete circle can be made. The weight is now
adjusted by clipping off the straight end with a scissors until the
desired deflection of the balance is obtained with it. When a number
of these hooks have been brought to the same weight within a few
mm. deflection, the second loop is made in each one. The hooks are
stored on a glass rack (H) which has a series of pegs (J) 0.5 mm.
in diameter projecting from the large horizontal tube at 3 cm.
intei-vals. The fine glass springs (K) prevent hooks from falling or
blowing off. The hooks are handled by a glass rod (L) about 1 mm.
in diameter drawn out at the end to 0.2 mm. A 5 mm. bend (M) is
made in the end of this rod to slip into the loop for transfer. During
attachment or removal of hooks at the end of the fiber, a straight rod
(N) is used to hold the fiber.

After the case has been closed for 1.5-2 min., readings may be
taken; successive observations have been found to agree to 0.03 mm.
One hook is kept as a standard weight. Calibration of the balance
is carried out by accurately pipetting 3-10 /xl. of standard salt
solution into the lower loop of a hook (if 1-2 fx\. of distilled water is
placed in the loop first, the transfer of the salt solution is easier),
drying the solution, and observing the deflection given by the known
weight of salt. Deflections given by various weights are plotted to
form a calibration curve.

In order to obtain the dry weight of microtome sections of tissue,
Lowry places a 3-5 (A. drop of water in the lower loop of a weighed
hook with a fine-tipped pipette, and then places the section in the
drop with a fine rod. Hooks with sections are put on the rack, dried
at 100° for 30 min., and reweighed. For the measurement of neutral
fat, the hooks with the dried sections may be kept in ethyl or
petroleum ether for 30 min., redried in the oven, and reweighed.

Quartz Torsion Balance. A quartz torsion balance was devised
by Lowry (1944) that has a capacity of 50-100 mg. and a sensi-
tivity of ±0.1 fig. The instrument is shown diagrammatically in
Figure 75. The beam (A) is a quartz tube 25 cm. long and about 1
mm. in diameter suspended between the horizontal quartz fibers (C) .

"-v.

192 CHEMICAL TECHNIQUES

The quartz stand (B) supports these fibers. Fine quartz loops in the
ends of the beam hold quartz hooks (E) from which the aluminum
foil pans (D) are suspended. The two arms of the beam need not
be of exactly the same length. The feet (G) of the standard are
sealed to the floor of a balance case with DeKhotinsky cement. It is
convenient to employ the usual mechanism in analytical balances
that lifts the beam when loads are added to, or removed from, the
pans. Electrostatic shielding is achieved by lining the inside of the
balance case with metal foil in which windows (H) are cut. In addi-
tion, the members of the balance are metalized by coating them with
a 5% solution of chloroplatinic acid in alcohol and, after drying,
heating with a "cool" flame to effect conversion to metaUic platinum.
Particular care is required to avoid overheating the fine quartz sus-
pensions. A strip of aluminum foil is used to ground the instrument
to the case.

Fig. 75. Quartz torsion balance.
From Lowry (1944)

After the case has been closed for at least 1 min., measurement of
the displacement of one end of the beam produced by the load placed
on a pan is made with a cathetometer (F) reading to ±:0.01 mm. In a
particular balance constructed by Lowry, a load of 10.8 ^ug. pro-
duced a displacement of 1.00 mm. The cathetometer may be focused
on any convenient landmark on one end of the beam. An illuminated
piece of white paper outside the opposite end of the balance case
furnishes a background that facilitates the measurement.

Calibration of the balance is carried out by cutting 5-10 cm. of
fine wire, weighing 1.5-2 mg., into ten nearly equal lengths, weigh-

GENERAL APPARATUS AND MANIPULATION 193

ing the ten pieces together on a microbalance, and then observing
the displacement given by each piece placed separately on a pan.
The sum of the individual displacements divided by the total weight
gives the sensitivity. The process must be repeated on the other pan
if both arms are to be calibrated. The balance will accommodate
larger weights if a tare is used as a counterbalance.

//. COLORIMETRIC TECHNIQUES

A. CAPILLARY TUBE TECHNIQUE

During the course of their classical investigations dealing with
the composition of glomerular urine, Richards and his group at the
University of Pennsylvania developed a simple and clever technique
of capillary tube colorimetry which enabled them to carry out
analyses on less than 1 fx\. liquid with an accuracy comparable to
that of macro procedures. The chief problem, as stated by Richards
et al. (1933), was "to introduce the minute amount of fluid to be
analyzed into a capillary tube without evaporation or contamination,
to dilute it quantitatively with water if necessary, to introduce into
the same capillary in quantitatively accurate proportions and with-
out mixing the one or more reagents required for production of color,
to effect mixture of the fluids in the capillary tube at a given moment,
and to compare the resulting color with those developed in standard
solutions treated simultaneously in identical or equivalent fashion."

The recent introduction of microcuvettes for the colorimetry of
small volumes of liquid in photoelectric apparatus (page 216) will
very largely displace capillary tube colorimetry because of the ob-
vious advantages of greater objectivity and accuracy of the analyses,
and the greater ease of manipulation in most cases. However, the
capillary tube technique and methods are included here because
there are instances in which the equipment for the cuvette methods
is not available, or the volumes to be handled are still too small to
permit the use of cuvettes, even of the micro variety. Furthermore,
some of the capillary tube methods might be adapted to cuvette
colorimetry when the equipment for the latter is available, and in
that case the assembly of the methodology of the former would also
be useful.

195

196 CAPILLARY TUBE COLORIMETRY

1. Apparatus

Capillary Tubes. For blood collections, plasma protein precipi-
tations, and for the making of pipettes, capillary tubing having an
outside diameter of 0.8 mm. and an inside diameter of 0.6-0.7 mm.
was employed by Richards et al. (1933). The capillaries in which
reactions were produced and colors developed were 0.5 mm. outside
diameter and 0.35 mm. inside. These smaller tubes must have very
uniform bores and hence it is necessary that they be drawn mechan-
ically.

Pipettes. The pipettes are drawn from the larger capillary tub-
ing. Their slender tips should have an outside diameter of about 50
[x; the over-all length should be about 10 cm. Liquid is drawn up and
expelled in the pipettes by means of an attached piece of rubber
tubing through which suction or pressure may be applied.

Microscope. A binocular microscope giving about fifteen fold
magnification with an optical field of about 1 cm. in diameter is
recommended. For the microscopic measurements a micrometer disc
is placed in one of the oculars or the disc is cemented to the glass
stage of the microscope. The disc should have a 10 mm. scale divided
in 0.1 mm. In order to reduce the chance of evaporation of fluids,
the glass stage, with the exception of the circle visible in the optical
field, is covered with wet filter paper.

Water Manipulator. For the introduction and movement of
columns of fluid in the capillary tubes, controllable suction or pres-
sure must be applied at one end. A small syringe having a piston 3
mm. in diameter moved by a micrometer screw serves this purpose.
The tip of the syringe is connected by rubber tubing with a short
glass or metal tube drawn out at one end to a tip small enough to
enter the capillary tube. The syringe, rubber tube, and tip are filled
with colored water, care being taken to exclude air bubbles, and
mounted on a level with the microscope stage. When water is forced
out of the tip into the capillary tube, a water seal is formed which
permits the movement of water into or out of the capillary. In this
fashion columns of liquid may be introduced into the capillary tube
from the other end and their movements can be easily controlled.

Other Accessories. A small centrifuge is required that will hold
the capillary tubes. A piece of unglazed milk glass (35 cm. X 35 cm.
X 4 mm.), two desk lamps fitted with 100 watt bulbs, and a sus-

APPARATUS AND MANIPULATIONS 19?

pended lamp equipped with a 150 watt bulb and Daylight glass
filter are also needed.

2. Manipulations

1. Connect a length of capillary tubing to the water manipulator,
and fix the tube on the stage of the microscope so that it is parallel
to, and lying on, the micrometer scale with its open end near the
edge of the optical field farthest from the manipulator.

2. Force water from the manipulator into the tube until half of
its length is filled.

3. Bring the tip of a pipette filled with the solution into the
optical field and insert it into the open end of the capillary tube.
Carefully blow the liquid out of the pipette, at the same time draw-
ing it into the tube by turning the piston screw of the water manipu-
lator. The volume of liquid introduced is determined by the length
of the column as measured on the micrometer scale visible through
the tube. When the appropriate amount of the solution has been
introduced move the column inward so that its distal meniscus is
near the center of the field.

4. In the same manner introduce columns of reagents, and, when
these have been added, break off the portion of the tube containing
all the columns (about 3-4 cm. long) and seal both ends quickly in
a minute gas flame. Set aside in a horizontal position. When breaking
off the tube, caution is required to avoid including any portion of
the tube which has been wetted with the manipulator water ( a dia-
mond point is useful for cutting the tubes at the proper place) and
when the ends are sealed, care must be exercised to avoid heating
adjacent liquid columns.

5. In order to mix the solutions, briefly centrifuge the sealed
tubes to bring the separated columns together, invert, and again
centrifuge. Then repeat the inversion and centrifugation. (This may
be simplified, see page 178.)

6. When it is necessary to heat the mixture, place the tubes in
a hot water bath.

7. Since it is essential that color comparisons be made with tubes
of the same diameter, the tubes to be compared should be obtained
from the same original length of uniform-bore tubing. When many
tubes are to be compared, the various original lengths of tubing re-
quired may not have the same diameters. In this case break a single

198 CAPILLARY TUBE COLORIMETRY

30 cm. length of unifonii tubing into 2 cm. pieces. Transfer the
colored solutions to these pieces by breaking off the sealed ends of
the tubes, inserting one end of a tube into small rubber tubing held
in the mouth, and placing the other end in contact with the piece
into which the solution is to be transferred. Apply gentle pressure to
effect the transfer and quickly seal the ends of the tube with plasti-
cine, taking care to avoid contact between the plasticine and the
solution in the tube. In order to prevent blowing the liquid from one
tube right through the other, the two tubes should be held in the
position of a wide-angled V during the transfer.

8. For the comparison of blue colors, place two desk lamps fitted
with 100 watt bulbs side by side about 6 in. over the milk glass plate.
The use of two lamps prevents shadows. For colors at the red end
of the spectrum suspend a 150 watt lamp provided with color filters
over the plate. Place the standard tubes, one at a time, beside the un-
known on the plate for comparison. It is sometimes helpful to cover
the tubes with a piece of white paper in which a rectangular window
has been cut so that the visible columns are of the same length.

note: In some cases it has been found that the intensity of the color
developed when minute quantities of test solution and reagent are mixed
in capillar}' tubes is not the same as when the liquids are mixed in the
same proportions in macro quantities. However, when this difference does
not exist, it is obviously less laborious to prepare the series of standard
color mixtures in macro volumes and transfer them to capillary tubes.

3. Methods

PREPARATION OF PROTEIN-FREE SUPERNATANTS

The following procedures were employed by Richards' group
( 1933) for frog plasma. The proportion of plasma to precipitating
reagent and the final dilution may be varied to suit the particular
kind of blood used. In general it is best to keep the dilution of plasma
as low as possible for capillary tube colorimetry.

Tungslic Acid Supernalanls

1. Collect the blood directly in one of the micropipettes. A few
grains of dry sodium oxalate may be placed in the pipette in advance.

METHODS 199

2. Seal off the larger end of the pipette in a minute gas flame
and centrifuge at once.

3. Cut the tube a little above the juncture of the cells and plasma,
let the plasma flow back from the cut end by gravity, and then seal
both ends of the tube taking care not to heat the plasma.

4. Attach a large capillary tube (0.6 mm. inside diameter) to
the water manipulator and fix it on the stage of the microscope.
Draw back 5 mm. from the end of a column of Vis N sulfuric acid
5.0 mm. long.

5. Introduce a 4 mm. column of plasma and add 10% sodium
tungstate to it until the column becomes 5 mm. long.

6. The two columns now in the tube are made to oscillate back
and forth several times by means of the water manipulator in order
to effect thorough mixing of the plasma and tungstate.

7. Break off the distal part of the tube and seal the ends. The
end nearest the acid is sealed last and held in the flame long enough
to make a small bulb.

8. Centrifuge the tube, bulb end down. Reverse and recentrifuge
at least six times for complete precipitation. The final centrifuga-
tion should be thorough and the material should be left in the nar-
row end of the tube.

9. Break the tube about 5 mm. above the surface of the fluid and
draw off the protein-free liquid into a pipette. Should a zone of hazi-
ness exist between the clear fluid and the precipitate, too much oxa-
late was used.

Trichloroacetic Acid Supernatants

1. For frog plasma phosphates a 4 mm. column of plasma is
placed in a larger capillary tube followed by 1 mm. of 90% trichloro-
acetic acid ( by weight ) . Seal the tube and centrifuge with the plasma
end down.

2. Immerse in hot water for a moment and centrifuge several
times inverting the tube each time.

3. Should the supernatant fluid be turbid, separate from the pre-
cipitate by cutting the tube, draw into another pipette, seal the large
end, and centrifuge at high speed.

4. Separate the sediment by cutting off the tube. The protein-
free liquid is ready for transfer to a mixing capillary for color devel-
opment.

200 CAPILLARY TUBE COLORIMETRY

Zinc Sulfate-Sodium Hydroxide Supernatanls

1. For frog plasma chlorides a 3.0 mm. column of plasma is
drawn in 1 cm. from the end of a 10-12 cm. capillary tube followed
by separate columns of 6.0 mm. of 0.1 N sodium hydroxide and 2.1
cm. of 0.64% zinc sulfate (ZnS04.7H20, freed from excess acid by
three recrystallizations from w^ater) .

2. Draw in the columns so that at least 2 cm. from the end of
the tube is empty, break off the portion of the tube containing the
liquids, and seal both ends in a flame.

3. Place in the centrifuge with the plasma uppermost and mix the
liquids by four centrifugations.

4. Immerse the tube for 30 sec. in water at 90-95° and centrifuge
once for 5 min.

5. Cut off the tube above the fluid and then cut off the part con-
taining the protein precipitate. The protein-free fluid is then ready
for analysis.

CHLORIDE

By the use of s?/m-diphenylcarbazide ( Cazeneuve reagent) West-
fall, Findley, and Richards ( 1934) increased the sensitivity of
Isaac's (1922) method for the determination of chloride and then
adapted it to capillary tube colorimetry. Their procedure allows
chloride determination in a fraction of a fA. of liquid containing 1 /xg
or less of sodium chloride with an average error of under 3.0%. The
principle of the method is that dry silver chromate will react with
chloride to precipitate silver chloride and leave the chromate ion,
which can be estimated by its yellow color. However, a much more
intense purple-red color will develop in the presence of diphenyl-
carbazide. For other methods see pages 224 and 281.

Westfall, Findley, and Richards Method for Chlorides

SPECIAL REAGENTS

Potassium Chromate Standards. Dissolve 3.321 g. of pure dry
potassium chromate (corresponding to 2.00 g. sodium chloride) in
1 1. distilled water. Dilute to prepare standard solns. in the range
10-70 milligram per cent sodium chloride at 2.5 milligram per
cent intervals.

CHLORIDE 201

Powdered Silver Chromate. Add slowly 200 ml. 5.5% potassium
chromate to 100 ml. boiling 10% silver nitrate soln. Add drops of
the chromate soln. until a slight excess is present as indicated by a
yellow color. Cool, wash the precipitate with water, and air-dry
on a Buchner funnel.

Diphenylcarbazide Reagent. Dissolve 0.5 g. syw-diphenylcarba-
zide {Eastman Kodak Co.) in 70 ml. 95% alcohol; add 25 ml.
glacial acetic acid, and make up to 100 ml. with distilled water.
This reagent is stable at 20° for only 3 hr.

PROCEDURE

1. Since it will be necessary to measure columns of liquid longer
than the diameter of the optical field of the microscope, mount a 15
cm. steel rule, graduated in 0.5 mm., on the microscope stage.

2. Fill a 10-12 cm. capillary tube (0.35 mm. inside diameter) to
nearly half its length with water from the water manipulator, and
adjust the tube so that its open end is in the optical field over the
zero mark of the stage micrometer and adjacent to the zero of the
steel rule.

3. Introduce a 2-3 mm. column of zinc sulfate-sodium hydroxide
supernatant (page 200) or other fluid to be analyzed and measure its
length accurately, then introduce just nine times as much distilled
water. If the concentration of the unknown corresponds to less than
0.1% sodium chloride, less water will be required; if higher than
0.7%, more water must be used.

4. Draw the liquid in 2 cm. from the open end, break off the
portion of the tube containing the added liquids, seal the ends in a
flame, and mix well by eight brief centrifugations, reversing the tube
after each one.

5. Seal one end of a larger capillary tube (0.6 mm. inside diam-
eter), place a few grains of dry silver chromate in it and tap the
tube to get the material down to the closed end. Take care that none
of the substance is left near the open end; cut off a little of the tube
if necessary.

6. Cut off the end of the first (smaller) tube above the column
of liquid, insert the open end into the silver chromate tube so that it
projects into it for about 1 cm. and fasten the two tubes together
with a ring of DeKhotinsky cement.

7. Centrifuge with the larger tube down for a moment so that

202 CAPILLARY TUHE COLORIMETRY

the liquid in the small tube is forced into contact with the silver
chromate in the larger tube.

8. Warm the cement, withdraw and discard the smaller tube, cut
away any of the larger tube to which the cement is adhering, and
seal the open end in a flame.

9. Drive the silver chromate back and forth through the liquid
by eight successive centrifugations. Continue the last centrifugation
for 5 min.

10. Make a pipette from the smaller capillary tubing and draw
the supernatant fluid into it.

11. Seal the larger end of the pipette, and force the liquid into
this end by centrifuging for 5 min.

12. Examine the tube under the microscope (magnification 50X)
to be sure the fluid is free of silver chromate particles. If not,
transfer to another pipette and centrifuge again.

13. Place a new piece of the smaller tubing 10-20 cm. on the
microscope stage with one end in the optical field and the other con-
nected to the water manipulator.

14. Introduce a 2.0 mm. column of the chromate liquid obtained
in step 12, and after measuring its length accurately draw it in at
least 2 cm. from the end of the tube.

15. Introduce a column of the diphenylcarbazide reagent four-
teen times the length of the chromate fluid, draw both columns in
2 cm. from the end, break off the portion of the tube containing the
liquid, seal the ends, and place in the centrifuge with the chromate
fluid uppermost.

16. Measure 4.2 ml. diphenylcarbazide reagent into each of three
test tubes. Start the centrifuge containing the capillary tube, and
as quickly as possible measure into the test tubes 0.3 ml. of each of
three standard chromate solns. covering the range of concentration
within which the unknown lies.

17. Centrifuge the capillary tube eight times, inverting it after
each centrifugation.

18. Fill a piece of capillary tubing at least 3 cm. long, having the
same diameter as that containing the unknown, from each test tube
and seal the ends with plasticine.

19. Compare the colors of the unknown and standards on a milk
glass plate under Daylite electric bulbs.

20. The result obtained gives a first approximation of the chloride

CHLORIDE AND SODIUM 203

concentration of the unknown. It may be necessary to repeat once or
twice with fresh portions of chromate supernatant in order that the
final color comparison may be made to standards differing from one
another by the equivalent of 1.25 milligram per cent sodium chlo-
ride. Smaller differences must be estimated.

SODIUM

A method for the determination of sodium in samples containing
as little as 0.3 fig., e.g., 0.2 /A. urine, with an average error of about
3%, was described by Bott (1943). Since this method employs a 6
ml. volume for the development of color, it is obvious that smaller
quantities might be measured if the colorimetry were performed on
smaller volumes. The method depends on the precipitation of the
sodium as sodium zinc uranium acetate, and the measurement of the
zinc in a solution of the salt by means of the red color it produces
with diphenylthiocarbazone. This principle was used by Deckert
( 1935) for the determination of zinc, and the technique of Bott
could be adapted to the measurement of small quantities of zinc.
For other methods see pages 265 and 270.

Bott Method for Sodium

SPECIAL REAGENTS

Water. The water employed in preparing all of the reagents is re-
distilled from an all-Pyrex still.

20% Trichlorocetic Acid. Made from acid that has been redistilled
from an all-Pyrex still.

93% Alcohol.

Ether. Redistilled.

0.01 N Sodium Hydroxide. Carbonate free.

Zinc Uranium Acetate Reagent. According to Butler and Tuthill
(1931): Prepare a soln. of 80 g. sodium-free uranium acetate,
U02(C2H302)2.2H20, and 48 g. or 46 ml. 30% (by vol.) acetic
acid in water to make a total of 520 g. Prepare a second soln. of
220 g. zinc acetate, Zn(C2H302)2-2H20, and 24 g. or 23 ml. of the
30% acetic acid in water to make a total of 520 g. Cover and warm
both solns. on a steam bath until, with stirring, soln. is complete.
Mix while hot, let stand 24 hr., and if no yellow precipitate ap-
pears add 0.2 g. precipitated uranyl zinc sodium acetate in order

204 CAPILLARY TUBE COLORIMETRY

to saturate the soln. Shake well and filter through quantitative
paper before using.

Magnesium Uranium Acetate Reagent. According to Blanchetiere
( 1923) : Dissolve 100 g. uranium acetate in 60 g. glacial acetic
acid and enough water to make 1 1. Dissolve 333 g. of magnesium
acetate in 60 g. glacial acetic acid and enough water to make 1 1.
Combine equal vol. of the two solns. Filter the reagent through
quantitative filter paper before use.

Diphenylthiocarbazone solution. Prepare immediately before use
by shaking 100 mg. of the compound {Eastman Kodak) in 5 ml.
of the sodium hydroxide soln. for 3 min. in a glass-stoppered ves-
sel the ground surfaces of which are thinly coated with paraffin.
Filter off the excess reagent on quantitative filter paper which
has been washed in redistilled water and dried before use. Dilute
1 vol. of the filtrate with 4 vol. of the sodium hydroxide soln. A
considerable variation in the quality of the compound from one
lot to the next has been observed.

Zinc Standards. Prepare pure sodium zinc uranium acetate by pre-
cipitating the sodium of pure sodium chloride with the zinc ura-
nium acetate reagent. Dissolve 0.235 g. of the triple salt in redis-
tilled water and make up to 1 1. This stock soln. contains 1 mg. zinc
per 100 ml. and it will keep for years in a Pyrex bottle in the dark.
Frequently prepare dilute standards containing from 10 to 70
microgram per cent of zinc by diluting the stock soln. with re-
distilled water.

PREPARATION OF FILTERS

1. Cut 3 cm. lengths of capillary tubing, 0.6 mm. internal di-
ameter. Cut ends squarely or the funnel openings to be made later
will be off center.

2. Partially seal one end of each piece of tubing by twirling in a
microflame. Use a microscope to observe the result. The opening
should be funnel shaped, about 0.1 mm. at the top and less at the
bottom (Fig. 76).

3. Prepare paper pulp by teasing apart the filter paper in re-
distilled water and drying at 105°. Bits teased off the dried pulp are
placed in the filter tubes and pushed down into the funnel end by a
thin capillary tube about 6 cm. long sealed at one end. Pack each bit

SODIUM

205

of pulp separately using, alternately, the open and sealed ends of
the thin capillary tube in order to obtain a filter well packed on the
sides and in the center. Make the packed filters 0.6-0.8 mm. thick,
and see that no spaces are present around the filter mat.

Fig. 76. Apparatus for determination of sodium. A and C are
approximately actual size. B is an enlargement (X20) of the
end of a filter tube. From Bott (1943)

4. Wash and test the filters by filling the tubes, with the aid of a
syringe and adapter, with redistilled water. Place them in a round-
bottomed centrifuge tube fitted with a mat of clean dry filter paper
on the bottom, centrifuge for about 1 min., examine the tubes, and
discard any which have not drained completely. Dry the filter tubes
at 105° in a clean vessel and store in covered weighing bottles kept
in a dust-free container.

206 CAPILLARY TUBE COLORIMETRY

PROCEDURE

1. As in the procedure for chloride (page 201), mount a 15 cm.
length of capillary tubing (0.35 mm. inside diameter, and check the
outside diameter with a stage micrometer — it should be just 0.5 mm.)
on the microscope stage and fix a 15 cm. steel rule beside it so that
the zero on the rule is opposite the 35 mark on the micrometer scale.

2. To one end of the tubing attach a water manipulator and place
the other end over the 30 or 40 mark of the stage micrometer so that
a 0.2-0.4 fA. sample will be in the center of the optical field.

3. Introduce a 2-4 mm. column of sample and draw it into the
tube just far enough to give a fully curved meniscus. Measure the
length of the sample column and pull it in about 5 mm.

4. Depending on the size of the sample, introduce rapidly from
a rather coarse capillary pipette, just filled with freshly filtered re-
agent, a 30-40 mm. column of zinc uranium acetate reagent. Meas-
urement of the reagent column need not be precise but do not move
the column back and forth, since evaporation of the liquid will give
high results. Draw the column in about 10 mm.

5. Cut off the portion of the tube containing the liquids, seal both
ends in a microflame without heating the liquids, centrifuge and in-
vert the tube ten times, allow to stand at room temperature for 10
min., and then centrifuge and invert five times more. During the 10
min. intervals examine the filter under the microscope and repack
it gently.

6. Examine the empty end of the precipitation capillary to make
sure no crystals have remained there. Cut off the end of the tube
containing the precipitate and insert it into the filter tube so that
the open end is about 6 mm. above the filter mat. Seal the two tubes
together with DeKhotinsky cement as shown in Figure 76, taking
care to avoid heating the reagent. Insert the tubes in a small hole in
a rubber stopper and fit into a test tube as in A of Figure 76.

7. Lower the assembly into a centrifuge cup by means of rubber-
tipped forceps, and spin rapidly for about 15 sec. Examine the capil-
lary and filter; usually a little liquid is found above the filter. Cut
off the sealed end of the capillary without disturbing the assembly,
centrifuge again for about 6 sec, and again inspect. No fluid should
appear above the mat. Centrifuge for 2 min. more to insure com-
plete draining.

SODIUM 207

8. Set the assembly in a wooden block. Dip the end of a clean
microfimnel (about 15 /A. capacity) such as pictured at the top of C
in Figure 76 into the magnesium uranium acetate soln. Fill the funnel
completely by pinching and releasing the attached rubber tubing.
Wipe off the outside, and slip the rubber tubing over the end of the
small capillary. Centrifuge for 6 sec, take off the funnel, and re-
centrifuge for 30 sec. Now repeat the process with two funnel fillings
of alcohol and two of ether. After evaporation of the ether, clean,
dry, yellow crystals should be seen on the filter mat.

9. Transfer the stopper and capillaries to a larger tube calibrated
to contain exactly 6 ml. (C, Fig. 76). Introduce a little redistilled
water into the small capillary, remove the funnel, and centrifuge for
a few sec.

10. Cut off the capillary about 1 cm. above the DeKhotinsky
cement, and proceed as before after attaching the funnel, filling the
funnel with water six to seven times. This should dissolve the pre-
cipitate and transfer the liquid completely into the tube which is
then filled with water to the 6 ml. mark.

11. From a 0.1 or 0.2 ml. Mohr pipette drawn out to a fine tip,
add 0.1 ml. diphenylthiocarbazone soln. to each unknown tube, and
also to each of three colorimeter tubes containing 6 ml. of water and
of two standards, respectively. Mix the contents of all the tubes and
transfer the unknowns to colorimeter tubes. The blanks should be
golden yellow, and the solns. with 10-70 microgram per cent of zinc
should vary from orange to cherry-red. Make colorimetric measure-
ments immediately. With the Evelyn colorimeter use Filter 565. The
calibration curve is linear, but at least one standard should be run
every time an unknown is run since variations may occur even
though the reagent is prepared the same way every time. The blank
should be found to be negligible, since measurements carried out on
redistilled water substituted for the sample gave maximum values of
only ±0.6 microgram per cent of zinc.

12. Since the inside of the very curved meniscus is used for
measurement of samples, make a vol. correction by the addition of
0.005 fA. for each meniscus. From the corrected vol., calculate the
final dilution (in a corrected sample of 0.2 lA. vol. the dilution is
30,000 times), and apply the following relation:

Na concn. _ 23.00 Zn concn. in ^ilnfi-nn
in sample " QdM ^ final soln. ^ a^ution

208 CAPILLARY TUBE COLORIMETRY

PHOSPHATE

A colorimetric capillary tube method for inorganic phosphate was
developed by Walker ( 1933) as an adaptation of the Kuttner ( 1927,
1930) modification of the Bell-Doisy phosphomolybdic acid method.
Walker's procedure enables analysis of as little as 0.08 fA. of liquid
containing less than 1 m^tig. phosphate phosphorus with a mean error
of about +0.1% and a mean deviation of about ±2.5% for solu-
tions of known concentration. A recent discussion by Sumner (1944)
of phosphomolybdic acid methods should be consulted. For other
methods see pages 124, 226, and 280.

Walker Method for Phosphate

SPECIAL REAGENTS

Standard Phosphate Solution. Prepare standards in the range of
1.5 to 7.0 milligram per cent phosphorus differing from one an-
other by 0.5 milligram per cent and below 1.5 milligram per cent
by 0.1 or 0.2 milligram per cent.

Molybdic-Sulfuric Acid Reagent. To 1 vol. of 10 A^ sulfuric acid
(282 ml. cone, acid, 95%, sp.gr. 1.84, to 1 1.) add 2 vol. distilled
water and 1 vol. 7.5% sodium molybdate soln. Store in a brown
glass-stoppered bottle.

Stannous Chloride Stock Solution. Prepare a 40% soln. in cone,
hydrochloric acid and store in a brown glass-stoppered bottle. Do
not use longer than one week.

Stannous Chloride Working Solution. Dilute the stock soln. 1:100
and prepare fresh each day.

PROCEDURE

1. Introduce a column of about 0.2 /xl. of the trichloroacetic acid
supernatant (page 199) or other phosphate soln. into a capillary tube
(0.35 mm. inside diameter) followed by an equal vol. of the molyb-
dic-sulfuric acid reagent and withdraw the two columns 3 cm. from
the end of the tube.

2. Introduce a third equal column of the stannous chloride work-
ing soln. and seal the ends of the portion of the tube containing the
liquids.

3. In a similar manner prepare tubes with the standard phos-
phate solns.

PHOSPHATE AND PHOSPHATASE 209

4. Centrifuge all the tubes at one time with the stannous chloride
soln. up.

5. Compare the colors on a milk glass background illuminated by-
two lamps arranged to avoid shadows. The colors fade by about
10% during the first few min. but this change occurs equally in all
of the tubes and hence need not interfere with the comparisons.

PHOSPHATASE

By an adaptation of the method of King (1932) to capillary tube
colorimetry, Weil and Russell (1940) worked out a procedure for
the determination of phosphatase which could be applied to less
than 1 ix\. plasma with an average deviation of ±3.0%. Their
technique employs capillary tube procedures for the determination
of the inorganic phosphate, but the enzymatic digestion procedure
is based on the use of pipettes, reaction tubes, and other apparatus
common to the titrimetric techniques. As in the method of Siwe
(1935b) (page 226), aminonaphtholsulfonic acid is used to reduce
the phosphomolybdate. For other methods see page 226,

Weil and Russell Method for Phosphatase

SPECIAL REAGENTS

Standard Phosphate Solutions. Prepare standards in the range of
0.02-1.00 /xg. phosphorus/15 /A. differing from one another by
0.02 or 0.04 ixg.

Molybdic-Sulfuric Acid Reagent. 5% ammonium molybdate con-
taining 15% by vol. cone, sulfuric acid.

Ajninonaphtholsulfonic Acid Solution. Dissolve 0.5 g. of the 1,2,4
acid, 30 g. sodium bisulfite, and 6 g. crystalline sodium sulfite in
water by shaking, and make up to 250 ml. Filter and, if filtrate
is not clear, leave overnight and again filter. Prepare fresh
every 2 weeks.

Veronal Buffer, pH 9.0, with magnesium. 0.0015 M magnesium
chloride in buffer consisting of 9.36 ml. 0.1 M sodium diethyl
barbiturate + 0.64 ml. 0.1 N hydrochloric acid.

Substrate Solution. 0.1 M sodium-/3-glycerophosphate.

10% Trichloroacetic Acid.

210 CAPILLARY TUBE COLORIMETRY

PROCEDURE

1. Pipette 3 fA. plasma into 21 /xl. water in a reaction tube of
250 /xl. capacity, and add 7 /xl. Veronal buffer with magnesium and
7 /xl. substrate soln. Mix with a magnetic stirring "flea" (page 179).

2. Set up a control experiment in which the substrate and buffer
are placed as a separate drop on the side of the tube where it
cannot touch the enzyme soln.

3. Place tubes in a rack in a desiccator containing water in the
bottom and a small bottle of chloroform to produce a vapor inhibit-
ing growth of microorganisms. The desiccator is kept at 37° and
the digestion is allowed to proceed for 4 hr.

4. Stop the reaction by setting the tubes in ice water, and add
10 /xl. 10% trichloroacetic acid to each.

5. Centrifuge, and pipette 15 /xl. of the supernatant into another
tube.

6. Add 7 /xl. of the molybdic-sulfuric acid reagent to the super-
natant.

7. Pipette 5 /xl. aminonaphtholsulfonic acid soln. on the side of
the tube as a separate drop.

8. Set up standards with 15 /xl. of the known phosphate solns.,
following steps 5-7.

9. Mix the drops on the side of the tubes with the rest of the
liquid using stirring "fleas."

10. Draw the solns. into capillary tubes of uniform lumen (inside
diameter 0.65 mm., length 30 mm.) and seal the ends with Duco
cement.

11. Compare the colors as in the Walker method.

REDUCING SUBSTANCES

Sumner's ( 1925) dinitrosalicylic acid method was adapted to
capillary tube colorimetry by Walker and Reisinger ( 1933) . In
this manner, quantities of glucose of the order of 0.1 /xg. in 0.2 /xl.
liquid (50 milligram per cent) can be determined with a maximum
error of 3 milligram per cent in duplicate measurements of solutions
of known concentration. For other methods see page 296.

REDUCING SUBSTANCES AND CREATININE 211

Walker and Reisinger Method for Reducing Substances

SPECIAL REAGENTS

Standard Glucose Solutions. Prepare solns. in the range 10-100
mg./lOO ml. in 5 mg. steps.

Sumner Reagent. Add 22 ml. of 10% sodium hydroxide to 10 g.
crystallized phenol. Dissolve in a little water and dilute to 100 ml.
Add 69 ml. of this soln. to 6.9 g. sodium bisulfite; then add a
soln. containing 300 ml. 4.5% sodium hydroxide, 255 g. Rochelle
salt (KNaC4H40r,.4H20) and 880 ml. 1% dinitrosalicylic acid.
Store in well-stoppered bottles and prepare fresh each week.

PROCEDURE

1. Introduce a 1.5-3.0 mm. column of tungstic acid supernatant
(page 198) or other unknown soln. into a capillary tube (0.35 mm.
inner diameter) followed by a second column (three times as long)
of the reagent. Seal both ends of the tube and mix by centrifuging.

2. Mix the standard solns. with reagent in test tubes employing
1 ml. glucose to 3 ml. reagent.

3. Immerse the capillary tubes and the test tubes together in
boiling water for 5 min.

4. Transfer the standard color solns. to capillary tubes.

5. Compare the colors on a white background under light
screened with Daylite glass.

CREATININE

Bordley, Hendrix, and Richards (1933) adapted Folin's method
for the determination of creatinine to capillary tube colorimetry.
As finally worked out, this adaptation enables analysis of about 0.5
fj}. of liquid containing 10-30 m/y.g. creatinine with an error of a few
per cent. For other methods see page 239.

Method of Bordley et al. for Creatinine

SPECIAL REAGENTS

Standard Creatinine Solutions. Prepare solns. containing 2.0, 2.5,
3.0, 3.5, 4.0, 4.5, 5.0, and 6.0 milligram per cent creatinine in 0.01
A^ hydrochloric acid. Add toluene as a preservative.

2 12 CAPILLARY TUBE COLORlMETftY

Saturated Picric Acid Solution. Prepare from pure picric acid.
10% Sodium Hydroxide. Prepare from Merck's reagent "from

sodium."
Folin Reagent. Freshly prepare before use by mixing 5 vol.

saturated picric acid with 1 vol. of the 10% sodium hydroxide.

PROCEDURE

1. Introduce separate columns of saturated picric acid (25 mi-
crometer divisions), tungstic acid supernatant (page 198) or other
unknown soln. (60 divisions), and 10% sodium hydroxide (5 divi-
sions) in that order into a capillary tube (0.35 mm. inside diameter) .

2. Seal off the ends of the tube and place in a closed box until
the other tubes are prepared.

3. Darken the room for the following operations.

4. Prepare as rapidly as possible the standard color solns. in
test tubes by adding 1 ml. Folin reagent to 2 ml. standard soln.

5. With no loss of time mix the liquids in the capillary tubes
by repeated centrifugations and plan to begin color comparisons
10 min. after the first centrifugation.

6. In this 10 min. interval transfer the contents of each capillary
tube and a portion of each standard mixture to pieces of capillary
tubing of uniform bore, and seal the ends of each piece with
plasticine.

7. Place each sealed piece of tubing in a labeled space on a
milk glass plate for color comparison.

8. Compare the colors in a dark room under a 200 watt bulb
equipped with a straw-colored light filter, or illuminate the milk
glass plate from underneath using a straw-colored filter between the
plate and the light source.

note: The particular order of procedure given must be followed since
Folin reagent darkens at a faster rate and more extensively in capillary
tubes than it does in larger volumes in test tubes. Furthermore, this change
is mtensified and accelerated by daylight, which makes it imperative to
protect the solutions from light. The yellow picric acid color interferes
with the comparisons of the red colors developed, and hence it is necessary
to use a straw-colored light filter. The color produced by 2.0 milligram
per cent creatinine is about the palest which can be reliably estimated in
the tubes used. The most advantageous colors are those produced in the
range 2.5 to 5.0 milligram per cent.

XJEIC ACID 213

URIC ACID

Bordley and Richards ( 1933) adapted Folin's ( 1930) method for
the determination of uric acid to capillary tube colorimetry with
the result that 0.03-0.5 fil. liquid containing 3-10 m/xg. uric acid can
be determined in solutions of known concentration with an average
error of about 5%. For other methods see page 239.

Bordley and Richards Method for Uric Acid

SPECIAL REAGENTS

Standard Uric Acid Solutions. Prepare stock soln. containing 1
mg./ml. Transfer 1 g. uric acid to a 1 1. volumetric flask. Shake
0.6 g. lithium carbonate in 150 cc. water for 5 min. to dissolve,
and filter. Heat the soln. to 60°, pour into the liter flask with
the uric acid, and shake for 5 min. Cool under running cold tap
water. Add 20 ml. 40% formalin and half fill the flask with
distilled water. Add a few drops of methyl orange soln. followed
by 25 ml. 1 N sulfuric acid added slowly and with shaking. The
soln. should turn pink before the last 2-3 ml. acid is added.
Dilute to vol., mix well, and store in a tightly stoppered bottle
protected from light. Prepare a series of standards containing 0.6,
0.8, 1.0, 1.2, 1.4, 1.6, and 2.0 mg./lOO ml.

Cyanide-Urea Solution. Dissolve 50 g. sodium cyanide in 700 ml.
water, add 300 g. pure urea and, when dissolved, transfer into a
2 1. flask. Add 5-6 g. calcium oxide and shake for 4-5 min. Filter
through Whatman No. 41 or similar paper. Add up to 1 g. finely
divided disodium phosphate; shake and filter. The soln. may be
used for at least 2 months if stored at room temperature and much
longer if kept cold.

Uric Acid Reagent. Dissolve 100 g. sodium tungstate in 200 ml.
water. Add slowly with stirring and cooling 20 ml. 85% phosphoric
acid. Pass a slow stream of hydrogen sulfide through the soln.
for 20 min. but after the first 3-4 min. add 10 ml. more of the
85% phosphoric acid. Filter through Whatman No. 41 or similar
paper, refiltering the first 40 ml. Transfer the filtrate to a separa-
tory funnel and shake for a few min. with 300 ml. alcohol. Transfer
the lower layer into a previously weighted 500 ml. flask and add
water to a total of 300 g. liquid. Boil a few min. to remove the

214 CAPILLARY TUBE COLORIMETRY

hydrogen sulfide. Add 20 ml. 85% phosphoric acid and slowly
boil for 1 hr. under a reflux condenser. Decolorize with a few drops
of bromine, boil off the excess bromine, and cool. To 12 g. lithium
carbonate and 25 ml. phosphoric acid, add slowly 150 ml. water.
Boil off the carbon dioxide and when solution is complete, cool
and mix with the cone, uric acid reagent and dilute to 1 1. Keep
in well-stoppered bottles protected from light.

PROCEDURE

1. Introduce into a uniform capillary tube (0.35 mm. inner
diameter) a 5 mm. column of tungstic acid supernatant (page 198)
or other soln. to be analyzed, 5 mm. cyanide soln., and 1 mm. uric
acid reagent, keeping the three columns separated by air spaces. Seal
off both ends of the portion of the tube containing the liquids.

2. In a similar manner prepare tubes with the seven standard
solns.

3. Mix the liquids simultaneously in all of the tubes by centri-
fugation.

4. Four min. after the mixing immerse the tubes for 1 min. in
boiling water.

5. Compare the colors.

UREA

Walker and Hudson ( 1937) adapted the capillary tube apparatus
to the determination of urea by the hypobromite method. This
adaptation enables the analysis of 0.3 /xl. liquid containing 2 to 25
milligram per cent urea nitrogen with an average deviation from the
macro method of ±3.8%. The measurement of known quantities of
urea added to dialyzed horse serum could be made with an average
error of 2.3%. For other methods see page 286.

Walker and Hudson Method for Urea

SPECIAL REAGENTS

Sodium Hypobromite Solution. Prepare according to Stehle
( 1921) : In a 50 ml. Erlenmeyer flask, mix 2 ml. of a soln. contain-
ing 12.5 g. sodium bromide and 12.5 g. bromine/100 ml., with 2 ml.
of a soln. containing 28 g. sodium hydroxide/100 ml. Gently

UREA 215

revolve the mixture in -the flask for 1 min. and set aside for 30
min. before use.

PROCEDURE

1. Fix a 15 cm. length of uniform capillary tubing (0.35 mm.
inner diameter) to the stage of the binocular microscope so that one
end is in the optical field and attach the water manipulator to the
other end.

2. Introduce a 6 mm. column of distilled water with a capillary-
pipette and draw it back 1 mm. from the end.

3. Introduce a 3 mm. column of tungstic acid supernatant (page
198) or other urea soln. to be analyzed, draw it in away from the
end, and seal the end with plasticine.

4. Accurately measure the length of the air column between the
two liquid columns with a filar micrometer temporarily substituted
for the right ocular which contains a disc micrometer. Two successive
readings must agree within 1 micrometer scale division (5 fx).

5. Replace the right ocular, cut off the end of the tube sealed
with plasticine, move the urea column back to the end of the capil-
lary with the water manipulator, and add 3 mm. of sodium hypo-
bromite soln. from a blunt-tipped pipette freshly filled just before
use. Should gas bubbles appear immediately upon the addition of
the reagent, discard the tube, and prepare fresh reagent.

6. Move the liquid column away from the end of the tube and
seal with plasticine.

7. Carefully remove the tube from the water manipulator by
cutting it about 6 cm. from its end, revolve it between thumb and
forefinger for a few sec. and set aside in a nearly vertical position
upon a plasticine mount.

8. After 2 hr. again revolve the tube for a few sec, place on the
microscope stage and accurately measure the length of the air column
with the filar micrometer.

9. Run a blank determination with distilled water and subtract
the increase in the length of the air column from that found above.
For each milligram per cent of urea nitrogen the increase averages
four scale divisions (20 /a) ; the increase in the blank averages five
divisions. Hence a soln. containing 10 milligram per cent urea
nitrogen should give an increase of 45 divisions.

216 CUVETTE COLORIMETRY

HYDROGEN ION CONCENTRATION

Capillary tube colorimetry has been employed for the measure-
ment of the hydrogen ion concentration of less than 1 /xl. liquid by
Montgomery (1935), who used quartz capillary tubes having an
internal diameter of 4-5 mm. When comparisons of indicator colors
given with protein-free buffer solutions were made, the error of
measurement was less than 0.02 pH. However when applied to
biological fluids, the indicator color may not be an accurate indica-
tion of the pH value. Montgomery (1935) observed that the capil-
lary tube method gave values for blood plasma from frogs and
Necturus which were consistently lower by an average of 0.11 pH
than those obtained with a glass electrode, a deviation which he
ascribed to the protein error of the indicator. It may be possible in
some cases of this nature to apply a correction factor. For electro-
metric measurements see page 183.

B. CUVETTE TECHNIQUE

1. Apparatus

General. Cuvettes for the colorimetric measurement of small
volumes of liquid have been designed for use with certain standard
colorimeters. Zeiss cuvettes having a capacity of 0.2 ml. are made
for the Pulfrich step photometer. The Evelyn photoelectric colorim-
eter has a micro attachment made to accommodate cells which re-
quire 0.15 ml. Adapters which enable 0.2 ml. cuvettes to be used with
the Coleman Junior spectrophotometer (model 6) are obtainable
from &. Ash (Lowry, Lopez, and Bessey, 1945). Quartz cuvettes
permitting the use of volumes of 0.05 ml., or less, with a special
adapter for the Beckman quartz spectrophotometer have been de-
scribed by Lowry and Bessey ( 1946) . With their adaptation meas-
urements can be carried out on 0.05 ml. volumes from about 225 to
1050 m^ with spectral widths of no more than 3 m/x. With 0.025 ml.
volumes a range of 235-935 m/x can be utilized with the 3 m^u,
spectral bands. The cuvettes and adapters may be obtained from
Pyrocell Manufacturing Co.*

* Since this writing a capillary absorption cell has been described by Kirk
et al. (see Bibliography Appendix, Ref. 40; see also Ref. 42).

APPARATUS

217

The degree of light absorption, and consequently the response in
the eye or in a photocell, is proportional to both the concentration
of the color substance and the length of the light path through the
solution. Therefore, a given quantity of color substance will effect
the same light absorption whether it is contained in a volume of
0.002 ml. and a 0.35 mm. light path is used (as in Richards' tech-
nique, page 195), or in a 0.060 ml. volume with a 10.5 mm. path.
Of course, the greatest absorption would be obtained by employing
the smallest volume with the longest light path.

i

Top %

I

t^ Light

^ Penny"

Light
Cuvette

^

^

Cuvette

; © ;

s y

Block

Diaphragm
(side)

Diaphragm (Type A)
(face)

°o*

— Holes for precision pins — '
of Beckman

o

■f

0o O ))

O

Adj

/
ustable

Diaphragm (Type B)

Fig. 77. Microcuvette and diaphragms.
Frovi Lowry and Bessey (1946)

Lowry and Bessey Adaptation of Beckman Spectropho-
tometer to Measurements on Small Volumes. The special cu-
vettes used have the same 1 cm. light path as the macro variety, but
the width of the chamber has been reduced to 2 mm. or less (Fig.
77) . A 0.05 ml. volume of liquid will fill the cuvette to a height of
about 2.5 mm. The height of the cell is 25 mm. and its outside cross-
sectional dimensions are the same as those of the macro vessel. The
inner cross-sectional dimensions of the macrocuvette are 10 X 10
mm. Cuvettes having an internal measurement of 1 X 10 mm. have
also been used; they require 0.03 ml. liquid, but their use is more
difficult.

218 CUVETTE COLORIMETRY

A diajihragm is placed in front of the cuvette to obtain a light
beam confined to a cross section of less than 2X2 mm. A beam of
this size can pass through the liquid without touching the meniscus
or the walls of the cuvette. The diaphragm (type A, Fig. 77) has
a metal disc the size of a penny through which a 1.0 to 1.4 mm. hole
is drilled about 1 mm. off center. Before the disc is fastened to the
metal sheet, it is held in the oi)ening from which the light enters
the cuvette and turned until the beam passes precisely in the middle
between the walls of the chamber when the cuvette is in place. The
disc is soldered at this angle to the sheet of metal (about 6X9
cm.) so that the hole coincides with a 3-4 mm. hole in the sheet 2.5
cm. from one end. The top of the sheet is bent at a right angle to
form a flange which lies on the top of the instrument. Wooden
blocks are used to raise the cuvettes so that the light beam just
misses the bottom of the chamber. The diaphragm is inserted and
removed by loosening the bolts which hold the phototube housing.
The carriage for the cuvettes should be oriented to bring the cuvettes
as near the diaphragm as jwssible. The cuvettes are numbered and
always set in the holder with the same orientation.

The type B diaphragm (Fig. 77) contains a sliding strip of brass
with pinholes which can move in a channel cut in the sheet metal.
The diaphragm is inserted between the cuvette carriage and the
body of the instrument and the sliding strip is moved until a pinhole
coincides with the center of the cuvette. The stop on the strip is
then adjusted with a bolt so that the pinhole can be brought to the
same position each time. The different-sized pinholes can be brought
into position without disturbing the adjustment. Blocks are used
to raise the cells as with the type A diaphragm. By removing the
brass strip the instrument can be used with macrocuvettes without
disturbing the metal sheet.

To obviate the effect of "play" in the cuvette carriage, the cells
should be moved into position from the same direction. In use, the
microcuvettes are left mounted in the carriage. Samples are intro-
duced with fine-tipped pipettes, and removed by suction with fine
tipped glass tubes. A macro cell may be used in the first position in
the carriage for the solvent or other blank solution.*

*See Bibliography Appendix, Ref. 32.

CALCIUM 219

2. Methods

CALCIUM

Sendroy (1942b) adapted iodometric reactions, previously used
in titrimetric measurements of calcium, to colorimetry. Using an
Evelyn macro photoelectric colorimeter, the method was applied
to volumes of serum down to 20 /xl. (about 2 fig. calcium). Further
refinement could be obtained if the colorimetry were carried out with
smaller volumes in microcuvettes. The calcium is precipitated as
the oxalate; the latter is washed, dried, dissolved in acid, and reacted
with an excess of eerie sulfate. The excess of eerie ion is made to
liberate iodine from potassium iodide and the yellow color thus
developed is measured. A precision of ±2^0 has been reported. An
alternative method is measurement of the blue color formed when
starch is added to the iodine solution. However, the many factors
which influence the blue color make it a less desirable choice even
though the relative color intensity of the blue is about 100 times that
of the yellow (Sendroy and Alving, 1942). The method to be de-
scribed was developed for serum, but it can be adapted to other
fluids. A thorough study of different procedures for the determination
of serum calcium was made by Sendroy (1944). For titrimetric
methods see page 272.

Sendroy Method for Calcium

SPECIAL REAGENTS

Saturated Ammonium Oxalate (about 3.5%). Prepare at room
temperature using analytical reagent grade of the salt.

2% Ammonium Hydroxide. Dilute 2 ml. cone, ammonium hy-
droxide (26% analytical reagent grade) to 100 ml.

Water-Alcohol-Ether Mixture. Mix equal vol. distilled water,
absolute ethyl alcohol, or redistilled 95% alcohol, and ethyl ether
(analytical reagent grade, or absolute, or redistilled U.S.P. grade).

1 N Sulfuric Acid (approx.). Dilute 27 ml. cone, acid, sp. gr. 1.84,
analytical reagent grade, to 1 1.

0.2 N and 0.1 N Sidfuric Acid (approx.). Prepare from the 1 N
soln.

220 CUVETTE COLORIMETRY

0.1 N Ceric Bisulfate (approx.). Dissolve 29 g. anhydrous eerie
bisulfate in 1 A^ sulfuric acid to make 500 ml. (The greater ease
of solution of the bisulfate makes it preferable to the sulfate. The
bisulfate is obtainable in about 92% purity from G. Frederick
Smith Chemical Co.) Store in amber bottles and protect from
light.

In preference to ceric sulfate, Kochakian and Fox (1944) have stressed the
greater sharpness of the end point in titration with ammonium hexanitrato-
cerate using setopaHne C as the indicator. Prepare a 0.01 A^ soln. by dissolving
6 g. ammonium hexanitratocerate, reagent grade, in about 200 ml. 1 N per-
chloric acid and dilute to 1 I. with the acid. Do not heat during preparation,
and store in a black bottle in the dark. Prepare a 0.05% setopaline C soln. by
adding 50 mg. of the dye (Eimcr and Amend) to 100 ml. distilled water and
warming on a hot plate or bath. Precipitation occurs on cooling, therefore the
soln. must be warmed before use and used while warm.

0.0035 N, 0.001 N, 0.0007 N, and 0.00035 N Ceric Bisulfate
(approx.). Prepare these solns. just when needed from the 0.1
N soln. Use 0.2 A^ sulfuric acid to dilute the 0.1 A'' to 0.0035 A^.
Use 0.1 N sulfuric acid for dilution to the weaker concentrations.
Prepare the 0.007 A^ and 0.00035 A^ solns. from the 0.0035 A^ soln.
Store in amber bottles and protect from light.

Standard 0.1 N Sodium Oxalate. Dissolve 3.3498 g. sodium oxalate,
analytical reagent grade, in 52 ml. 1 A^ sulfuric acid and add water
to make 500 ml. Store in amber bottle; the soln. is stable for at
least 6 months.

Standard 0.0005 N, 0.00025 N, and 0.0002 N Sodium Oxalate. Pre-
pare fresh when needed from the 0.1 A'' soln. by dilution with
water.

0.5% and 1% Potassium Iodide (approx.) Prepare fresh for use
from analytical reagent grade of the salt. (When tested with
starch, no trace of free iodine should be present.)

95% Ethyl Alcohol. Filter through two layers of ashless filter paper
on a Buchner funnel.

2% and 1% Starch (Lintner Soluble). Prepare the 2% soln. in
saturated sodium chloride soln. every 2 weeks by making a paste,
diluting to vol. and boiling for 5-10 min. Prepare the 1% soln.
fresh for use from the 2% soln. by diluting with water.

CALCIUM 221

PROCEDURE

1. Mix 5 vol. distilled water with 1 vol. serum in a 12-15 ml.
centrifuge tube; run in duplicate. Add 6 vol. distilled water to
another centrifuge tube to be treated in a parallel manner as a
standard; run in duplicate.

2. Add 1 vol. saturated ammonium oxalate to each tube, stir by
tapping, cover tubes to keep out dust, and let stand at least 16 hr.

3. Centrifuge for 5 min. at 2600 R.P.M. and carefully siphon
off all but about 0.2 ml. of the supernatant with an upturned capil-
lary, the tip of which is kept immersed.

4. Wash the entire inner surface of each tube with 3 ml. 2%
ammonia added slowly from a pipette moved around the top of the
tube.

5. Tap the tubes until the precipitates just begin to move up;
then again centrifuge and withdraw the supernatant.

6. Add 1 ml. of the water-alcohol-ether mixture; stir and mix
well. Add 3 ml. more of the mixture and mix gently to keep a mini-
mum of the precipitate in the upper portion of the liquid. Centrifuge
and withdraw the supernatant.

7. Repeat the washing with the water-alcohol-ether mixture as
in step 5.

8. Place the tubes in an oven at 100-110° at an angle of about
15° for 0.5-1.0 hr. to dry completely.

9. To each of the two standard tubes add different vol. of the
standard oxalate soln. (See Table VI.) Add the sulfuric acid to
all the tubes (see table) and heat for 5 min. in a beaker of water
kept below boiling.

10. Remove tubes, let cool to room temperature and add the eerie
bisulfate (Table VI). Mix well, cover the tubes, and let stand at
room temperature for 30 min. or in a water bath at 70° for 10 min.

11. Transfer solns. to vessels in which color development and
dilution to final vol. (Table VI) are carried out. To facilitate trans-
fer, coat a part of the outer rim of the tube with a thin film of
paraffin. Wash out tubes with 4 ml. portions of water to make the
transfer quantitative.

12. Add potassium iodide (Table VI) with a minimum of
agitation necessary to mix well. After 60 sec. add filtered alcohol
and then water to bring to final vol.

222

33

to

s

>

(N

O

o

a
w

01

O .

o S

w a

3

S
O

CO

3
T3

«
o

c

s
O

■ a

o

O "5

o o

Q O

I

00

o a

ffi

«

o o o

(M (N C<l

05

o o o

00 GO 00

o >ra
o o

Q O

I

00

o o o

o c^

o o

Q o

I

CO
00
1/5

o o o

o
o

o
o

CD

o o o

'^ tJ^ tJ<

5> (M C^ (N

^

oO •

°* g
o

^ o o

O "^ ^

•s-9 «■

(M

O O iC
(M C<i (N

o feO .

^^ 03 CI —

o c 2; °

o

o
o

d
in

3
t>

=0 "H

— • 33

s C

^ 33

e

33

c

03

CO CO CO

odd

LO IC

d d

o o

o o o

■^ "^ "^

CO CO CO

<6 'S <6

CO
o
o
o

d

o o

^

o

CO
83

lO lO

d d

lO lO IC

d d d

b- o o

o

O ^H 1— I

o
d

lO

(N

o
o

o

3 ^

I §

• ^ ^

!2 S 33

o ^ n

d c» Ph

<

d

o o o
-- — (N

! ^

o o

g o

8 °

3 ^

^ - ^

CO ^ ^

_: 33 3
■3-^4)

S 33

d CK Ph

d
o"

CO

O !»

.i a.

^ 3

03 -

ci ^~^

Co

"-o

00-*

?§s

o 2

^ a

00-3

(M 2

CO 2

^ a

o^

OJO

MM

3^

2S

«^o

■^co

£ =»

■3 <o

•- 3

^t

■*^ ,s

"l^-s

<B.S

.■t^.3

a-^

tc :s

c

?^:£

-S u)

• <-i

u ^^

Oi .^

-iJ .^

a a

tc

CO o3

03 ^

tiD(-5

a>

T30

CD CD

03

.3 .

!0 0

>

a-T3

03

c

.^-^

-^- 3

0

3 to

.— 0 .

^ '^

m. th
0. 40
ivelv

f^oo

aZ"£

»3 IC

3 .

.n £ a

•- 0

^^ S CO

co;§2J

fi^

.^•7

ooy2 >

0) s

^^ a

> *s

No.
(icon
0)m

wa

s 3

33'" 0

"^.3

i: 3cD

3 &

-Ui /vJ

let Ul
lynd
imum

3 S

0 3

II

0 ji X
L- > 33

>H a

bC ^

r/3 "•

Cornin
to 635

-1

a;cD 0

iJ «

^COqO

CQii

H 3"^

r^

' i-g

A c

CALCIUM 223

13. Prepare simultaneously reagent blanks containing sulfuric
acid, potassium iodide, and alcohol in the same concentrations as in
the standards and unknowns. Use these blanks to set galvanometer
at 100 just before reading the standards and unknowns.

14. Read 10 ml. portions at 25 ±5° in the Evelyn colorimeter
with filters indicated in the table. The yellow color may be read at
any time within 1 hr. after addition of the potassium iodide. Data
are given in the table for use of blue color with starch if filters for
yellow color are unavailable. In the latter case, add water to about
80% of vol. after the potassium iodide has been added, add the
starch slowly to the soln. at 25 ±1°, mixing by rotation, add water
to vol., and read color promptly at 25 ±1°. The blue-color method
is not reliable for samples of serum smaller than 0.05 ml.

Result. Since blue color readings are not reproducible from day
to day, a new calibration curve must be obtained for each day's
work. Galvanometer readings plotted semilogarithmically are linear
functions of oxalate concentration. For calibration of yellow-color
readings, the percentage transmission readings of 90, 80, 70, 60, 50,
40, 30, 25, 20, 15, and 10 correspond to the respective values of
iodine, in milliequivalents per liter in the color solns., of 0.0022,
0.0048, 0.0077, 0.0111, 0.0151, 0.0200, 0.0267, 0.0308, 0.0362, 0.0433,
and 0.0544. To redetermine the calibration curve, treat 0.5 to 0.7 ml.
of 0.133 mM potassium iodate with 2.4 ml. of 0.085 M phosphoric
acid and 1.2 ml. of 5% potassium iodide, dilute to 40 ml. with
filtered 95% alcohol, and then to 100 ml. with water. Read with
filter No. 586-5.

Calculate the concentration of the calcium (in milliequivalents
per liter) in the unknown from:

j-C-C^s^, _ ^j^^^-j^

where Si, So, and u refer to standards and unknown sample, C =
(C204^")si + {C204^~)s2, and D = V/v where V is the volume (in
ml.) at final dilution of color soln. and v is the volume (in ml.) of
original sample used. The C and D values are given in the table and
the (lo) values are obtained from calibration curve data correspond-
ing to the readings observed.

224 CUVETTE COLORIMETRY

When 0.02 ml. samples of serum are used it has been found neces-
sary to correct the yellow color readings for the effects of traces of
residual serum. The correction varies with the reading and is to be
subtracted from the latter: for readings of 15, 20, 30, 40, 50, and 60
subtract corrections 0^-^, 0-, 0^, l'^, V, and P, respectively.

example: Analysis of 0.02 ml. samples of a serum gave yellow color readings
of 36\ 36^ for the standards, 23^ and 55". The former were corrected to 35^ by
subtracting 1°. Values for iodine in milliequivalents per liter from the calibra-
tion curve were 0.0230 for the serum, and 0.0325 and 0.0130 for the standards.
Then the calcium concentration in the original serum (in milliequivalents
per liter) was:

[(°°^°° + 0.0325 + 0.0130^ _ „^23g-| ^ ^ ^ ^^^

In the case of blue-color readings a semilogarithmic plot is made with
milliequivalents of oxalate per liter from 0 to 0.0171 as abscissa and galvanom-
eter readings from 10 to 100 ordinates. A straight line is drawn between
the points representing the readings of the two standards, Si and S2, at concen-
trations of 0.016 and 0.004 milliequivalent oxalate per liter. Oxalate values for
serum analyses, obtained by interpolation of their galvanometer readings on
this line, times D give directly milliequivalents calcium per liter in the sample.

example: Analysis of 0.05 ml. samples of a serum gave blue color readings of
32^ and 32*, for the standards, 2V and 49^. A straight line was drawn through
the two latter values located at 0.016 and 0.004 milliequivalent oxalate per liter,
respectively. Interpolated values for the serum were 0.01005 and 0.01013 milli-
equivalents per liter. Then the average calcium concentration in the original
serum was:

0.01008 X 500 = 5.04 milliequivalents per liter.

CHLORIDE

Colorimetric chloride methods have not been specifically adapted
to histochemical work. However, the procedure of Sendroy (1939b,
1942a), which was designed for use with the macro Evelyn photo-
electric colorimeter, could be adapted to the smaller quantities
sufficient for use with microcuvettes. Even with the macro apparatus,
10 lA. serum is adequate for analysis. The principle of the Sendroy
method is conversion of the chloride in acid solution to its silver
salt by shaking with solid silver iodate; the iodate liberated by the
chloride is made to act on potassium iodide, and the yellow color of
the iodine set free is measured using filter No. 420 with the Evelyn
instrument. For other methods see pages 200 and 281.

CHLORIDE 225

Sendroy Method for Chloride

SPECIAL REAGENTS

Approxwiately 0.085 M Phosphoric Acid. Test to make sure soln.
is halide free.

Caprylic Alcohol.

Silver lodate Powder, C. P. Test for presence of potassium iodate
according to Sendroy (1939a): (1) Solubility measurement —
lodate analysis of a saturated soln. of silver iodate should give a
value not exceeding 0.21 mM/1. (2) Analysis of a standard
chloride soln. — Analyze a known 100 mM chloride soln. diluted
twenty times with 0.085 M phosphoric acid.

5% Potassium Iodide. Test with starch to be sure no trace of free
iodine is present.

PROCEDURE

1. Dilute the sample chloride soln. in 0.085 M phosphoric acid,
or in tungstic acid soln. if protein is present, at between pH 2.0 and
3.0 to a final concentration of between 3 and 12 mM/\.

2. Add solid silver iodate (10 mg./ml.) to duplicate portions in
15 ml. centrifuge tubes, and shake vigorously for 2 min.; then either
filter through halide-free paper or centrifuge for 1 min. at over
3000 R.P.M. The soluble iodate is now equivalent to the chloride in
the sample.

3. Sendroy (1942a) recommended that the system given in
Table I, B, of the paper by Sendroy and Alving (1942) be followed
for the measurement of chloride in serum and blood. In this schedule
0.5-13 ml. iodate soln. containing 0.8 mM is added to 2.4 ml. 0.085 M
phosphoric acid and 1.2 ml. 5% potassium iodide; the vol. is made
up to 100 ml. with water.

4. Promptly after the color has been developed, transfer the
tubes to a 25° water bath for 3-5 min. Wipe the tubes clean and
dry; set the Evelyn instrument with filter No. 420 to read 100 with a
blank soln., and then read the unknowns. The blank soln. may be
prepared by omitting the iodate from the reaction mixture.

5. A calibration curve has been given by Sendroy and Alving
(1942), but it is usually well to obtain one's own curve with the
particular reagents and instrument used. The calibration may be
made with standard potassium iodate solns.

226 CUVETTE COLORIMETRY

PHOSPHATE AND PHOSPHATASE

Siwe (1935) has described the use of the Pulfrich step photometer
with filter No. 72 (red) for the colorimetric measurement of in-
organic phosphorus in small amounts of blood by conversion to
phosphomolybdic acid and reduction by aminonaphtholsulfonic acid.
The same reaction was employed by Weil and Russell (1940) in
their phosphatase method ( page 209) . At about the same time Lund-
steen and Vermehren (1936) developed a micro procedure for the
determination of inorganic phosphate and alkaline phosphatase in
blood plasma based on Miiller's (1935) amidol reduction of phos-
phomolydic acid. The Pulfrich instrument with filter No. 72 was also
used in this case, and for most measurements the 10 mm. cells were
employed, but the 20 mm. cells were required for weaker colors. 50
fA. of blood are needed for a duplicate determination, and since the
procedure might be adapted to tissue extracts as well, it will be
described.

Conditions for the determination of inorganic phosphate in the
presence of labile phosphate esters, such as phosphocreatine, acetyl
phosphate, and ribose-1-phosphate, were established by Lowry and
Lopez (1946). As these authors pointed out, the usual procedures
for measurement of inorganic phosphate in tissue extracts represent
the sum of the inorganic phosphate and the phosphate of the labile
esters hydrolyzed by the reagents employed in the determination.
The procedure of Lowry and Lopez is based on the reduction of
phosphomolybdate by ascorbic acid at pH 4.0.

Bessey, Lowry, and Brock (1946) utilized as the substrate p-nitro-
phenyl phosphate, which had been studied by King and Delory
( 1939) , and applied to phosphatase determinations by Ohmori
(1937) and Fujita (1939). Bessey et al. were able to determine the
phosphatase in as little as 5 ju.1. serum using 0.5 ml. of solution for
the colorimetry. The advantage of this substrate is that it is color-
less and yields the yellow salt of p-nitrophenol when the phosphate
group is split off. Thus the color develops in proportion to the degree
of the hydrolysis and no additional reagents are required for the
color development. This advantage is also to be found in the use
of phenolphthalein phosphate, which was employed by Huggins and
Talalay (1945). However, alkaline phosphatase splits the p-nitro-
phenyl phosphate 25-30 times faster than the phenolphthalein com-

PHOSPHATE AND PHOSPHATASE 227

pound, 15% faster than phenyl phosphate, and 2-3 times more
rapidly than glycerophosphate, according to Bessey, Lowry, and
Brock. Either acid or alkaline phosphatase may be determined with
the substrate; it is only necessary to carry out the colorimetry in
alkaline solution, since the free nitrophenol, which would exist in
acid solution, is colorless. For other methods see pages 124, 208, 209,
and 280.

Lundsteen and Verniehren Method for Inorganic Phosphate
and Phosphatase

SPECIAL REAGENTS

Substrate. Combine 8 ml. 1 A^ ammonium hydroxide, 12 ml. 1 N
ammonium chloride, 1 g. disodium-;S-glycerophosphate, 2 ml. 1 M
magnesium chloride, and make up to 100 ml. with water.

10% Trichloroacetic Acid.

Acid-Molybdate Solution. Combine 100 ml., 7.5% ammonium
molybdate, 45 ml. 10 A^ sulfuric acid, and 105 ml. water.

Amidol Solution. Dissolve 15 g. sodium sulfite and 1.5 g. Amidol
(Agfa) in 100 ml, water. Store in the dark and cold, and dilute
five times before use. After about 2 weeks it turns red and can no
longer be used.

PROCEDURE

1. If blood is used, pipette 50 ^1. into 1 ml. of 0.9% sodium
chloride soln. and centrifuge out the cells.

2. To 200 ix\. of the supernatant fluid or a tissue extract add 200
fj\. substrate soln. and place in thermostat for the digestion period
(for plasma 24 hr. at 37°).

3. To another tube containing the same ingredients add 300 /*!.
10% trichloroacetic acid for a control experiment.

4. Stop the reaction by adding 300 fA. 10% trichloroacetic acid,
and centrifuge out the precipitate from both the enzyme and control
tubes.

5. To 400 /xl. of the supernatant in each case ^dd 100 /xl. acid-
molybdate reagent and 100 fA. Amidol soln.

6. Measure the color intensity after the soln. has stood for 15
min., and obtain the quantity of free phosphate from a previously
determined calibration curve constructed from measurements with
known amounts of phosphate.

228 CUVETTE COLORIMETRY

Lowry and Lopez Method for Inorganic Phosphate in Presence
of Lahile Phosphate Esters

SPECIAL REAGENTS

Protein Precipitant. 5% trichloroacetic acid, or 3% perchloric
acid, or (with very labile esters) saturated ammonium sulfate
which is 0.1 A^ with respect to acetic acid and 0.025 A^ with respect
to sodium acetate (pH 4).

0.1 N Sodium Acetate.

Acetate Buffer (pH 4) . 0.1 X to acetic acid and 0.025 N to sodium
acetate.

1 % Ascorbic Acid.

1% Ammonium Molybdate in 0.05 N Sulfuric Acid.

0.05 mM Standard Phosphate Solution.

PROCEDURE

1. Deproteinize the sample with ice-cold protein precipitant. If
either of the acid precipitants is used, bring the extract rapidly to
pH 4.0 to 4.2 by adding 4 vol. of 0.1 .V sodium acetate. (Most labile
esters are fairly stable at this pH.)

2. Dilute the extract with the acetate buffer until the inorganic
phosphorus concentration is 0.015 to 0.1 mM (0.05 to 0.3 milligram
per cent) . Dilute ammonium sulfate extracts at least five times.

3. Add 0.1 vol. 1% ascorbic acid and 0.1 vol. 1% acid-molybdate
soln. to each vol. of extract. If used within 15 min. of their mixing,
the ascorbic acid and molybdate may be combined.

4. Carry out the colorimetric reading at 5 and again at 10 min.
after the addition of molybdate using light between 650 and 950
vcijx (maximum absorption at 860 m/x) .

5. Take simultaneous readings of the standard soln. and blank,
which should be prepared parallel with the unknown. Should a
difference be observed in the readings of the unknown at 5 and 10
min. compared to the standard, extrapolate the values to zero time.

note: Lowiy and Lopez have found the reaction to be delayed in the
presence of certain tissue extracts. In these cases standardization must be
obtained by adding a known quantity of inorganic phosphate to an aliquot
and using the difference in the readings effected by the added phosphate
for the standardization. Dilution overcomes the inhibitory effect to some
degree. Thus, brain and muscle extracts should be diluted to a volume
150-200 times that of the tissue, and, in the case of liver 300-500 times.

PHOSPHATE AND PHOSPHATASE 229

Furthermore, acceleration in color development may be accomplished by
increasing the molybdate to 1.5% in 0.05 A'' acid, and the ascorbic acid
concentration to values that do not exceed a final concentration of 0.2%.

Bessey, Lowry, and Brock Method for Phosphatase

SPECIAL REAGENTS

Buffer-Substrate Solution. (pH 10.3 to 10.4). Prepare soln. A by
dissolving 7.50 g. (0.1 mole) glycine and 95 mg. (0.001 mole)
magnesium chloride in 700 to 800 ml. water; then add 85 ml. 1 N
sodium hydroxide and dilute to 1 1.

Prepare soln. B, which is 0.4% disodium p-nitrophenyl phos-
phate in 0.001 A^ hydrochloric acid. (The authors reported that the
Eastman Kodak Co. product contained about 50% inert material;
hence twice the quantity of this preparation should be used.
Purification may be carried out by recrystallization from hot S7%
alcohol.) Adjust the pH of soln. B to 6.5 to 8.0 with acid or base,
if necessary. Test for free nitrophenol by diluting 1 ml. with 10
ml. 0.02 A^ sodium hydroxide and measuring the absorption at
415 m^. If the extinction is greater than 0.08 {i.e., transmission
less than 83% for 1 cm. liquid, or 70% for 2 cm.) remove the
free phenol by extracting two to three times with equal vol. water-
saturated butyl alcohol followed by three extractions with water-
saturated ether. (All butyl alcohol must be removed since it in-
hibits phosphatase activity.) Aerate off the traces of ether and
store in the cold.

Mix equal vol. of solns. A and B; adjust the pH to 10.3 to 10.4
if necessary with strong sodium hydroxide or hydrochloric acid,
and store in the cold, or better, in the frozen state. When 2 ml.
diluted with 10 ml. 0.02 A^ sodium hydroxide has an extinction
greater than 0.1 for 1 cm., either discard or extract it with butyl
alcohol, as above, and readjust the pH.

Standard Solutions. Prepare solns. containing 1, 2, 4, and 6 milf
p-nitrophenol (molecular weight 139.1) per liter.

PROCEDURE

1. Place 5 ^1. serum in the bottom of a 6 X 50 mm. serological
tube, immerse in ice water, and rapidly add 50 jxl. of the ice-cold
buffer-substrate soln. with a constriction pipette. Alix by tapping
with the finger.

230 CUVETTE COLORIMETRY

2. Digest at 38° for 30 min., and then place in ice water and add
0.5 ml. 0.02 A^ sodium hydroxide with sufficient force to mix the solns.

3. Transfer to a cuvette and measure the color intensity using
light at 400-420 m/^.

4. Add 2-4 ix\. cone, hydrochloric acid and take a second colori-
metric reading. The difference in the optical densities gives the
corrected density of the unknown.

5. Run standards and blanks by treating 5 /xl. vol. of the
standards and distilled water in the same manner as the serum.
Construct a standard curve from the corrected optical densities. If
the same pipettes are used for both the standards and unknowns, the
exact pipette vol. need not be known.

Bessey et al. employ a "millimole unit" which is defined as the
phosphatase activity which will liberate 1 milf nitrophenol/liter/hr.
1 millimole unit is approximately equal to 1.8 Bodansky units. For
sera weaker in phosphatase, such as those from adults, the vol. serum
and reagent may be doubled without increasing the vol. alkali; this
will yield a color of nearly double the intensity.

NITROGEN AND AMMONIA

General

As in the more macro methods, the determination of nitrogen in
histochemical investigations involves conversion of the total nitro-
gen to ammonium sulfate by digestion. The digest may be Ness-
lerized directly, or it may be alkalized and the liberated ammonia
absorbed in acid and measured either colorimetrically or titrimetri-
cally. However, the determination of the very small quantities in-
volved requires unique treatment. Since the preliminary procedures
are common to both the colorimetric and titrimetric methods, it
will perhaps contribute to greater integration and clarity if the
chief developments in both forms of analysis are considered together
in chronological order at this point.

At about the same time, Linderstr0m-Lang and Holter ( 1933b)
and Conway and Byrne (1933) reported methods for the micro-
estimation of ammonia which depended on the transfer, by diffu-
sion, of the ammonia from an alkalized solution to one of standard
acid followed by titration. Linderstr0m-Lang and Holter employed a

NITROGEN AND AMMONIA 231

tube having a total capacity of about 250 /xl. for the diffusion process.
The sample was placed in the bottom of the tube and the ammonia
from it was allowed to diffuse into a drop of standard acid placed
in the upper part of the tube to form a seal across the lumen (Fig.
46). Conway and Byrne used a special diffusion cell (Fig. 41) re-
quiring considerably more of the liquids; hence it was not suitable
for the accurate measurement of much less than 14 jug. ammonia
nitrogen. The Linderstr0m-Lang and Holter method had a precision
of 0.005 fxg. nitrogen and it could be used for the determination of up
to about 28 /xg. The following year Gibbs and Kirk ( 1934) employed
a modified Conway-Byrne procedure, which they used for the esti-
mation of from 1.5-8.3 fig. ammonia nitrogen. Conway (1935b)
subsequently described a refinement of the diffusion cell method
which had an ultimate standard deviation of 0.02 /xg. ammonia
nitrogen.

Levy (1936) developed a technique for the determination of total
nitrogen based on the direct Nesslerization of the acid-digested
sample. The complete treatment was carried out in the same vessel
and the final solution was transferred for colorimetry to a micro-
cuvette having a capacity of 0.2 ml. Levy's method was adapted for
quantities of nitrogen in the range 0.5-6.0 /xg. and the average devia-
tion observed was 0.03 jug.

A titrimetric method for total nitrogen employing features of both
the Linderstr0m-Lang and Holter and the Conway techniques was
published by Needham and Boell ( 1939) . These investigators used
a single vessel with a special cap for all the operations, i.e., digestion,
diffusion, and titration. The method was adapted to 1-20 /xg. of
nitrogen and the standard deviation in control experiments was 0.3
fig. In order to refine the earlier colorimetric method of Borsook
(1935), Borsook and Dubnoff (1939) also borrowed features of the
Linderstr0m-Lang and Holter technique as well as the diffusion cell
of Conway and Byrne to develop a method for total nitrogen, am-
monia, and other nitrogenous compounds. The procedure of Borsook
and Dubnoff for 5-10 fig. total nitrogen involves acid digestion,
transfer of an aliquot to a diffusion cell, and finally electrometric
titration of the excess acid. The standard deviation was around
0.05 fxg.

Levy and Palmer ( 1940) adapted the hypobromite method for am-
monia to the iodometric estimation of nitrogen without diffusion.

232 CirV'ETTE COLORIMETRY

The principle of this method, which was originally proposed by
Rappaport (1935), is iodometric measurement of the excess hypo-
bromite remaining after reaction with ammonia according to the
equation: 3 NaOBr + 2 NH3 -^ N2 + 3 H2O + 3 NaBr. The pro-
cedure of Levy and Palmer can be used for total nitrogen in the
range 500-5 /xg. or, if the microvolumetric techniques of Linder-
str0m-Lang and Holter are used, down to 0.5 [xg. After digestion in
small test tubes the material is diluted with water, made alkaline
with a neutralizing reagent, treated with an excess of hypobromite,
and this excess is measured by the addition of potassium iodide and
titration with thiosulfate. The entire treatment can be carried out in
one small tube.

As an improvement on the diffusion procedure of Bentley and
Kirk (1936), Tompkins and Kirk (1942) described a specially con-
structed unit for both digestion and diffusion designed for the meas-
urement of samples containing 0.5-20 /xg. nitrogen. A probable error
of not more than about 1% was claimed by the authors, but this was
challenged by Hawes and Skavinski (1942), who stated that the
diffusion time employed by Tompkins and Kirk is sufficient to trans-
fer only about 90% of the ammonia in samples containing less than
10/xg. nitrogen.

Hawes and Skavinski ( 1942) described another modification of
the diffusion cell technique. They employed a test tube for both di-
gestion and diffusion. A small helix of platinum wire, which was
sealed to a glass tube held in a rubber stopper, was used to hold a
drop of the ammonia-absorbing solution. AIM primary sodium
phosphate solution, rather than the commonly used saturated boric
acid solution, was employed to absorb the ammonia because of the
greater absorption capacity of the former. The volume of phosphate
solution need not be measured ; it is sufficient merely to dip the helix
into the stock solution. After the absorption is complete, the helix is
immersed in water and the solution is titrated with standard acid
to an end point between pH 4.3 and 4.7. Electrometric titration is
advantageous but an indicator may be used; in the latter event a
bromocresol green-methyl red mixture is superior to the methylene
blue-methyl red combination. As used by Hawes and Skavinski, the
method enables the determination of from 10 to 100 fig. nitrogen.

Russell ( 1944) , by utilization of the Conway-Byrne diffusion cell
for the collection of ammonia, refined the phenol-hypochlorite

NITROGEN AND AMMONIA 233

inetliud for the colorimetric determination of ammonia, which had
been used earlier by Van Slyke and Hiller ( 1933) and Borsook
(1935). By the reaction of ammonia with phenol and hypochlorite
in alkaline solution an intense blue product is formed, which is be-
lieved to be indophenol or a closely related compound. The method
has been applied to 1.5 ml. ammonia solution containing 0.5-6.0 ixg.
nitrogen. Larger or smaller volumes may be used to extend the range
of the method.

Boell (1945) employed a method for the estimation of 1-50 /xg.
tutal nitrogen which utilized a digestion similar to that of Levy
( 1936) , transfer of an aliquot of the digest to a diffusion unit made
up of ordinary laboratory glassware rather than the special cells of
Conway and Byrne, and titration with a simplified microburette.
The standard deviation was about 0.1 [xg. nitrogen.

The method of Linderstr0m-Lang and Holter (1933b) for am-
monia has been adapted to the determination of total nitrogen and
subjected to exhaustive critical trial and investigation in the Carls-
berg Laboratory. Changes have been introduced as dictated by
experience and a description of the procedure used in 1939 was
given in a publication of Bottelier, Holter, and Linderstr0m-Lang
(1943). A final summary of the method after further improvements,
with a full treatment of each step, was the subject of a paper by
Briiel, Holter, Linderstr0m-Lang, and Rozits (1946). In principle,
the final method consists merely of the digestion of the sample,
transfer of the digest to the bottom of a paraffin-coated tube, and
titrimetric measurement of the ammonia in a manner similar to
that originally employed by Linderstr0m-Lang and Holter but with
improvements in certain of the details.

The preceding survey indicates that various procedures may be
used for every step in the nitrogen determination. The choice of
method may depend to some degree on the prejudices and prefer-
ences of the experimenter, but the method of Briiel et al. ( 1946) is
recommended as the most foolproof and reliable.

Titrimetric Methods. The Levy and Palmer (1940) method
has the great advantage of employing a simple small tube for all of
the chemical steps, and the diffusion process is avoided altogether.
Certainly the digestion of the sample can be performed in a small
tube regardless of the ensuing procedure. If an ammonia diffusion is
to be employed, it is simpler to use small tubes for it rather than

234 CUVETTE COLORIMETRY

the specially constructed vessels of Conway and Byrne (1933),
Needham and Boell (1939), or Tompkins and Kirk (1942). In this
regard, advantages might be claimed for the method of Briiel et al.
(1946), in which both diffusion and titration are carried out in the
same simple tube, or the Hawes and Skavinski (1942) method, in
which the digestion and diffusion are conducted in the same tube.
However, the former requires transfer of digest and the latter
transfer of acid. The method of Briiel et al. ( 1946) is the most precise
of all (0.005 /xg. nitrogen), and the one most thoroughly tested for
its reliability. This method is described on page 283.

Colorimetric Methods. Levy's (1936) method for the direct
Nesslerization of the digest has the advantage that all of the chemi-
cal operations may be carried out in the same vessel and the diffu-
sion process is eliminated. On the other hand, the phenol-hypo-
chlorite reaction used by Russell (1944) is highly sensitive, and
ammonia, in the quantities that can be determined by Levy's
method, can be analyzed with ordinary macro equipment. However,
diffusion of the ammonia from the digest is essential for a proper
phenolhypochlorite reaction.

Digestion of Sample for Determination of Total Nitrogen

The digestion can be conveniently performed in small tubes of
resistant glass. In the earlier methods after the sample had been
introduced, a small Hengar granule {Hengar Co.) was sometimes
added to prevent bumping, the digestion mixture was introduced,
and the tubes were placed in a drying oven at 120-130° for a few
hours, or at 105-110° overnight, to drive off most of the water. The
digestion was continued by heating the tubes in a sand bath, or in
a copper or aluminum block with holes drilled so that the tubes
could be inserted to a depth not exceeding one third of the tube
length.

Briiel, Holier, Lintlerstr0m-Lang and Rozits Procedure for
Digestion of the sample. In the procedure of Briiel et al. ( 1946) ,
which should be given preference, the digestion tubes are 4 cm.
long, 1.8 mm. inner diameter, and 2.4 mm. outer diameter. The
tubes are cleaned by boiling in sulfuric acid-chromic acid mixture,
rinsed thoroughly with distilled water run through each tube by
means of a thin capillary extending to the bottom, and the water is

NITROGEN AND AMMONIA 235

removed by suction through a thin capillary. The tubes are dried
in vacuo at 100° for 10 min. The rims of the tubes are never touched
with the fingers after removal from the cleaning solution and the
clean tubes are stored in a desiccator or in petri dishes in a room
where no ammonia is allowed and smoking is prohibited. To the
sample in a tube about 4 fx\. of the following mixture is added with
a constriction pipette, properly centered so that the solution deliv-
ered to the bottom of the tube is not drawn up between the pipette
and the inner wall:

1 g. copper sulfate (CuS04-5H20)

10 g. potassium sulfate

0.2 g. sucrose

5 ml. cone, sulfuric acid

Dilute to 100 ml.

(The sucrose is added to make sure that the reduction of the nitrog-
enous impurities in the reagents also takes place in the blanks.)
The constriction pipette is emptied under constant pressure, which,
for the greatest accuracy, is so adjusted that the pipette automati-
cally empties when the tip touches a wall or liquid surface. Before
use the pipette is rinsed inside and out with distilled water. After the
digestion solution is added to the sample the tubes are placed in a
vacuum desiccator over phosphorus pentoxide and the water is
allowed to evaporate at room temperature. To prevent creeping the
bulk of the water is removed at 150 mm. mercury (about 24 hr.) ;
then the drying is continued at 0.1 mm. mercury (another 24 hr.) .

By means of a mouth-operated horizontal pipette with a vertical
delivery tip, 1 [A. cone, sulfuric acid containing 10 mg. selenium/ml.
is added to the charred sample in the digestion tube. (About each
10 mm. of the graduated length of the pipette corresponds to 1 [A.;
a strip of graph paper may be used for the graduation. The selenium-
acid solution is prepared by boiling until clear.) Proper centering of
the delivery tip is essential to prevent the liquid from being drawn
up between the tip and the wall of the tube. Should the liquid be
drawn up as far as the rim where contamination may occur, the
experiment should be discarded. It is also essential to rinse the
pipette both inside and outside each day before use.

The digestion is carried out in small flasks (Fig. 78), which may
be made by forming a constriction in the middle of insulin ampules.
1 ml. cone, sulfuric acid containing 0.4 g. potassium sulfate is placed

236

CUVETTE COLORIMETRY

in the bottom of the flask, and before being used for the first time
the solution is boiled in the flask to drive off water and gases. A
small glass ball (Fig. 79) is used to close the flask when not in use.
The flasks are heated conveniently in a copper block supplied with
an electric coil. Holes in the block permit the flasks to be inserted to
a depth of about 14 mm.; the temperature is kept constant at 295°.
At this temperature the acid is clear; it fumes but does not boil.

; u

/l2 mm.\

) C

)

> (. ;/A

)

14-5 mm.

Fig. 78. Arrangement for
digestion.

Fig. 79. Glass Fig. 80. Removal

stopper for diges- of tubes from diges-
tion flask. tion flask.

From Brilel et al. (1946)

To insert the digestion tubes the flask is removed from the block,
allowed to cool for 30 sec, and then the tubes are inserted by means
of forceps and arranged in a row around the circumference of the
flask (Fig. 78). After replacement of the flasks in the block the
digestion is watched for a few minutes to see whether liquid seals
form across the lumen of the tubes and rise upward. If they rise
higher than the middle of a tube the flask is removed from the block
for a minute or two and the seals collapse. After the initial stage the
digestion is allowed to proceed unattended for 5-6 hr. The tubes are
removed from the slightly cooled flasks by means of a conical glass
rod as indicated in Figure 80. They are rinsed on the outside with
distilled water, dried with a clean towel, and stored in a desiccator
over phosphorus pentoxide until ready for the determination of
ammonia.

NITROGEN AND AMMONIA 237

Digestion Mixtures Used by Other Workers. 1 ml. cone, sul-
furic acid, 0.625 g. potassium sulfate, and 0.075 g. selenium; dilute
about 5 times. — Levy (1936).

1% selenium dioxide and 1% copper sulfate (CuSO4.5Ho0) in
concentrated sulfuric acid and water (1:1) ; after digest is colorless,
cool, add 1 drop saturated potassium persulfate, and continue diges-
tion for 15 min. after the solution becomes water clear. — Borsook
andDubnoff (1939).

3 g. copper sulfate, 1 g. potassium sulfate, and 0.1 g. selenium
dioxide to 300 ml. cone, sulfuric acid. — Needham and Boell ( 1939) .

Cone, sulfuric acid and water (1:1) ; when fuming starts, add 1
drop of 30% hydrogen peroxide. After digest is clear, cool, add a
couple of drops of saturated potassium persulfate, and continue
heating; destroy peroxides by adding water, after cooling, add
another Hengar granule, and place in an oven at about 120° for
30 min. — Levy and Palmer (1940).

Cone, sulfuric acid and water (1:1) saturated with potassium
sulfate and made 0.1% with respect to copper selenite. The latter is
prepared by mixing a strong copper sulfate soln. with one of sodium
selenite and collecting the precipitate. — Tompkins and Kirk (1942).

18 A^ sulfuric acid containing 0.1% selenium dioxide and 0.1%
copper sulfate; use 30% hydrogen peroxide if needed. — Hawes and
Skavinski (1942).

I g. copper sulfate, 1 g. potassium sulfate, 1 g. selenium dioxide,
and 100 ml. 50% sulfuric acid; add saturated potassium persulfate
to the cooled mixture if required. — Boell ( 1945) .

Levy Nesslerization Method for Nitrogen

SPECIAL REAGENTS

Digestion Mixture. See above.

Nessler Reagent (Folin and Wu). Add 15 g. potassium iodide and

II g. iodine to 10 ml. distilled water and introduce an excess of
mercury ( 14-15 g.). Shake well for 7-15 min. or until the dissolved
iodine has nearly all disappeared. Cool in running water when the
solution begins to become pale and continue shaking until the
greenish color of the double iodide (HgIo.2KI) appears. Decant
the solution and wash out vessel with distilled water. Dilute the
solution and washings to 200 ml. Add 75 ml. of this solution and

238 CtfVEtTE COLORIMETRY

75 ml. distilled water to 350 ml. 10% sodium hydroxide (within
1% of 2.5 A^) to make the final Nessler reagent.

PROCEDURE

1. Place a microtome section of tissue in 7 jxl. water in the bottom
of a digestion tube (Fig. 39, page 167). This tube has a total capac-
ity of about 2.5 ml. Add 50 fA. of the Levy digestion mixture and
drive off most of the water at 130° (about 1.5 hr. required). Digest
until water- white (Levy uses a small mobile gas flame),

2. After the tube has cooled, hold almost horizontally and pipette
700 fA. distilled water directly into the digest, delivering the water
with as much force as possible. If complete solution is not attained
at once, it will occur on standing.

3. Add the Nessler solution in a standardized manner (Levy,
1936) : First, "A piece of glass tubing is drawn out to a long capil-
lary and clamped vertically above a platform which may be raised
and lowered by a rack and pinion. It is connected to a source of
air supply at 20 cm. water pressure. The capillary is of such diam-
eter that a rapid stream of air bubbles passes through when im-
mersed in water," Then, the reaction tube is placed in a holder and
set on the platform under the capillary tube with air flowing through
it. Fill a 300 fA. constriction pipette with Nessler reagent and place
one end of a rubber tube connected to the pipette in the mouth, and
hold the pipette in the right hand. "The pinion of the platform is
now turned with the left hand to raise the tube about the bubbler so
that a stream of air bubbles passes from bottom to top of the solu-
tion. Just as soon as the bubbler reaches the bottom the Nessler
reagent is blown in. The platform is then lowered. The entire process
takes five seconds." Tilt the tube back and forth to collect any
solution in the bulb,

4. Make the colorimetric measurement from 10-90 min. after
Nesslerizing. Prepare a blank by following the preceding steps, but
omitting the sample. (Levy used a Pulfrich photometer with filter
S43. The microcuvettes which hold 0.2 ml. were employed.)

Russell Plienol-Hypoclilorite Method for Ammonia

SPECIAL REAGENTS

Alkaline Phenol Reagent. Add some water to 25 g. crystalline
phenol and pour in with stirring 54 ml, 5.0 A^ sodium hydroxide.

UKIC ACID, CREATINE, CREATININE, AND ALLANTOIN 239

Make up to 100 ml. and store in a brown bottle in a refrigerator.

Hypochlorite Solution. Dissolve as much as possible of 25 g. of
ground and sifted calcium hypochlorite in 300 ml. hot water. With
stirring, add 135 ml. 20% potassium carbonate soln. which had
been boiled to drive out ammonia. Heat briefly to about 90°, cool,
and dilute to 500 ml. Filter a little of the soln. and test for calcium
ion by adding some of the potassium carbonate soln. and heating
in boiling water a few min. In the absence of calcium ion the
solution remains crystal clear. If the test is positive, add more
carbonate until a negative test is obtained. Filter the final soln.
and store in small brown bottles in a refrigerator. This soln.
should be water clear and contain 1.30-1.40% free chlorine. Test
for free chlorine by adding 10 ml. water, 2 ml. 5% potassium
iodide, and 1 ml. glacial acetic acid to 2.00 ml. of the hypochlorite
solution. Titrate with 0.100 A^ thiosulfate; 7.5-8.0 ml. should be
required. Occasionally retest the solution. The soln. may also be
prepared from Clorox (page 58),

0.003 M Manganous Chloride or Sulfate.

PROCEDURE

1. Place 1.5 ml. sample in neutral or acid solution not stronger
than 0.01-0.02 A^ (containing 0.5-6.0 /xg. ammonia nitrogen) in a
colorimeter tube, cooled by an ice bath. Add 1 drop manganous salt
soln., 1 ml. cold alkaline phenol reagent, and 0.5 ml. cold hypochlorite
solution. Mix by gentle rotation and place at once in a boiling water
bath for about 5 min.

2. Cool, dilute to a convenient volume such as 6 or 10 ml., and
measure the color intensity with light of about 625 m/x.

URIC ACID, CREATINE, CREATININE, AND ALLANTOIN

Borsook (1935) reported colorimetric methods for uric acid,
creatine, creatinine, and allantoin which were modifications of pro-
cedures already in use, but which are in the_ category of micro
methods, even though no special micro equipment is required. The
colorimetric measurements were carried out spectrophotometrically
in cuvettes taking 3.0-3.5 ml. With the present availability of micro-
cuvettes, these methods could be adapted to the analysis of much
smaller quantities of the substances by substituting smaller test
tubes for the 125 X 9 mm. (inside dimensions) tubes used.

240 CUVETTE COLORIMETRY

The uric acid method is based on precipitation of the suljstance
with zinc, solution of the precipitate in dilute acid and water, addi-
tion of cyanide followed by Benedict's arsenophosphotungstic acid
reagent, and development of the color by a procedure which yields a
clear solution. The method was designed for the analysis of 1 ml. of
sample having less than 1 milligram per cent uric acid, i.e., less then
10 /xg. For another method see page 213.

The creatinine method involves absorption of the compound on
Lloyd reagent, removal of impurities, and liberation of the creatinine
by the same alkaline picrate in which the color is developed. Creatine
is determined by the increase in creatinine after conversion to the
latter. With the 3 ml. cuvettes which Borsook used, the smallest
quantity of creatinine which could be measured was 1 ju,g. The
absolute error with all concentrations was ±0.1 jug. For another
method see page 211.

The allantoin method depends on the enzymatic conversion of
allantoin to allantoic acid (in the presence of cyanide to prevent the
formation of allantoin from uric acid), acid hydrolysis of the
allantoic acid to urea and glyoxylic acid, and colorimetric measure-
ment of the latter. With 2 ml. of sample, the least that can be
measured is 0.05 milligram per cent (1 /xg.) with an error of ±5%.

Sure and Wilder (1941) employed the micro attachment on the
Evelyn photoelectric colorimeter for the measurement of creatine
and creatinine. Gold-plated plungers had to be used because the
ordinary cadmium-coated plungers were corroded by the saturated
picric acid. The error in the conversion of creatine to creatinine by
the procedure employed varied from —0.79 to +2.66% in test
experiments. 1 ml. blood filtrate was used by the authors for each
analysis.

Borsook Method for Uric Acid

SPECIAL REAGENTS

2.5% Zinc Chloride.

10% Sodium Carbonate.

N/14 Hydrochloric Acid.

5% Sodium Cyanide (containing 2 ml. cone, ammonium hydroxide

per liter. Prepare fresh every 6—7 weeks.
Benedict's Arsenophosphotungstate Reagent. Dissolve 100 g. pure

URIC ACID, CREATINE, CREATININE, AND ALLANTOIN 241

sodium tungstate in 600 ml. water. Add 50 g. pm'e arsenic pent-
oxide followed by 25 ml. 85% phosphoric acid and 20 ml. cone,
hydrochloric acid. Boil for 20 min., cool, and dilute to 1 liter.
Folin's Stock Uric Acid Standard Soln. Place 1 g. uric acid, weighed
to 1 mg., in a 1 1. volumetric flask. Add 150 ml. water to 0.6 g.
lithium carbonate in a 250 ml. flask and shake until dissolved.
Filter, and heat the filtrate to 60°. Warm the flask containing the
uric acid in running hot water and pour the warm lithium carbon-
ate soln. into the volumetric flask containing the uric acid; wash
down crystals adhering to the neck. Shake until the uric acid has
dissolved (about 5 min.), cool under the tap, add 20 ml. 40%
formaldehyde, and half fill the flask with water. Add a few drops
of methyl orange and then pipette in slovdy, with shaking, 25 ml.
1 A^ sulfuric acid. Dilute the pink soln. to 1 1. Store the reagent in
v/ell-stoppered bottles and keep in the dark. Diluted standards
made from this soln. will keep for several days, but do not use
them sooner than 1 hr. after they are made.

PROCEDURE

1. Pipette 1-2 ml. of sample, 1 ml. water, and 0.05 ml. 2.5%
zinc chloride into a Pyrex test tube ( 125 X 9 mm. inside) provided
with a ground-glass stopper. Mix well by inversion several times.

2. Add 0.4 ml. 10% sodium carbonate and again mix by inver-
sion.

3. Centrifuge the test tube, pour ofT the supernatant, and take
up the last drop from the lip of the tube with filter paper.

4. Add 0.5 ml. N/14: hydrochloric acid, 1.5 ml. water, and 1 ml.
cyanide soln.

5. Stopper the tube and shake well to dissolve all precipitate
and leave a clear colorless soln.

6. Add 0.2 ml. arsenophosphotungstate soln. and again mix well
by inversion.

7. Place the stoppered tube in a 37° water bath for 40 min., and
then in an ice water bath for 15 min.

8. Centrifuge, transfer some of the supernatant to an absorption
cell, and determine the color spectrophotometrically at 610 mix.

9. In a parallel manner, treat four standard solns. covering the
range 0 to 1.0 milligram per cent uric acid in order to obtain a
calibration.

242 CUVETTE COLORIMETRY

Borsook Method for Creatine and Creatinine

SPECIAL REAGENTS

0.1 N Hydrochloric Add.

0.01 N Hydrochloric Acid.

Lloyd Reagent {Eli Lilly and Co.).

Alkaline Picrate Solution. Combine 10 parts saturated picric acid

and 1 part 10% sodium hydroxide.
Creatinine- Zinc Chloride Standard. Dissolve 1.61 g. creatinine-zinc

chloride (Benedict, 1914) in N/14: hydrochloric acid and make

up to 1 1. This soln. has 1 mg. creatinine/ml. Make up fresh at

least once a month.

PROCEDURE

1. Convert creatine to creatinine: Place 1-5 ml. of sample in a
Pyrex test tube ( 125 X 9 mm. inside) provided with a ground-glass
stopper, add one fourth the vol. of 0.1 A?" hydrochloric acid and mix
by inversion. Insert a piece of thread into the neck of the tube and
stopper. Then autoclave for 20 min. at 30 lb. (130°). Omit this step
for preformed creatinine.

2. After cooling, add 30-40 mg. Lloyd reagent and, with thread
removed, stopper tightly and shake continuously for 10 min.

3. Centrifuge, pour off supernatant, and remove the last drop
from the lip of the tube with filter paper.

4. Resuspend the precipitate in 1 ml. 0.01 A^ hydrochloric acid
and use another 1 ml. portion of the acid to wash down the stopper
and the sides of the tube.

5. Again centrifuge and discard the supernatant, removing the
final drop from the lip with filter paper.

6. Add 3 ml. sodium picrate soln. to remove the creatine from
the Lloyd reagent, and shake gently for 10 min. to develop the color
fully.

7. Centrifuge, and measure the color of the liquid spectrophoto-
metrically at 525 m/x. A linear relationship exists between the
absorption and the creatinine concentration from 0 to 2.0 milligram
per cent.

8. Adsorb standards on Lloyd reagent and treat as above from
step 2 on.

tmiC ACID, CREATINE, CREATININE, AND ALLANTOIN 243

Sure and Wilder Method for Creatine and Creatinine

SPECIAL REAGENTS

Saturated Picric Acid.

10% Sodium Hydroxide (carbonate free).

PROCEDURE

1. Pipette 1 ml. of sample (blood filtrate) into a 15 ml. centri-
fuge tube and add 0.5 ml. picric acid soln.

2. To convert creatine to creatinine, cover the tube with lead foil
and autoclave for 40 min. at 20 lb. pressure. Omit this step for
preformed creatinine.

3. Cool, add 0.1 ml. of the sodium hydroxide, and allow to stand
for 10 min.

4. Run a control on the reagents alone.

5. Using the Evelyn photoelectric colorimeter, set the control at
100 with filter 520-M, and then obtain readings of the unknown. Do
not allow the plungers to remain in contact with the soln. for longer
than 1-2 min. in order to avoid corrosion.

6. Obtain values from a previously established calibration curve.

Borsook Method for AUantoin

SPECIAL REAGENTS

Enzyme Powder. Use urease preparation (Squibb) from soy bean
meal (Van Slyke and Cullen, 1914).

Ammonium Carbonate-Sodium Cyanide. Dissolve 1.153 g. ammo-
nium bicarbonate, 0.891 g. ammonium carbonate, and 0.46 g.
sodium cyanide in water and make up to 200 ml.

10% Trichloroacetic Acid.

2% Sodium Tungstate.

N/15 Sulfuric Acid.

0.5% Phenylhydrazine Hydrochloride in A^/14 hydrochloric acid.
Dissolve commercial phenylhydrazine hydrochloride in water and
decolorize by boiling with activated charcoal. Filter the hot solu-
tion, and, after the filtrate is cooled in an ice-salt bath, precipitate
the phenylhydrazine hydrochloride by addition of cone, hydro-
chloric acid, or by dry hydrochloric acid gas. Filter the precipitate
off by suction; wash once quickly with very cold hydrochloric acid,

244 CUVETTE COLORIMETRY

and place in a desiccator over calcium oxide in the dark. Make

up the soln. just before use.
1.25% Potassium Ferricyanide. Prepare just before use.
Standard Solutio)i. Prepare fresh each week a soln. of 1 mg. allan-

toin/ml.

PROCEDURE

1. Into a Pyrex test tube (125 X 9 mm. inside) provided with a
ground-glass stopper place 10 mg. dry enzyme powder, 0.5 ml.
carbonate-cyanide soln., 2 ml. of the sample to be analyzed, and a
drop of chloroform.

2. Stopper the tube and let stand at 37° for 12 hr. An occasional
shaking aids in the solution of the enzyme.

3. Transfer 2 ml. of the mixture into another test tube and add
0.2 ml. 10% trichloroacetic acid and 0.1 ml. 2% sodium tungstate.
Mix after stoppering by inverting a few times. Then add 0.1 ml.
iV/15 sulfuric acid and again mix by inversion.

4. Place tube in a large beaker of water at room temperature
and heat the water quickly to 90° for 5 min. Cool quickly in ice
water for 2 min.

5. Add 0.3 ml. 0.5% phenylhydrazine soln., stopper and shake
vigorously.

6. Set in a water bath at 60° for 5 min. and quickly cool in ice
water.

7. Centrifuge; carefully float a drop of alcohol on the surface
of the liquid, and again centrifuge to throw down floating particles.

8. Carefully pipette 2 ml. clear supernatant into another dry test
tube and cool it in a dry ice-alcohol mixture contained in a beaker
surrounded by an ice-salt bath maintained at — 15 to — 20°.

9. Cool some cone, hydrochloric acid in the Dry Ice-alcohol
mixture, and, after the tube has been cooling for about 10 min., add
1.5 ml. of the cold hydrochloric acid to it.

10. Stopper, and continually invert in the air until the frozen
material melts.

11. Just before the last of the frozen material disappears add 0.2
ml. potassium ferricyanide soln. and mix quickly by several inver-
sions.

12. Set in an ice-salt bath for 5 min. and then place the tube in
a beaker of water at room temperature.

ASCORBIC ACID 245

13. After 10 min., promptly measure the color (it slowly fades
and becomes turbid) spectrophotometrically at 535 m/x. From 0 to
1.5 milligram per cent allantoin there is a linear relationship between
absorption and concentration.

14. Run the standard soln. in the same manner and at the same
time as the unknown.

ASCORBIC ACID*

The method of Roe and Kuether (1943) for assay of ascorbic acid
was adapted by Lowry, Lopez, and Bessey ( 1945) to determinations
on amounts of blood serum down to 10 fd. Measurements in the
range 0.3 to 1.4 milligram per cent have been made with a standard
deviation, in single determinations, of the order of 0.03 milligram
per cent. Ascorbic acid is converted to dehydroascorbic acid, the
latter is treated with 2,4-dinitrophenylhydrazine, and the osazone
formed is made to yield a colored dehydration product through the
action of sulfuric acid. Pijoan and Gerjovich ( 1946) pointed out that,
while this method is reliable for use on blood, its application to
tissues must be made with caution in regard to oxidation products of
ascorbic acid. This follows since the phenylhydrazine reaction is not
specific for dehydroascorbic acid but can react with structures, such
as diketogulonic acid, which bear no antiscorbutic properties. Before
applying the procedures to tissues, it would be desirable, and perhaps
necessary, to ascertain in advance whether interfering substances
were present in the tissue. A titrimetric method, employing dichloro-
phenol indophenol, which measures ascorbic acid directly and not
dehydroascorbic acid, is given on page 300.

Lowry, I^opez, and Bessey Method for Ascorbic Acid

SPECIAL REAGENTS

Osazone Reagent. Prepare a soln. of 2% dinitrophenylhydrazine
and 0.25% thiourea in 9 N sulfuric acid; centrifuge or filter
through sintered glass if a precipitate develops. Store in a re-
frigerator and discard after 1 month.

65% Sulfuric Acid. Add 70 ml. of the concentrated acid to 30 ml.
water.

* See Bibliography Appendix, Refs. 33, 34, and 43.

246 CU\^TTE COLORIMETRY

1% Suspension of Norit in 5% Trichloroacetic Acid. First treat
the Norit by placing 200 g. in a large flask, add 1 1. 10% hydro-
chloric acid, heat to boiling, filter with suction, transfer the cake
of Norit to a large beaker and add 1 1. distilled water; stir well,
filter, and repeat the whole procedure until the washings give a
negative or very faint test for ferric ion. Dry overnight at 110-
120°. Some grades of activated carbon may not require washing;
this can be determined by running a blank test on trichoroacetic
acid washings of the carbon. If these give no more color than the
acid alone, the washing of the carbon is unnecessary. Suspend 5
g. of the iron-free Norit in 100 ml. 5% trichloroacetic acid. After
the Norit has settled, decant the supernatant and restore the
volume with 5% trichloroacetic acid. Repeat several times to
eliminate some of the very fine floating carbon particles. It is
necessary to prevent carbon from getting into the final sample
since carbon contamination may result in low values. If difficulty
from floating is encountered, add 1 vol. 2% gelatin to 10 vol. of
the acid suspension just before use. Once a week or so, replace
the supernatant by fresh acid to avoid the possibility of contamin-
ation with heavy metals that may slowly leach out of the Norit.

PROCEDURE

1. Place 10 /xl. serum in the bottom of a serological tube (6 X
50 mm.) ; add 40 jxl. of the acid-charcoal suspension, and mix by
tapping the tube. In pipetting the charcoal suspension, first blow
through the pipette to suspend the material and then fill and empty
it rapidly to prevent the charcoal from settling out. Employ a con-
striction pipette with a tip and constriction two to three times wider
than normal to avoid plugging.

2. Cap the tube with a piece of Parafilm or a stopper and centri-
fuge 10 min. at 3000 R.P.M.

3. Transfer 30 fA. of the supernatant to another serological tube;
add 10 fx\. of the osazone reagent, and mix by tapping.

4. Cap the tube and set aside at 38° for 3 hr.

5. Chill the tube in ice water and add 50 fil. ice-cold 65% sul-
furic acid. Mix very well, and, after 30 min. at room temperature,
measure the color intensity at 520 m/x using the 0.05 ml. cuvette with
a Beckman spectrophotometer (page 216). If the vol. is increased
the 0.2 ml. cuvette may be used.

V

ASCORBIC ACID AND GLYCOGEN 247

6. Prepare a standard and blank by adding 4 ml. of the acid-
charcoal suspension to 1 ml. aliquots of freshly prepared 1 milligram
per cent ascorbic acid soln. and water, respectively. Centrifuge and
treat 30 /xl. aliquots in the same fashion as the unknowns. Take care
to avoid floating charcoal, which is more of a problem in the absence
of serum. Correct both standard and unknown for the blank and cal-
culate the result; only the single standard is required since the color
is directly proportional to the concentration of ascorbic acid.

note: Serum may be stored safely for several days in a refrigerator or
for several weeks at — 20° after the acid has been added. The supernatant
acid extract may be separated and stored safely in a refrigerator for at
least several weeks, and presumably at —20° for an indefinite period. The
tubes must be well sealed with rubber stoppers to prevent evaporation.
When samples are stored in the frozen state it is preferable to separate
the supernatant before freezing in order to avoid the troublesome tendency
for the charcoal to float as a result of the subsequent necessity for stirring.
The rapid loss of ascorbic acid in blood at ordinary temperatures makes it
imperative to keep the material cool until it is acidified.

In the preceding method of Lowry et al. the danger of charring
has been minimized by the use of 65% sulfuric acid instead of the
85% acid used in the original method of Roe and Keuther; however,
it is still necessary to cool the reaction mixture. Bolomey and Kem-
merer (1946) suggest the substitution of glacial acetic acid for the
sulfuric acid in order to avoid the danger of charring without the
necessity of cooling. The color intensity developed with the acetic
acid is about half that obtained with the sulfuric acid, which may or
may not be important.

GLYCOGEN

While colorimetric methods for glycogen have not been developed
specifically for histochemical studies, the procedure of Boettiger
( 1946) can be adapted to the micro scale necessary.

In the method of Boettiger, the glycogen obtained by alcohol pre-
cipitation of an alkaline digest of the tissue is dissolved, heated with
an acid solution of diphenylamine, and the color which is developed
is measured (a filter No. 635 is used with the Evelyn photoelectric
colorimeter) . This method enables duplicate determinations on 5-10
jug. glycogen, and if reduced to the volumes required in micro-

248 CUVETTE COLORIMETRY

cuvettes, will enable the corresponding refinement. The chief diffi-
culty is the erratic behavior of the diphenylamine reagent, which
necessitates a new calibration each time the determinations are car-
ried out.

The method of van Wagtendonk et al. ( 1946j is based on the color
development which occurs when Lugol solution is added to the gly-
cogen isolated from the tissue. A Klett-Summerson photoelectric
colorimeter was used with filter No. 54 and the measurements were
made in the range 0.05-2 mg. glycogen in a total volume of 5 ml.
Morris (1946) has pointed out that the color developed with iodine
varies considerably with temperature, and therefore temperature
control is required for accurate work. The concentration of iodine
is also a factor that affects the color intensity, and the importance
was stressed for standardization with glycogen obtained from the
same source as the material to be analyzed. Morris is of the opinion
that the precautions required to render the iodine method sufficiently
accurate constitute a major disadvantage. Nevertheless, if tem-
perature is controlled and care is taken to maintain a constant iodine
concentration the method should yield reproducible results. If, in
addition, the standard solution is prepared from glycogen native to
the tissue to be analyzed, sufficient accuracy should be obtained.

For work on the histochemical level, the tissue may be digested
with alkali and the glycogen precipitated with alcohol according to
steps 1-7 in Heatley's procedm'e (page 299). The glycogen thus iso-
lated can be treated in the manner used by Boettiger or van Wag-
tendonk et al. Some preliminary work will be required, no doubt, to
obtain the proper color intensities for the apparatus employed.

Boettiger Method for Glycogen

SPECIAL REAGENTS

Diphenyla7nine Reagent. The purest diphenylamine must be used.
Oxidized crystals are brownish and impart a blue color to the re-
agent. The compound can be purified by dissolving in alcohol at
55°, and crystallizing out by cooling and adding a little water. The
product is dried and stored in a glass-stoppered dark bottle in a
cool place. Glassware used for the reagent must be free of all
organic matter, hence it must be cleaned without soap and kept
dust-free. Add 100 ml. of glacial acetic acid to 3 g. diphenylamine,

GLYCOGEN 249

and, when completely dissolved, add 60 ml. cone, hydrochloric
acid, with stirring. Store in a glass-stoppered dark bottle in a cool
place, and discard when the reagent begins to acquire a bluish
color. Add a few crystals of sodium hyposulfite to improve the
keeping quality.
Standard Glycogen Solutions. Prepare from glycogen reprecipi-
tated from aqueous soln. by alcohol.

PROCEDURE

1. To the glycogen which has been centrifuged down and washed,
add water to make a solution of suitable concentration — this will
have to be determined for the particular case.

2. Add 5 vol. diphenylamine reagent to 2 vol. glycogen soln.
Mix and centrifuge to eliminate any insoluble material. Avoid get-
ting the reagent on the sides of the tube where it will evaporate
during the heating and leave a film that will not go into solution.

3. Heat the tubes exactly 40 min. in boiling water and plunge
into cold water for at least 3 min.

4. Run glycogen standards parallel with the unknowns.

5. Read color intensities (filter No. 635 with the Evelyn colorim-
eter) within 1 hr. after removal from the bath, using a water blank,
and obtain the values of the unknowns from a calibration curve de-
rived from the standards.

Van Wagtendonk, Simonsen, and Hackett Method for Glycogen

SPECIAL REAGENTS

Lugol Solution. Dissolve 1 g. iodine in a soln. containing 2 g. potas-
sium iodide in 20 ml. water. Store in a well-stoppered dark bottle.

Standard Glycogen Solution. Dissolve 25 mg. glycogen {Eastman
Kodak Co., White Label) in 25 ml. 35% potassium hydroxide.
(See note below.)

PROCEDURE

1. To a given vol. glycogen soln., diluted to an appropriate de-
gree as determined in advance, add 0.01 vol. Lugol soln., and mix
well.

2. Read the color at once (filter No. 54 with the Klett-Summer-
son colorimeter) using a blank consisting of the Lugol soln. diluted
100 times with water.

250 CUVETTE COLORIMETRY

3. Obtain the results from a calibration curve derived by using
various amounts of the standard glycogen soln. Subject the glycogen
standards to the same steps of initial precipitation and color develop-
ment as the unknowns.

note: Constancy of temperature and iodine concentration are essential,
and the glycogen standard should be prepared from glycogen obtained from
the same source as the sample to be analyzed. See page 248.

VITAMIN A AND CAROTENE

By means of 2 mm. quartz microcuvettes and an adapter for the
Beckman spectrophotometer (page 217), which was used for the ab-
sorption measurements, Bessey et al. (1946) succeeded in determin-
ing the vitamin A and carotene in as little as 35 fA. blood serum.
Various volumes of serum greater than 35 /xl. may be used as long as
proportional volumes of the reagents are also employed. The pro-
cedure to be described is based on the use of 60 ix\. The method in-
volves saponification and extraction with solvents of low volatility,
measurement of the absorption by the small volumes at 328 m^u ( for
vitamin A) and 460 m/^ (for carotene), destruction of the vitamin A
absorption by treatment with ultraviolet irradiation, and finally re-
measurement of the absorption of 328 m/x. By measurement of the
absorption of 328 m/x, before and after the destruction of the vitamin
A, absorption at this wavelength due to other substances will not
interfere with the determination.

Method of Bessey et al. for Vitamin A and Carotene

SPECIAL APPARATUS

Ultraviolet Apparatus. A diagram of the apparatus used for the
ultraviolet irradiation is shown in Figure 81. A mercury discharge
lamp (General Electric B-H4) with purple emelope-filter and
transformer are used to furnish the radiation. The brightest part
of the lamp is placed opposite to the lower half of the tube, and
the shadow of the electrode support is not allowed to fall on any
tube. The tubes must be cooled during the irradiation by a moder-
ate air current from a fan.

Mixing Apparatus. The device for mixing the liquids in the narrow
tubes is made by cutting off the head of an eight-penny nail,

VITAMIN A AND CAROTENE

251

slightly flattening the end for a distance of 10-15 mm., inserting
the nail in a small high-speed hand drill with the end projecting
about 20 mm., and mounting the drill vertically with the nail up.
The liquids are mixed by touching the side of the tube near the
bottom to the rapidly rotating nail. If this apparatus is not avail-
able mixing may be effected by adding a 1 cm. length of stainless
steel wire (0.041 in. diameter — may be obtained from Newark
Wire Cloth Co.) and shaking. For the extraction with the kero-
sene-xylol, the open ends of the tubes are sealed in a flame and
then shaken vigorously. Care is taken to prevent contamination
of the tops of the tubes by serum which would be charred when
the tubes are sealed.

Fig. 81. Arrangement for ultra-
violet irradiation. Mercury lamp (A)
is held vertically in a clamp, base up,
with the other end extending 3 or 4
cm. into a hole 8 cm. in diameter,
B-B, in a large block of wood, C,
which serves as a base. Semicircular
racks {D, D') are provided for hold-
ing the glass tubes in a circle equi-
distant from the lamp (6 cm. from the
center of the lamp). These racks may
be made from pieces of quarter inch
plywood held about 2 cm. apart, with
the upper piece drilled to hold the
tubes. Twenty or thirty holes may be
drilled in each rack along a semicir-
cular line. From Bessey et al. (1946)

' A

^

1

1
1

1

1

-t

D'

u

1 c

B

SPECIAL REAGENTS

1 N Potassium Hydroxide in 90% Alcohol. Add 1 vol. 11 A^ potas-
sium hydroxide to 10 vol. absolute alcohol. Prepare on the day it
is used. If the color develops rapidly or if the reagent gives a blank,
reflux the alcohol with potassium hydroxide and distill.

252 CUVETTE COLORIMETRY

Kerosene-Xylol. Mix equal vol. xylol, C. P., and water white odor-
less kerosene (obtainable from Einier and A7nend).
Anhydrous Propionic Acid.

PROCEDURE

1. Place 60 fxl. serum and 60 [A. alcoholic potassium hydroxide
in a test tube 100 X 3 mm. (Prepare the tubes by cutting 200 mm.
lengths of glass tubing, 3-3.5 mm. internal diameter, cleaning with
boiling half -cone, nitric acid, rinsing, drying, and dividing in the
middle with a blast lamp flame to yield two tubes.) If the liquids do
not run to the bottom, send them down with a whipping motion.

2. Mix the liquids, immerse the tube in a 60° water bath for 20
min., cool, and add 60 /A. kerosene-xylol.

3. Extract by holding the tube at a 45° angle against the whirl-
ing nail so that the contents are violently agitated for 10-15 sec.
Then, when the tube is at room temperature or a little below, centri-
fuge for 10 min. at 3000 R.P.M.

4. Cut the tube with a file just above the kerosene-xylol layer
and pipette this layer into the cuvette with a constriction pipette,
taking care to avoid the inclusion of any of the aqueous liquid which
would cause turbidity. (Use an uncalibrated 50-60 fA. constriction
pipette (page 172) with a fine tip and a fine constriction because of
the low surface tension of the organic solvents.)

5. Read the absorption at 460 and 328 m/x, and then transfer the
liquid to a soft glass test tube 40 mm. long and 2.5-3.0 mm. internal
diameter.

6. Irradiate with ultraviolet for 30-60 min. (Find the proper
time by testing with known vitamin A solns. The time should be six
to eight times that required to destroy half the vitamin in pure soln. )

7. Transfer the liquid back into a cuvette and take a second
reading at 328 m/A. Rinse the pipette with anhydrous propionic acid
before the transfer to eliminate traces of moisture which would cause
turbidity.

CALCULATION

^460 X 480 = microgram per cent carotene
(^328 — -2^328 after irradiation) X 637 := microgram per cent

vitamin A

VITAMIN A AND CAROTENE 253

where E — optical density for 1 cm. cuvette = 2 — log per cent
transmission with 1 cm. cuvette. The factor 637 is based on an E
(1%, 1 cm.) of 1720 for vitamin A palmitate in alcohol at 328 m/x,
calculated as free alcohol. The factor 480 is based on an E (1%,
1 cm.) of 2080 for /3-carotene (Sniaco) in kerosene-xylol.

///. TITRIMETRIC TECHNIQUES

A. MICROLITER BURETTES

The microliter burettes employed in histochemical procedures
fall into two general groups. In one a capillary glass tube is
calibrated so that the volume of liquid delivered can be determined
by observing the position of a meniscus. These burettes are usually
modifications of the Brandt-Rehberg (1925) instrument, which is
arranged to move the column of solution by the pressure of a
mercury thread controlled by a screw. Mercury in a reservoir is
displaced by turning in the screw and the displaced mercury moves
into the glass capillary. Instruments of this general type have been
described by Pincussen ( 1927) , Linderstr0m-Lang and Holter
(1931, 1933a), Kirk (1933), Sisco, Cunningham, and Kirk (1941),
Links (1934), and Boell (1945). In the Heatley (1935, 1939)
microburettes the pressure is supplied by leveling-bulb arrange-
ments, and both Conway (1934) and Hawes and Skavinski (1942)
employ hydrostatic pressure in their instruments. The Conway
burette was modified by Ramsay ( 1944) for use under anaerobic
conditions (page 279).

In the other general group of burettes a calibrated capillary tube is
not used, but the screw, usually in the form of a micrometer, is cali-
brated instead. These are essentially modifications of the instrument
described by Widmark and Orskov ( 1928) . Krogh and Keys ( 1931) ,
Kirk (1933), and Krogh (1935) employed a fine screw to move the
plunger of a small glass syringe for the accurate delivery of small
volumes of liquid (page 174). Trevan (1925), Dean and Fetcher
(1942), and Hadfield (1942) used the spindle of a micrometer to
operate the plunger. Probably the best micrometer burette is that
designed by Scholander ( 1942) and later improved by Scholander,
Edwards, and Irving (1943). In this instrument the spindle of the
micrometer is used to displace the mercury in the reservoir. An

255

256

TITRIMETRIC METHODS

advantage of this group of microburettes is that their accuracy is
independent of the lumen of the capillary.

Linderstr^ni-Lang and Holler Burettes. These instruments
possess an approximately fivefold refinement of the original Brandt-
Rehberg ( 1925) instrument, and they have been constructed in two
main forms. The type 1 burette ( Linderstr0m-Lang and Holter,
1931), shown in Figure 82, has a cahbrated glass capillary tube

Fig. 82. Burette, type 1.

From Linderstr0m-Lang

and Holter (1931)

Fig. 83. Burette, type 2.

From hinder At rOm-Lang

and Holter (1933a)

58 cm. long, having a total capacity of 100 ix\. and graduated in
divisions of 0.2 ix\. Estimations may be made to 0.02 /xl. When the
screw in the bottom is turned in, the mercury is forced up into the
capillary, which, in turn, forces the liquid out of the burette. The
tip of the burette is dipped into the liquid to be titrated in order that
quantities less than a drop may be added. Readings are taken from
the meniscus of the top of the mercury column. In filling the burette
the tip is dipped into the standard solution and the screw is reversed.
The top of the mercury column is in contact with the standard
solution.

MICROLITER BURETTES

257

In the type 2 (Lindersti'0m-Lang and Holter, 1933a), shown in
Figure 83, the mercury is separated from the standard solution by
an air space. This instrument is used for solutions which might be
affected by contact with mercury. When the screw S is manipulated,
the right mercury column, which is open to the air, is raised or
lowered, and this results in a small positive or negative pressure over
the left column. In this way liquid can be delivered from, or drawn
into, the burette.

The type 1 burette can be connected to a permanent reservoir of
standard solution as shown in Figure 84 ( Linderstr0m-Lang and
Holter, 1933b).

^

L^

8. Burette, type
with reservoir.
Frovi hinder str0m-Lang
and Holter (1933b)

Fig. 85. Glass bead used

to exclude air during titration.

From hinder sir 0m-Lang, Weil

and Holter (1935)

Reduction of evaporation and protection from the air during
titration is afforded by the loosely fitting glass cap around the tip
of the burette held suspended by two threads (Fig. 82). This effect
may also be obtained by passing the tip of the burette through a
glass bead, P (Fig. 85) , which rests on the top of the titration vessel
(Linderstr0m-Lang, Weil, and Holter, 1935). The glass bead may
be dipped into paraffin oil first in order to effect a better seal to the
titration tube. In order to carry out titrations in an atmosphere free

258

TITRIMETRIC METHODS

from carbon dioxide, Schmidt-Nielsen (1942) designed a soda lime
container that fits on the titration tube as shown in Figure 86.

It may be observed in Figures 64 and 84 that a titration table
is employed which is adjustable both vertically and horizontally.
An opal-glass background, illuminated by a small electric bulb, is

Reservoir

Fig. 86. Titration with "desiccator" to maintain
carbon-dioxide-free atmosphere.
From Schmidt-Nielsen (1942)

provided to facilitate the observation of the color of the solution.
The electromagnet placed to the left of the titration table is used
for magnetic stirring in the manner described in the section dealing
with stirring devices (page 179).

MICROLITER BURETTES

259

Tlie complete titration assembly is available from A. H. Thomas
Co. and E. Petersen, Carlsberg Laboratory.

Fig. 87. Burette, front and rear views:
From Sisco, Cunninghavi, and Kirk (1941)

Kirk Burette. In this instrument (Fig. 87) the mercury is
separated from the standard solution by air, and the readings are
taken from a scale behind the capillary tube rather than from

260

TITRIMETRIC METHODS

graduations on the tube itself. The burette has a total capacity of
about 0.1 ml. and is capable of a precision in reading of ±0.03 /tl.
(Sisco, Cunningham, and Kirk, 1941). (Available from Micro-
chemical Specialties Co.)

Heatley Burette. The essential differences between the Heatley
( 1935) burette and the preceding models are that the mercury
displacement is effected by means of a leveling bulb rather than a
screw, the standard solution is in contact with paraffin oil, and
delivery is made directly from a stock bottle of the solution. A
diagram of the instrument is shown in Figure 88. The tube leading

100 B 0

I I I I I I— I I I I I

TIT

Fig. 88. Burette.
From Heatley (1935)

from the delivery tip extends to the bottom of the stock bottle which
has a 2 oz. capacity and is lined with paraffin. The stopper of the
stock bottle is a cork infiltrated with paraffin, and the 5 mm. vertical
tube H ends flush with the bottom of the cork. The upper part of H,
the capillary connection {M) , the three-way stopcock {E), reservoir
(F), and the space (D) are all filled with paraffin oil. The space
under D and part of the capillary tube {B) contain mercury. The
leveling bulb arrangement (C), by which the pressure is regulated,
also contains mercury. An air space exists between the leveling
bulbs and the mercury thread in the capillary. No air is permitted
between the mercury under D and the delivery tip. Interchangeable
delivery tips fitted through a ground-glass joint may be used. When
not in use, the tip is dipped into a tube of the titration solution

MICROLITER BURETTES

261

covered by a layer of paraffin oil. This serves to protect the tip,
and if the protecting tube and the tip are sealed together by a rubber
connection, siphoning-over of the solution is prevented. By adjusting
C so that only a small positive pressure is applied, the surface
tension at the fine tip will prevent liquid from escaping, and delivery
will occur only when the tip is immersed in the solution to be titrated.
This, of course, is the principle used for the other microburettes
that have been described. Stock bottles can be interchanged without
affecting the capillary in any way. One of the instruments that
Heatley constructed had a capacity of 0.1 ml. over a 25 cm. scale,
and the capillary was divided in 1.0 [x\. graduations. The relatively
large volume of the stock bottle and the air space between the
mercury in the leveling bulb and that in the capillary would make
this burette particularly prone to errors arising from temperature
fluctuations during titration.

Fig. 89. Micrometer burette.
From Scholander, Edwards, and Irving (1943)

Boell Burette. One of the easiest burettes to construct is that
of Boell (1945). The instrument consists of a horizontal calibrated
capillary tube drawn out to a fine delivery tip at one end, bent
vertically and pointing downward; the other end of the capillary is

262 TITRIMETRIC METHODS

connected to a rubber tube that acts as a mercury reservoir. One end
of the rubber tube is plugged to retain the mercury, and a screw
clamp is used to force the mercury out of, or to draw it into, the
reservoir. An air space is left between the mercury and the titration
liquid. While lacking some of the refinements of other models, it
can be made into a serviceable instrument.

Scholander Burette. A diagram of this instrument is shown in
Figure 89; an all-steel micrometer, with its anvil removed, is used.
A burette (.4) for titration, or a burette (B) for calibration of
pipettes and syringes may be fitted by a ground-glass joint to the
chamber containing the micrometer spindle. The volume of the
bulbs in the burettes should approximate the volume that the spindle
can displace. A medium-heavy grease is applied to the micrometer
spindle, and the spindle chamber is fixed in position by means of a
set screw (3) which presses against the steel disc (1) having a
recessed punch mark; a lightly greased paper or fiber gasket (2)
seals the open ground end of the chamber held against the spindle
bearing. The gasket should fit the spindle tightly. With the spindle
retracted until flush with the bearing face, the chamber is filled with
mercury through the open ground socket, taking care to remove all
air bubbles. The air bubbles adhering to the walls can be removed
by touching the bubbles with the end of a fine steel wire and leading
them out. Bubbles at the ground socket are avoided by placing
several drops of water or titration liquid over the mercury in the
socket before inserting the upper part of the burette. Extra mercury
can be drawn in through the tip, if necessary. The spindle chamber
should be made as small as required to just clear the spindle. By
keeping the chamber volume small, and with proper handling of the
instrument, temperature errors can be reduced to the point where a
water j acket is not necessary.

The micrometer employed has a total spindle excursion of 25 mm.
marked off in 2500 scale divisions. Estimations are made to one fifth
of a division. Calibration may be carried out by weighing delivered
quantities of water, and relating their volumes to the number of
scale divisions required to deliver them. The total capacity of the
burette can be delivered with an accuracy of 1 part in 6000 to 7000.
With the ordinary spindle, volumes can be measured with an over-all
accuracy of about 0.1 /xl. By replacing the spindle with a Vie in. drill
rod, a refined burette can be constructed capable of measuring

MICROLITER BURETTES 263

delivered amounts with an accuracy of about 0.02 fA. The burettes
are commercially available from Emil Greiner Co. and from 0.
Hebel, Edward Martin Biological Laboratory, Swarthmore College.
Loscalzo and Benedetti-Pichler Burette. Loscalzo and Bene-
detti-Pichler (1945) described the titration of microgram samples
in volumes of 0.05-0.50 fil. using a burette (Fig. 90) consisting of

TP ST TR ^^Li n

Ref.

HW

^ T

L V_

SH

STB

TV

TP

MS

H

Fig. 90. Titration of microgram samples. Upper drawing shows burette with
remote control, 7X natural size. Center drawing shows open titration cone,
7X natural size. At bottom, burette, schematic: TP, tipi; ST, shaft; TR, taper;
SK, shank; Ref., reference mark; SH, shank of titration cone; TV, titration
cone proper; STB, shaft of burette; HW, meniscus of hydraulic water; MS,
of standard solution; A. air column; H, pipette holder of metal with rubber
washer (horizontal shading. From Loscalzo and Benedetti-Pichler (1945)

a piece of capillary tubing drawn out to a fine tip. The titrations are
carried out in a moist chamber on the stage of a low-power micro-
scope and the operations are controlled by a micro manipulator.
Measurements of the quantity of the solution delivered from the
burette are made by observing the displacements of the meniscus
on the scale of an eyepiece micrometer used with the microscope.
The solution is moved by the pressure transmitted from a leveling-
bulb arrangement (Fig. 91). The method of handling the small
volumes of liquid was developed by Benedetti-Pichler (1937) for
micro qualitative analysis, and it was expanded later by Benedetti-
Pichler and Rachele (1940), and Benedetti-Pichler and Cefola
( 1942) . Details of the apparatus and manipulations will not be given
since there have been no histochemical applications of titrations in
volumes of 0.05-0.50 /i.1. However, the possibilities in this respect are
somewhat intriguing.

264

TITRIMETRIC METHODS

Fig. 91. Leveling bulbs for use with micropipettes.
From Loscalzo and Benedetti-Pichler (1945)

Housing for cuvette holder

Cuvette holder, 4 x l4 cm

Electromagnetic stirrer

Micro cell, 8 mm. outside diameter

Photo cell

Light
source

Path of
ight beam

Spring clip to press
cuvette into place

Electromagnet
with protruding core

Fig. 92. Detailed diagram of microcuvette in holder,

showing relationship of electromagnetic stirrer to light beam.

From Zamecnik, Lavin, and Bergmann (1945)

PHOTOMETRIC END POINTS AND METHODS 265

B. PHOTOMETRIC END POINTS

For the objective determination of end points in micro titrations,
Zamecnik, Lavin, and Bergmann (1945) employed a photoelectric
apparatus. A Pfaltz and Bauer fluorophotometer (model A) was
used, and the titration vessel was a square, 6X6 mm. ungraduated
hemometer tube 3 cm. long. The tube was placed in an adapter
which fitted into the cuvette holder and the titration was carried out
with the vessel in the instrument (Fig. 92). Electromagnetic stirring
was employed as shown. The end point was indicated by the galva-
nometer reading which corresponded to the maximum rate of de-
crease in light transmission per microliter of standard titration
solution.

The apparatus was applied to the Linderstr0m-Lang acetone
titration of amino groups using naphthyl red as the indicator. A
Wratten filter 77 was used in order to obtain light in the region of
540-550 uifji for the photometry. Actually, the acetone titration can
be performed satisfactorily by visually matching the color to the
arbitrary end point color chosen for the blank (page 304), but there
could be instances in which it would be of advantage to have an
objectively fixed end point.

C. METHODS

SODIUM AND POTASSIUM (COMBINED)

A method has been described by Linderstr0m-Lang (1936) for
the estimation of sodium plus potassium in small samples of bio-
logical material containing less than 4 X 10"^ milliequivalent of the
alkalis with a precision of 1 X 10 ^^ milliequivalent. The determina-
tion involves ashing the sample with a reagent to convert the
sodium and potassium to chlorides, removal of other chlorides, and
electrometric titration of the residual chloride with silver nitrate.
During the removal of extraneous chlorides by heating, a slight loss
of sodium and potassium chloride occurs even when the temperature
is held as low as 330°, the sublimation temperature of ammonium
chloride. However, this loss is fairly reproducible and it is usually

266 TITRIMETRIC METHODS

sufficient to compensate for it by multiplying the result by the
factor, 1.04.

From the results of various tests, it was shown that the ashing
reagent removes all sulfate and phosphate; the presence of calcium,
magnesium, and iron in the sample does not disturb the determina-
tion.

Linflerstr0ni-Lang Method for Sodiiiin and Potassium
SPECIAL APPARATUS

Quartz Tubes. These are of clear quartz having an inner diameter
of 3.8 mm., an outer diameter of 6 mm., and a length of 20 mm.
The cleaning of the tubes requires a special procedure in this case:
Completely fill with dil. (1 iV) hydrochloric acid (to dissolve
barium carbonate). Wash out and fill with 2 N ammonia (to
dissolve silver chloride). Wash out, fill, and heat with cone,
sulfuric acid (to dissolve barium sulfate), and finally wash several
times with boiling water. In all operations, avoid touching the
rim of the tubes with the fingers.

Incineration Oven. The oven, shown in Figure 65, is made of a
solid copper block containing holes 18 mm. deep and about 7 mm.
in diameter. Two electric heaters placed at the sides of the block
enable a temperature of 440-460° to be maintained. A rheostat
is used to obtain lower temperatures and to regulate the heating.
The sides and bottom of the oven are insulated with asbestos.

Hand Pipette for Ashing Reagent (about 10 fil. capacity). Made
of a piece of capillary tubing with a rather wide aperture to avoid
plugging b}^ barium carbonate, which forms during pipetting.

SPECIAL REAGENTS

Ashing B.eagent. Prepare a soln. containing 1.2% barium chloride

(BaClo.2H20) and 3.2% barium hydroxide (Ba(OH)o.8H20).
Precipitation Reagent. Dissolve 5.7 g. ammonium carbonate in

100 ml. 1.5 N ammonium hydroxide.
Titration Medium. Prepare a soln. containing 1.20% sodium

nitrate and 0.40% secondary sodium phosphate (Na2HP04.2H20)

in 0.10 N nitric acid.
Silver Nitrate Solution (0.02 N). Dissolve 0.3398 g. silver nitrate

in 100 ml. titration medium.

SODIUM AND POTASSIUM ( COMBINED) 267

PROCEDURE

I. Transfer sample of tissue containing 1 X 10"^, to 4 X 10~^
milliequivalent of the alkalis into the bottom of a quartz tube. If
necessary, about 8 ix\. water can be placed in the tube to receive the
tissue.

2. Add about 10 ix\. ashing reagent with the simple hand pipette
and place in an oven at 106° to evaporate the water.

3. When dry, place in the incineration oven at 440-460° for 30
min. The grayish-white ash may contain dark particles of carbon,
but these do not influence the analysis.

4. Add 100 ^1. distilled water to the ash with a calibrated con-
striction pipette, and stir carefully with a thin glass rod ( 1-2 mm.
diameter.

5. Cap the tube and centrifuge for 5 min. at 2000 R.P.M.

6. Draw off 70-80 fA. of the supernatant with a calibrated
constriction pipette and transfer to a glass reaction tube of the
usual type (page 166).

7. Add 15 ju.1. precipitation reagent with another standardized
constriction pipette and introduce a stirring "flea."

8. Cap the tube and let stand 2 hr., stirring the mixture from time
to time; then remove the "flea" with the electromagnet.

9. Centrifuge as before and transfer 70-80 lA. supernatant to a
quartz tube using the same pipette as previously.

10. Evaporate the water at 106° and incinerate for 1 hr. at 360°.

II. Add 50 ix\. titration medium and titrate the chloride with the
silver nitrate soln. using the electrometric procedure (page 282).

12. Run control experiments, omitting the sample, to obtain a
correction for alkalis in the reagents; the titration value of these
control runs should not exceed 0.2 /xl. silver nitrate soln.

13. Standardize the silver nitrate soln. by carrying a known
quantity of pure potassium chloride through the steps in the
procedure.

note: If in the above procedure it is assumed that pipettes having a
capacity of 101.1, 73.0, and 15.2 n\., respectively, were employed the calcula-
tion becomes:

101.1 X 88.2

Original content = titration value X

73.0 X 73.0

268 TITRIMETRIC METHODS

POTASSIUM

Norberg (1937) developed a method for the determination of
less than 1 X 10"^ milliequivalent of potassium ( < 3.91 fig.) with
a precision of about 1 to 2 X 10 *" milliequivalent in samples of
biological material containing between 1 X 10"^ and 3.5 X 10"'*
milliequivalent. The method is based on precipitation, as the
chloroplatinate, of the potassium extracted from incinerated bio-
logical material, isolation of the precipitate by centrifugation, con-
version of the chloroplatinate to the iodoplatinate, and titration of
the latter with thiosulfate. Tests revealed that the presence of
sodium in concentrations 150 times that of the potassium had no
influence.

A method described by Cunningham, Kirk, and Brooks (1941)
is suited to the analysis of biological material when the ratio of
sodium to potassium does not exceed twenty. For quantities of
potassium over 2 ^g. the error does not exceed 0.5%, and over 0.7 //.g.
it does not exceed 3%. As in the Norberg method, the potassium
extracted from the incinerated sample is precipitated as the chloro-
platinate. Cunningham et al. collect the precipitate on a filter stick,
dissolve the substance, reduce it with sodium formate, and titrate
the chloride thus produced electrometrically.

Only the Norberg method will be described, since it has the
advantage of employing a simple indicator titration rather than
the electrometric procedure, and the separation of the chloroplatinate
by centrifugation would also appear to be a little simpler than
filtration on a specially constructed filter stick.

Norberg Method for Potassium

SPECIAL APPARATUS

Apparatus jor Ashing the Sample. That employed in the combined

sodium and potassium method (page 266) of Linderstr0m-Lang

is used.
Glass Precipitation Tubes. These may be either of the forms shown

in Figure 36 (page 167).
Equi'pment lor Removal of Supernatant Fluid over the Precipitate.

Illustrated in Figure 37 (page 167). Suction is applied at A

POTASSIUM 269

through a rubber tube. The low-power microscope aids in the
careful removal of the liquid.

SPECIAL REAGENTS

Ashing Reagent. 1.2% barium chloride (BaCl2.2H20) soln, con-
taining 3.2% barium hydroxide (Ba(OH)2.8HoO).

Precipitatioji Reagent. Prepare 0.02 M chloroplatinic acid by dis-
solving 0.1 g. of the substance (H2PtCl6.4H20) in distilled water
and making up to 10 ml. The soln. is stable for at least a month
if stored in a refrigerator.

Pure Absolute Alcohol.

Phosphate Buffer (pH 6.98). Combine 4 ml. M/15 primary potas-
sium phosphate, 6 ml. M/15 secondary sodium phosphate, and
90 ml, distilled water.

2 N Potassium Iodide. Dissolve 3.32 g. potassium iodide in water
and make up to 10 ml. Filter if necessary. The soln. must contain
no free iodine.

0.02 N Sodium Thiosulfate (0.496% NaoS203.5H20). Standardize
preferably by treating 7-8 fA. 0.01 N potassium chloride in the
same fashion as the soln. of the unknown.

Green-Light Filter Solution. Add 4 ml. M/15 phosphate buffer,

pH 6.36 (7.5 ml. primary potassium phosphate -f 2.5 ml. secondary
sodium phosphate) to 6 ml. 0.04% bromothymol blue. Do not use
the soln. longer than a week, since the color eventually changes.

PROCEDURE

1. Ash the sample as described in the combined sodium and
potassium method of Linderstr0m-Lang (page 267) and extract the
ash with 100 /xl. water in the same manner.

2. Pipette an aliquot of about 100 /A. of the centrifuged clear
extract containing 1 X 10"^ to 3.5 X 10"^ milliequivalent potassium
into a precipitation tube and add 10 ix\. precipitation reagent with
a hand pipette.

3. Evaporate the water at 90-95°; it does not matter if the
heating continues for 1-2 hr. after the water has been driven off.

4. Wash the residue with 100 /A. absolute alcohol, added with a
hand pipette. Use a platinum needle 0.8 mm. thick to stir up the
solid and rinse the needle with 10 /xl. absolute alcohol before remov-
ing it from the tube.

270 TITRIMETRIC METHODS

5. Cap the tube with a short piece of clean rubber tubing plugged
at one end with a glass ball. The rubber should have no rough
surfaces which might rub off and contaminate the precipitate with
particles. Centrifuge for 5 min. at 2000 R.P.M. (radius 14 cm.).

6. Draw off the supernatant liquid with the capillary tube
arrangement leaving less than 5 ju.1. covering the bottom of the tul)e
to a depth of about 1 mm.

7. Repeat the washing process and dry the precipitate by heating
at 95° for 30 min.

8. Add 45 [A. boiling hot phosphate buffer, pH 6.98, with a hand
pipette to dissolve the precipitate. Cap the tube and set aside for
6-12 hr. to insure complete soln.

9. Add 15 ^1. 2 N potassium iodide with a hand pipette, introduce
a stirring "flea," cap the tube, and mix the contents.

10. After 30 min. and up to 24 hr., titrate the liquid with 0.02 N
thiosulfate to the disappearance of the rose color. Carry out the
titration in the green light obtained by passing the light from an
electric bulb fixed on the back of the titration stand through a cell
filled with the green-light filter soln. Compare the end point color
to that of the tube of water placed beside the titration tube.

SODIUM

Lindner and Kirk ( 1938) described a method for the determina-
tion of sodium which can be applied to small samples containing
0.13-4.13 /xg. sodium. The standard error, on the average, is reported
as a few tenths of a per cent. The method depends on the precipita-
tion of sodium as sodium zinc uranyl acetate, isolation of the pre-
cipitate, reduction of the uranium with cadmium, and titration of
the reduced uranium with eerie sulfate. The procedure involves
several quantitative transfers, not of aliquots, but of total material.
Hence scrupulous care must be exercised to avoid loss at each of
these steps (see page 165).

The procedure of Clark et at. (1942) might be adapted to the re-
quired micro level for histochemical work. In their procedure, the
sodium zinc uranyl acetate is simply dissolved and titrated with
sodium hydroxide using phenolphthalein as indicator. Each sodium
atom in the sodium zinc uranyl acetate is equivalent to nine of the
alkali molecules.

SODIUM 271

Sodium may also be determined by difference using both the Lind-
erstr0m-Lang method for sodium plus potassium (page 266) and the
Norberg method for potassium alone (page 268). A colorimetric
method for sodium is given on page 203.

Lindner and Kirk Method for Sodium

SPECIAL REAGENTS

Calcium Hydroxide.

0.1 N Hydrochloric Acid.

Zinc Uranyl Acetate Solution. Prepare as described on page 203
and let stand 24 hr. Filter through a sintered-glass bacteria-proof
filter. Should turbidity appear, refilter.

Wash Solution. Saturate 95% alcohol with sodium zinc uranyl
acetate and filter absolutely clear as above.

Asbestos. Grind washed and ignited Italian asbestos in a mortar
until fine. Boil in successive portions of eerie sulfate soln. acidi-
fied with sulfuric acid. Wash well with water and store in an all-
glass container.

5% Sulfuric Acid. Prepare from acid redistilled in a glass ap-
paratus.

0.01 N Ceric Sulfate Solution. Standardize against pure sodium
oxalate or potassium ferrocyanide.

Indicator Solution. 0.0025 M phenanthroline ferrous sulfate or
0.1% setopaline C (see page 275).

Standard Sodium Chlojide Solutions.

Spirals of Cadmium. Cut a thin ribbon from a stick of the metal
with a lathe tool. Flatten the helix formed to a disc^having a total
surface of about 2 cm.^; leave one end extending upward to serve
as a handle. This reductor may be used almost indefinitely.

Water. Redistill all water used from an all-glass apparatus.

PROCEDURE

1. Incinerate the sample at about 450° for several hr. in a small
platinum crucible.

2. Add about 50 /nl. 0.1 A'' hydrochloric acid to the cooled clean
white ash.

3. Transfer with washing by means of a large capillary pipette
to a centrifuge cone of about 0.2 ml. capacity. Add a little calcium
hydroxide with a toothpick and mix well to precipitate phosphate.

272 TITRIMETRIC METHODS

4. After standing 2-4 lir., add a drop of water and centrifuge for
at least 5 min. at about 5000 R.P.M.

5. Transfer the supernatant quantitatively, with washing of the
precipitate, to a porcelain titration dish (Fig. 44, page 169) using
a capillary pipette.

6. Evaporate to a small vol. just short of dryness, and while
still warm, add 1 ml. zinc uranyl acetate reagent. Stir and warm for
about 5 min.

7. Cover with a Petri dish and let stand 12-24 hr. for complete
crystallization. If reagent crystallizes during this period, place a
container of water under the Petri dish to prevent evaporation from
the sample.

8. Suck off the supernatant through a sintered-glass filter stick
covered with fine asbestos (Fig. 61). Wash the porcelain dish and
precipitate thoroughly with the alcoholic wash soln.

9. Remove the asbestos to the porcelain dish with a drop of 5%
sulfuric acid, which is also used to wash the end of the filter. Rinse
with a little water.

10. Insert the cadmium spiral, warm and stir for 5 min.; then
remove and thoroughly wash the cadmium with water.

11. After cooling for 5 min., titrate the reduced uranium with
eerie sulfate using one of the indicators.

12. Run a blank determination on the reagents by repeating the
entire procedure without a sodium sample.

CALCIUM

Two methods for the microestimation of calcium have been given
by Siwe (1935b) and Lindner and Kirk (1937a) have described still
another. All the methods employ precipitation of calcium as the
oxalate. In one of Siwe's methods the precipitate is dissolved in acid,
an excess of permanganate is added followed by potassium iodide,
and the iodine liberated by the excess permanganate is titrated with
thiosulfate. In the other method the oxalate is converted to carbon-
ate by heating, an excess of acid is added, and the excess titrated
with alkali. The Lindner and Kirk procedure uses an excess of eerie
sulfate to react with the oxalate dissolved in acid, followed by titra-
tion of the excess with ferrous ammonium sulfate. Kirk and Tomp-
kins (1941) compared oxalate titrations by different methods and

CALCIUM 273

found that the eerie sulfate procedure was superior to the direct
permanganate titration when the latter is used with setopaline C to
sharpen the end point. However, no comparison was made with the
iodometric permanganate method as used by Siwe. Siwe employed
his methods for measurements of the calcium in 50 ju,l. samples of
serum and reported a precision of ±3.5% for both procedures.
Lindner and Kirk stated that their method was adapted to the
determination of 0.5-12 /xg. calcium with a precision of less than
d=0.5% on all but the lowest amounts of calcium.

The procedure followed by Sobel and Kaye ( 1940) , which substi-
tutes an iodometric for the acidimetric titration of excess acid in the
Siwe method, could be adapted to histochemical work. This involves
ignition of the calcium oxalate precipitate to convert it to the car-
bonate, solution of the carbonate in an excess of acid, and iodometric
determination of the excess acid by addition of an excess of potas-
sium iodate and potassium iodide, followed by titration of the liber-
ated iodine with thiosulfate. By employing very dilute thiosulfate
(0.0007 N) as little as 4 /xg. calcium could be measured using a 5 ml.
burette for the titration. By carrying out the oxalate precipitation
at pH 3.0 to 3.5, the authors minimized coprecipitation of magne-
sium. Conversion of the oxalate to carbonate was considered advan-
tageous since the oxalate precipitate could then be washed with am-
monium oxalate to reduce the loss during the washing, and any excess
of oxalate ion in the precipitate would be volatilized in the ignition.
See page 219 for a colorimetric method.

Siwe Iodometric Permanganate Method for Calcium

SPECIAL REAGENTS

Ammonium Hydroxide (cone).

5% Ammonium Oxalate.

25% Nitric Acid.

1% Potassium Iodide.

0.2% Soluble Starch.

0.01 N Potassium Permanganate.

0.01 N Sodium Thiosulfate.

PROCEDURE

1. Place a soln. of the sample having a vol. of about 50 jA. into

274 TITRIMETRIC METHODS

a reaction tube, add a very small drop of ammonium hydroxide, and
drop in a mixing "flea."

2. Warm to about 50-60°, mix well, and add 50 ix\. ammonium
oxalate soln. Let stand 2-6 hr.

3. Centrifuge, remove the supernatant with a pipette, and wash
the precipitate twice with 100 lA. portions of ice-cold water.

4. Centrifuge 15-30 min. and pipette off the supernatant.

5. Dissolve the precipitate in 25 fxl. of the nitric acid by stirring
and heating for 30 sec. in boiling water.

6. Add 50 /xl. permanganate soln., and after 1-1.5 min. add 25 /xl.
iodide soln.

7. Titrate the iodine liberated with thiosulfate until the yellow
color is almost gone. Then add a very small amount of starch soln.
and continue the titration until the blue color just disappears.

8. Run a control by titrating as above but omitting oxalate from
the mixture.

NOTE : It has been the experience of the Carlsberg Laboratory workers that
a serious loss of iodine can occur by evaporation in iodometiic methods
unless a Hquid seal is placed across the lumen of the reaction tube. There-
fore it would be advisable to follow the technique given on page 311,
steps 4-5.

Siwe Acidimetric Method for Calcium

SPECIAL REAGENTS

Ammonium Hydroxide (cone).
5% Ammonium Oxalate.
0.01 N Hydrochloric Acid.
0.01 N Sodium Hydroxide.
1% Phenolphthalein.

PROCEDURE

1-4. Same as steps in the preceding permanganate method. Use
a heat-resistant glass reaction tube.

5. Heat on a sand bath or in an oven at 550-600°, not exceeding
the latter, for 30 min.

6. After cooling, pipette in 50 /xl. 0.01 N hydrochloric acid and
heat for 30 sec. in boiling water.

7. Add a very small amount of phenolphthalein and titrate with
0.01 A'' sodium hydroxide to the first pink color.

CALCIUM 275

8. Run a control by titrating 50 jul. of the acid with the alkali to
the same end point.

note: The precautions given on page 257 for the exclusion of atmospheric
carbon dioxide during the titration should be followed. It would also be
advantageous to substitute tetramethylammonium hydroxide for the
sodium hydroxide in order to avoid the difficulties occasioned by the use
of the latter in fine-bore microburettes; thymol blue will serve as a
better indicator than phenolphthalein (page 291).

Lindner and Kirk Cerinietric Method for Calcium

SPECIAL REAGENTS

Water. Distill all water used for reagents or otherwise from an all-
glass apparatus, and likewise distill all sulfuric acid used.

4% Ammonium Oxalate. Filter through fine asbestos on a sintered-
glass filter.

Wash Solution. Saturate 10% (by vol.) ammonium hydroxide with
freshly precipitated and washed calcium oxalate. Filter through
asbestos on a sintered-glass filter. Prepare fresh from time to time
to avoid the formation on long standing of a very fine suspension
that cannot be properly filtered.

Saturated Sodium Acetate.

0.01 N Ceric Sulfate.

0.01 N Ferrous Ammonium, Sulfate in 0.1 N Sulfuric Acid. Store in
a dark bottle under illuminating gas and withdraw portions as
needed. The soln. is stable in contact with air for 2 weeks or longer.

0.0025 M Phenanthroline Ferrous Sulfate. Prepare from a stock
soln. of 0.025 AI which is made up by dissolving 1.485 g. o-phen-
anthroline in 100 ml. of 0.025 M ferrous sulfate.

The advantage of using nitro-o-phenanthroline ferrous sulfate has been
emphasized by Salomon et al. (1946). A 0.05% soln. of lisamine green {Bntish
Drug House Ltd.) gives a smaller blank than the o-phenanthroline ferrous
sulfate, according to Nimmo-Smith (1946), who also recommends the latter
reagent because it is cheaper.

0.1 N Hydrochloric Acid.

6 N Sulfuric Acid.

Asbestos. Treat all asbestos used as follows: Wash and ignite
Italian medium fiber asbestos, and grind in a mortar. Boil in acidi-
fied ceric sulfate soln. and wash well with water on a sintered
glass filter. Store a suspension of the material in water.

276 TITRIMETRIC METHODS

PROCEDURE

1. If sample must be ashed, place in a small platinum crucible
and heat in a micro furnace not exceeding 450°. Dissolve ash in a
drop of 0.1 A'^ hydrochloric acid and transfer the liquid to a porcelain
titration dish (Fig. 44, page 169) using a pipette. Rinse crucible and
pipette with water to make the transfer quantitative (see page 165) .

2. Warm the acidified soln. of the sample in the titration dish, in-
sert a thread stirrer, and add, with mixing, a vol. of 4% oxalate
soln. equal to that of the sample.

3. Add 0.1 ml. saturated sodium acetate and continue heating for
5 min., adding a little water to keep up the vol.

4. Let stand overnight protected from dust.

5. Filter the calcium oxalate with a micro external filter stick
(1.5 mm, diameter, and covered with a thin layer of asbestos). Use
about 30 drops of wash soln. to rinse the dish and the precipitate on
the filter (see page 177).

6. Transfer asbestos pad to titration dish by using a drop of 6 A^"
sulfuric acid. Wash the filter with a few drops of the sulfuric acid
followed by a few drops of water.

7. Heat the dish for 2 min. while stirring to dissolve the oxalate,
and then let cool for 5 min.

8. Add a measured excess of standard eerie sulfate, stir for 3 min.
and add 5 /xl. of the phenanthroline indicator. Titrate the excess with
ferrous ammonium sulfate.

9. Run a control on the reagents alone.

IRON

Kirk and Bentley ( 1936) developed a micro method for the esti-
mation of iron which involves the reduction of the iron by cadmium
amalgam, addition of a measured excess of eerie sulfate, and titra-
tion with ferrous ammonium sulfate using phenanthroline ferrous
sulfate as the indicator. The method is adapted to the measurement
of 2-15 fig. iron. With blood or simple solutions a mean error o(
1-2%, and a maximum error of 4%, has been reported.

Ramsay (1944) reported that in his laboratory all reductions
with pure metals and amalgams gave blanks amounting to several
micrograms of iron. Accordingly, he employed a titanometric method.
Small amounts of copper have been found to interfere with the

IRON 277

analysis, but Ramsay succeeded in circumventing this difficulty
by a selective extraction of the iron. As described, the method may
be used for the determination of 10-200 ng. iron. When applied to
analysis of 0.2 ml. portions of blood, the coefficient of variation was
0.45%. A bother characteristic of the titanometric method is that
frequent standardization is required and the titration must be carried
out in a manner which will prevent access of oxygen to the solution;
furthermore, the standard solution must be protected from oxygen
at all times. Ramsay employed a horizontal burette arranged as
shown in Figure 93. The standard solution is kept in contact with
an atmosphere of hydrogen, and a stream of carbon dioxide bubbles
is used for stirring.

With the development of microcuvettes, the application of the
colornnetric methods for iron, e.g., the a,a'-dipyridyl or thiocyanate
reactions, to the small quantities considered in histochemical work
may be expected to offer more sensitive analyses, and possibly by
simpler jiroccdures.

Kirk and Bentley Method for Iron

SPECIAL REAGENTS

5% Sulfuric Acid (iron free).

3% Cadmium Amalgam.. Dissolve 3 g. pure cadmium in 100 g.

mercury and store in a well-stoppered, deep, narrow vessel to

minimize air oxidation.
0.01 N Ceric Sidfate.

0.01 N Ferrous Ammonium Sulfate. Store as indicated on page 275.
0.025 M Phenarifhroline Ferrous Sulfate. Dilute ten times before

using (page 275).

PROCEDURE

1. If the sample requires ashing, place in a platinum crucible (8
mm. deep, 12 mm. top diameter) and heat for 10 min. just below
visible redness. The small muffle furnace described on page 180 is
very well suited to tne purpose. Heating at too high a temperature or
for too long may cause formation of insoluble ferric oxide. The iron
concentration in blood is so high that a five fold dilution is necessary
to obtain a convenient sample (3-15 /^g. iron) . Addition of some iron-
free sodium hypochlorite soln. to the water used in the dilution
serves to dissolve clot fragments and assist in the ashing. Drive off

278 TITRIMETRIC METHODS

the water by heating on a hot plate before proceeding to the ashing.

2. Dissolve the ash in 5% sulfuric acid, and, if the insoluble form
of ferric oxide is present, remove the acid extract and heat the moist
residue on a hot plate until sulfur trioxide fumes appear. Then dis-
solve the residue in 5% sulfuric acid and add the liquid to the first
extract.

3. Pipette soln. of sample into a microreductor (Fig. 48, page
171) and adjust acidity so that when diluted to vol. the acid con-
centration will be 0.1 A^. Usually 40-75 /xl. 5% sulfuric acid is re-
quired for solns. of ashed samples.

4. After the vol. is made up to the mark, add 30 ul. 3% cadmium
amalgam; stopper and shake at intervals or continuously for 10 min.

5. Pipette about twice the vol. of 0.01 N eerie sulfate needed to
react with the unknown into a dish (Fig. 44, page 169). Add a meas-
ured quantity of indicator soln. and start the thread stirrer.

6. Open the reductor and pipette an aliquot into the eerie sulfate
soln.

7. Titrate with ferrous ammonium sulfate to a permanent reddish
tint.

8. Run a control by titrating the eerie sulfate without a sample.

Ramsay Method for Iron

SPECIAL REAGENTS

Concentrated Sulfuric, Perchloric (60%), and Nitric Acids. Test
the nitric acid for iron and distill from glass, if necessary.

50% Potassium Thiocyanate.

0.02 to 0.05 N Titanous Sulfate (depending on the quantity of iron
and the bore of the burette). Pour 200 ml. freshly boiled distilled
water into bottle F (Fig. 93) with stopcocks C, D, and G closed.
Mix the proper amount of com. 15% titanous sulfate or chloride
with 10 ml. water and 10 ml. cone, sulfuric acid. Boil the mixture
vigorously for 1 rain. Pour at once into F; stopper immediately.
Open stopcock D and let stand overnight for any precipitate
to settle. Then close D, open C, and allow a brisk stream of hy-
drogen to bubble through .4. Close C, and open first D and then G
to force the standard soln. over the siphon into the burette. A soft
vacuum grease (W. Edwards and Co.) was used for the stopcocks,
but perhaps silicone grease would be better. Standardize daily
against a standard ferric ammonium sulfate soln.

IRON

279

Standard Ferric Ammonium Sidfate. Prepare a soln. in 3 A?" sulfuric
acid such that 5 ml. requires a vol. of titanous sulfate equivalent
to 100-200 mm. on the burette.

Sodium Chloride (Powdered).

50% Formic Acid in Ether. Redistill each component from glass,
and mix just before using.

Fig. 93. Modified Conway burette. A, pressure-regulating vessel; B, C, D,
G, and H, stopcocks; E, the burette, standard-bore capillary tubing against
mm. scale (10 cm. o 0.05 ml.); F, Ti2(S04)3 reservoir, capacity 200-250 ml.;
/, digestion and titration flask. From Ramsey (1944)

PROCEDURE

1. Add the sample (0.2 ml. blood) to 0.2 ml. each of the nitric
and perchloric acids and 0.5 ml. sulfuric acid in a digestion flask.
Add a boiling chip and heat with shaking over a micro flame. The
mixture darkens, then clears, and just before the last of the perchloric
acid boils off the mixture becomes a clear deep greenish-yellow for
3-5 sec. Continue heating for 3 sec. after the disappearance of this
color.

2. If copper is present, dilute the cooled digest with 2 ml. water
and transfer as completely as possible to a narrow test tube. Rinse
the flask with two portions of 1 ml. water and add the rinsings to the
test tube. Add 0.5 ml. sulfuric acid and saturate the soln. with sodium
chloride. Shake the tube twice with 2 ml., and twice with 1 ml. 5%

280 TITRIMETRIC METHODS

formic acid in ether. Transfer the ether extracts back to the digestion
flask and completely evaporate the ether over warm water. Digest
again after the addition of 0.5 ml. sulfuric acid and a drop or two
each of the nitric and perchloric acids. Cool the clear digest and add
5 ml. water.

If too little copper is present to interfere, omit this step. (The
separation of copper is not necessary for blood analysis.)

3. Add 1 ml. 50% potassium thiocyanate to the diluted digest,
bubble carbon dioxide through the soln., open stopcock H, and deliver
the titanous sulfate until the last trace of pink just disappears. To
control the rate of delivery adjust the height of the liquid column in
the tube leading from A to B, and carry out the last of the titration
with C closed and B open.

PHOSPHORUS

A method for the measurement of phosphorus in the range of 0.5-
10.0 jxg. was given by Lindner and Kirk (1937b). The method con-
sists of conversion of the phosphorus to phosphate by ashing, precipi-
tation as ammonium phosphomolybdate, isolation of the precipitate
by filtration through a sintered-glass filter stick with an asbestos
mat, solution of the precipitate in a measured excess of sodium hy-
droxide, addition of an equivalent amount of acid, and finally titra-
tion of the excess acid with alkali to a phenolphthalein end point.
The end point is not sharp and the precision of the method leaves
much to be desired.

In an early review, Glick ( 1935a) reported that, with Linderstr0m-
Lang and Holter, he had developed a method for inorganic phosphate
capable of determining 5 /xg. phosphorus with an error of less than
1%, and 1 )ug. with an error of about 5.5%. In this method the phos-
phate was precipitated as magnesium ammonium phosphate, the pre-
cipitate was centrifuged, washed with alcohol and then acetone,
and dissolved in an excess of standard hydrochloric acid. The excess
acid was titrated in an acetone medium with ammonium acetate
using naphthyl red indicator. Phosphoric acid does not ionize in ace-
tone ; therefore only the free hydrochloric acid is titrated. The acetate
ion is used like an alkali in this case since it will react with hydrogen
ions to form acetic acid which, in acetone, does not ionize.

Both of these titrimetric methods lack the ease and simplicity

PHOSPHORUS AND CHLORIDE 281

characterizing the colorimetric procedures (page 226), and, since
the former methods do not offer sufficient compensating advantages
in accuracy, in the opinion of the writer, the colorimetric methods
deserve preference.

CHLORIDE

In chronological order, the following methods have appeared for
the determination of chloride in very small amounts of biological
material.

Linderstr0m-Lang, Palmer, and Holter ( 1935) described an elec-
trometric titration method that can be carried out with a precision of
0.02 ju,l. 0.02 A^ silver nitrate. The sample for analysis should contain
preferably 0.4-2.0 fig. chlorine. For use on tissue, a preliminary
ashing procedure was employed which could be conducted in the
titration tube itself to avoid transfer.

Conway (1935) employed a method based on oxidation of the
chloride to chlorine by an acid permanganate reagent, absorption
of the chlorine by potassium iodide solution, and estimation of the
iodine liberated by either titration or colorimetry, depending on the
quantity present. The reaction is made to take place in a diffusion
cell. Conway reported that, for quantities of chloride corresponding
to about 0.3 mg. chlorine, the coefficient of variation for a single
determination is 0.5%. Down to 7 ju,g. chlorine the coefficient of
variation increases to 4-5%, and when the method is adapted to
0.7 fxg. the coefficient of variation becomes 6—7%.

Wigglesworth ( 1937) used a method based on addition of excess
silver nitrate, filtration, and back-titration with thiocyanate. The
method is applicable to as little as 0.3 jul. tissue fluid and the
precision is ±: 6%. Dean (1941) modified the Wigglesworth method
and carried out the reaction and titration in a drop placed on a
waxed slide.

Cunningham, Kirk, and Brooks (1941) described an electrometric
titration method similar to that of Linderstr0m-Lang, Palmer, and
Holter (1935). A shallow dish is employed as the titration vessel
and silver-silver amalgam electrodes, a stirring needle, and the
burette tip all dip into the solution. To determine the end point of
the titration, the volume of standard silver nitrate solution added
is plotted against the resulting E.M.F. values. A sample containing

282 TITRIMETRIC METHODS

0.5-30 /j.g. chloride is used. For more than 2 ^g. the error does not
exceed 0.5%, and for more than 0.5 fig. it does not exceed 2%.

Since there is little difference between the two electrometric
methods, only that of Linderstr0m-Lang, Palmer, and Holter will
be described. It has a simple electrode system (the filled burette
serves as one half of the cell), and the titration can be carried out
in the glass reaction tube used for the ashing of the sample. Cun-
ningham et al. (1941) demonstrated that plasma can be titrated
directly, without ashing, by the electrometric method. Colorimetric
methods are given on pages 200 and 224.

Linderstr^m-Lang, Palmer, and Holter Electrometric Method
for Chloride

SPECIAL APPARATUS

The arrangement of the titration set-up is illustrated in Figure
69. A and A' are silver wires that serve as electrodes. They are
connected to a potentiometer so arranged that the galvanometer
gives no deflection when the potential difference across the electrodes
is -f-210 millivolts, corresponding to a silver concentration in the
titration tube of 0.05 X 10^ M. This concentration would be ob-
tained by adding 0.013 fA. 0.02 A^ silver nitrate to the 50 /xl. liquid
employed in the titration tube. The burette, which is of the type 2
variety (Fig. 83, page 256) can be read to 0.02 [A.

SPECIAL REAGENTS

Ashing Solution. Dissolve 1.2 g. sodium nitrate and 0.4 g. secondary
sodium phosphate (Na2HP04.2H20) in water and make up to
14ml.

0.10 N Nitric Acid. This soln. and the titration soln. below were
changed from 0.17 to 0.10 N nitric acid, as suggested by Linder-
str0m-Lang ( 1936) in a later publication.

Titration Solution. Dissolve 0.3398 g. silver nitrate (0.02 N) in
100 ml. 0.10 N nitric acid containing 1.20 g. sodium nitrate and
0.40 g. secondary sodium phosphate (Na2HP04.2H20).

PROCEDURE

1. Place the tissue sample in a heat-resistant tube (Fig. 35,
page 167) containing 7 [x\. ashing soln. Evaporate the water in an
oven at 106°, and ash the material by careful heating in a flame.

CHLORIDE. NITROGEN AND AMMONIA 283

When a crucible oven is available, first heat to about 300° for 10
min. and then continue to heat over the flame. Employ the coolest
flame that will burn away the brown tar which forms.

2. Add 50 /xl. 0.10 N nitric acid with a hand pipette.

3. Titrate the liquid in the tube with the titration soln. keeping
the galvanometer circuit closed and closing the titration circuit by
bringing the liquid in contact with the burette tip. Observe the
deflection each time the tip touches the liquid and a little of the
titration soln. runs out. Close the titration circuit only momentarily
to prevent polarization. When the deflection changes direction, the
end point has been reached.

NITROGEN AND AMMONIA

A discussion of various methods for the titrimetric determination
of total nitrogen and ammonia has been included in the section
dealing with the colorimetric methods (page 233). Since the method
developed at the Carlsberg Laboratory is the most precise (0.005
ixg. nitrogen) and has had its worth established by the most critical
trial over many years, the procedure finally recommended, and
given by Briiel, Holter, Linderstr0m-Lang, and Rozits ( 1946) , will
be the only one presented. In its present form the range of the method
is 0.1-1 fig. nitrogen, but larger quantities may be determined by
increasing the concentration of the reagents. The digestion of the
sample by the method of these investigators has already been de-
scribed on pages 234-236). To avoid the use of alkaline solutions
in the microburette, standard acid is employed for the titration
after an excess of dilute alkali has been added to the acid used to
absorb the ammonia.

Briiel, Holter, Linderstr^m-Lang, and Rozits Method for
Nitrogen and Ammonia

SPECIAL MATERIALS

Paraffin Block. Smooth the surface of a block of paraffin by scrap-
ing with a knife. Store the block under distilled water, and dry
with filter paper when removed from the water for use.

SPECIAL REAGENTS

18 N Sodium Hydroxide.
0.015 N Sulfuric Acid.

284 TITRIMETRIC METHODS

Alkaline Indicator Solution. Add 0.5 ml. M/15 secondary sodium
phosphate to 2 ml. 40 milligram per cent bromocresol green (pH
4.6) and 2.5 ml. boiled water. Prepare fresh each day and keep
in well-closed vessel.

0.01 N Hydrochloric Acid containing 10 ml. 40 milligram per cent
bromocresol green/100 ml.

Color Standard. Prepare by titrating a blank (45 /xl. water + 7 fi\.
0.015 A'' sulfuric acid + 18 /xl. indicator soln.) with the 0.01 N
hydrochloric acid until the color matches that of the indicator
in citrate buffer at pK 4.6. The slight difference in shade of the
indicator in citrate buffer makes the titsated blank the better
color standard.

PROCEDURE

1. Pipette a drop A of distilled water (about 20 /xl.) onto a clean
paraffin block (Fig. 94).

Fig. 94. Use of the paraffin block
in transfer of the digest.
From Brilel et at. (1946)

Paraffin block

2. Draw up about one third of A into a hand pipette without a
constriction (Fig. 55, page 175) and carefully introduce the stem
of the pipette into the digestion tube until the tip touches the center
of the concave bottom of the tube. See that the pipette stem is
properly centered in the tube so that liquid is not drawn up along
the walls. Warm the tube just before adding the water to prevent
crystallization of potassium sulfate, which might block the pipette.

3. Blow out the water into the digested sample (prepared as
described on pages 234-236) and immediately suck up the mixture
into the pipette. Deposit the liquid as a drop (B) on the paraffin
block.

4. Draw up another third of A into the pipette and use the water
to wash the walls of the digestion tube. Draw the liquid back into

NITROGEN AND AJ^MONIA 285

the pipette. The last of the Hqiiid usually follows the tip of the
pipette as it is withdrawn along the wall of the tube and can be
sucked up near the rim. Add the liquid in the pipette to B.

5. Repeat the rinsing of the tube with the last third of drop A.
With care, the last of A can be removed from the paraffin without
drawing air through the pipette tip. Add the liquid to B.

6. Draw up drop B completely into the pipette and transfer it
to the bottom of the paraffined distillation tube (page 169 and Fig.
35, page 167). It is essential that the pipette does not touch the
side walls of the tube. Accordingly, clamp the pipette vertically
and raise up the tube on a pipetting stand so that the centering
of the pipette stem will be proper. Deposit the liquid in the bottom
of the tube. It is usually permissible to ignore the film of liquid
adhering to the walls of the pipette after it has been emptied.

7. Add 9 (A. 18 N sodium hydroxide directly to the liquid in
the bottom of the paraffined tube with the same type of pipette
used for the previous steps. Again it is essential to avoid contact
between the pipette and the upper walls of the tube. Use the pipette
stand for the operation as in the previous step. Immediately after
addition of the alkali place a seal of 45 fA. water across the neck
of the upper part of the vessel, using a constriction pipette of the
type shown in Figure 58, as described on page 174.

8. Pipette 7 /A. 0.015 iV sulfuric acid into the water seal at
once using a constriction pipette (Fig. 57, page 175) which is
emptied by constant air pressure adjusted so that the pipette will
deliver when the tip touches the water. This arrangement insures
greater accuracy.

9. Stopper the tube with a rubber cap plugged with a drawn-out
piece of glass tubing, rather than a piece of glass rod, to prevent
displacement of the seal (Fig. 46, page 171).

10. Immerse the bottom half of the tube in a thermostat at
40° for 1.5 hr.

11. Add 18 fA. alkaline indicator soln. to the receiving seal by
means of a constriction pipette which is emptied by constant air
pressure.

12. Titrate with the 0.01 A^ hydrochloric acid to the color of
the color standard using a "flea" stirrer. About 1.5 lA. of the acid will
be required to bring the blank without absorbed ammonia to the
end point.

286 TITRI METRIC METHODS

13. Run controls in which the complete analysis is duplicated
without the unknown sample.

UREA

Urea has been determined by the urease method in a number
of laboratories by making use of one or another of the micro
procedures for the estimation of ammonia. In the urease method,
the enzyme is added to decompose the urea, and the ammonia thus
formed is absorbed in acid and measured.

Procedures employing diffusion cells were described by Conway
(1933), Gibbs and Kirk (1934), Borsook (1935), and Kinsey and
Robison ( 1946) . The various forms of diffusion cell used by these
investigators have been described (page 168). The method of
Kinsey and Robison (1946), which can estimate as little as 1 fig.
urea in 4 fA. plasma or aqueous humor with a precision of ±5%,
will be described. The methods which do not utilize the diffusion cell
for the estimation of ammonia can also be applied in the present
case. Thus, the method of Linderstr0m-Lang and Holter ( 1933b)
(page 230) was developed with the application to the measurement
of urea in mind as one of the potentialities. A colorimetric method
is given on page 214.

Kinsey and Robison Method for Urea

SPECIAL REAGENTS

Glycerol-Boric Acid. Add 25 ml. pure glycerol to 100 ml. 25%

glycerol saturated with boric acid.
Urease Extract. Dissolve 1 g. powdered double strength urease

(Squibb) in 100 ml. boiled saturated sodium chloride.
Saturated Sodium Metaborate in Saturated Potassium Chloride

(boiled).
0.002 N Hydrochloric Acid.
Gramercy Universal Indicator (Fisher Scientific Co.) diluted with

15 parts distilled water.
Mineral Oil.

PROCEDURE

1 . Measure the sample into the central container of the diffusion

UREA AND UREASE 287

cell (Fig. 43, page 168). Add 1 drop urease soln., stir with a "flea"
or otherwise, and allow to react for 20 min. at room temperature.

2. Place a few drops of mineral oil in the well, and pipette 5
fx\. glycerol-boric acid into the center of the covering vial.

3. When the digestion is completed, add 1 drop metaborate soln.
to the reaction mixture, and immediately place the covering vial over
the central container. Stir the digestion mixture and allow 60 min.
at 30° for the diffusion.

4. Remove the covering vial, blot off the adhering mineral oil,
add 0.5 ml. diluted indicator, and titrate with 0.002 A'" hydrochloric
acid to the color of a control containing no urea. (The latter should
be yellow; a greenish appearance indicates some ammonia has
been introduced, probably from the sodium chloride used in the
urease soln. or from the potassium chloride-metaborate soln.) Allow
about 1 min. between the addition of the final increment of acid
and the matching of the end point.

note: The colorimetric procedure of Russell (1944) (page 238) may be
substituted for the titration: Wash out the glycerol drop from the vial
with three 0.5 ml. portions of water into colorimeter tubes. Add 0.05 ml.
0.003 M manganese sulfate or chloride to each tube. Chill the tubes and
add 1.0 ml. alkaUne phenol reagent (25% phenol in 2.7 N sodium
hj^droxide) and 0.5 ml. hypochlorite soln. Place the tubes in a boiling
water bath for 5 min., cool, dilute to a convenient vol. and read in a
colorimeter.

UREASE

Procedures for the determination of urea can, of course, be adapted
to the measurement of urease. This was done by Linderstr0m-Lang
and Holter (1940, page 1144), who based the method on their
ammonia determination (page 230). 7 fA. enzyme soln. and 7 fA.
substrate (1.2 g. urea, 50 ml. 0.15 N sodium hydroxide, and 50 ml.
0.3 M primary potassium phosphate, pH 6.8) were employed. The
ammonia was liberated from the digested mixture by drawing into
it 7 /xl. 2 A'" sodium hydroxide, which had previously been placed on
the side of the reaction tube. The distillation of the ammonia and
the titration were then carried out (page 285).

The determination can also be carried out by measuring the am-
monia in an acetone titration according to the procedure used by
Linderstr0m-Lang and S0eberg-Ohlsen (1936). To the reaction

288 TITRIMETRIC METHODS

mixture given above, 150 /xl. acetone containing naphthyl red is
added, and titration with 0.05 A^" alcoholic hydrochloric acid is per-
formed in the usual manner (page 303).

AMIDE, PEPTIDE, AND NITRATE NITROGEN

Borsook and Dubnoff (1939) described methods for amide,. pep-
tide, and nitrate nitrogen which were adapted to 0.1 ml. samples
containing as little as 0.3 jag. nitrogen. These methods all involve
determination of ammonia, a diffusion cell technique, and an electro-
metric titration, a glass electrode being used by the authors.
Whether or not one wishes to follow this particular procedure for
the estimation of ammonia, the initial steps in the methods of Bor-
sook and Dubnoff can still be utilized; the details are given below.

Borsook and Dubnoff Methods for Amide, Peptide,
and Nitrate Nitrogen

Amide Nitrogen

SPECIAL REAGENTS (in addition to those required for determination of
ammonia)

S N Sulfuric Acid.

2.9 N Sodium Hydroxide.

PROCEDURE

1. Pipette 0.2 ml. sample into a Kjeldahl digestion tube. Add
0.1 ml. 3 A^ sulfuric acid, and mix.

2. Cover with a tin-foil cap and place in a boiling water bath
for 3 hr.

3. After the hydrolysis, cool and add 0.1 ml. 2.9 N sodium
hydroxide, mixing well during the addition.

4. Determine the ammonia nitrogen and subtract from this
value the ammonia nitrogen present before hydrolysis as measured
by a separate analysis. The difference is the amide nitrogen.

Nitrate Nitrogen

SPECIAL REAGENTS (in addition to those required for determination of
amide and ammonia nitrogen)

Devarda Alloy (finely powdered).
S5% Sodium Hydroxide.

AMIDE, PEPTIDE, AND NITRATE NITROGEN 289

PROCEDURE

1. To the soln. remaining after all ammonia has been distilled
out, in the determination of amide nitrogen, add 4 mg. Devarda
alloy. Tilt the vessel so that the alloy is separated from the liquid.

2. Then introduce 0.1 ml. 25% sodium hydroxide, and allow the
ammonia formed to diffuse into a fresh drop of standard acid. 1-2
hr. is sufficient for the reduction of the nitrate, and the distillation
of the ammonia will be complete within this period if the diffusion
cell of the authors is employed.

3. Determine the amount of ammonia which was absorbed.

4. Employ two sets of controls, one in which the sodium
hydroxide is added without the alloy, and a second in which the
alloy is added to a control mixture containing the reagents for the
amide determination, but no sample. Substract the sum of these
two controls from the titer obtained in step 3.

Peptide Nitrogen

SPECIAL REAGENTS (in addition to those required for measurement of
amino acid and ammonia)

Peptidase Preparation (ammonia free). Seed skim milk contain-
ing 5% dextrose with spores of Aspergillus wentii and incubate
for 6-8 days at 30°. Culture the mold in 1 1. Erlenmeyer flasks
with a 2 cm. depth of medium. A satisfactory growth is obtained
when the heavy pad of mold has begun to crack and draw away
from the wall of the flask. Remove the pads; weigh, and freeze
for several hr. to inactivate any arginase which may be present.
Grind the pads in a mortar with sand and add enough water to
give a suspension containing about 10% dry matter. (Determine
the moisture on a small sample of the pad.) Bring the pH to 7 to
7.5 and cover the suspension with toluene. After standing 4-6 hr.
at room temperature, filter through sailcloth with the aid of a
vacuum. Add an equal vol. acetone to the filtrate, and dissolve
the precipitate which forms in one fourth the original vol. water.
Centrifuge, and shake the supernatant with permutite to remove
ammonia. Store this enzyme soln. under toluene in a refrigerator.
The activity will be maintained for months even though the soln.
will darken on standing.

PROCEDURE

1. Add an equal vol. of a protein-free extract (trichloroacetic

290 TITRIMETRIC METHODS

acid filtrate), brought to pH 6.0, to the enzyme soln., and incubate
overnight under tokiene.

2. Determine the ammonia and carry out a formol titration.
Muhiply the ammonia found by 1.1 and subtract this value from
the formol titration value at the second stage. From the figure
obtained, subtract the sum of the formol titer of the enzyme soln.
and the free amino nitrogen before the enzymatic hydrolysis to
arrive at the peptide amino nitrogen.

ACID, ALKALI, AMINO, AND CARBOXYL GROUPS

Titration of acid or alkali in the conventional manner with
micro apparatus, using indicators or electrometric means for deter-
mining the end point, requires little discussion. Usually it is
advisable to use a color standard for matching the end point when
indicators are employed. The use of standard acid in capillary
burettes offers no difficulties ; however, standard solutions of metallic
hydroxides often give rise to the complications of carbonate
precipitation in the fine lumen, and for this reason have been
avoided by the Carlsberg Laboratory group. The difficulty has been
circumvented in some cases by the use of tetramethylammonium
hydroxide for the standard solution or by the addition of excess
alkali and back-titration with acid. With alkaline end points, pro-
tection from the carbon dioxide of the air must be afforded. Methods
for accomplishing this have been given on page 257. For electro-
metric titrations, various arrangements may be employed, such as
the metallic wire electrode of Linderstr0m-Lang, Palmer, and Hol-
ter (1935) (Fig. 69, page 184) or the open-cup glass electrode of
Sisco, Cunningham, and Kirk (1941) (Fig. 70).

For the direct raicrotitration of amino groups, Linderstr0m-Lang
and Holter (1932) developed an acidimetric acetone titration
method, which is described in their procedure for proteolytic enzymes
(page 303). Linderstr0m-Lang and Duspiva (1936) employed an
alkalimetric alcohol titration method for the direct micro estimation
of carboxyl groups, and this measurement is described in their
protease method (page 304). Glick (1934) (page 309) determined
carboxyl groups by their neutralization in an excess of alkaline
buffer and acidimetric titration to an acid pH. An indicator formol

LIPID 291

titration was described by Weil (1936) (page 306), and an electro-
metric formol procedure by Borsook and Dubnoff (1939). Sisco,
Cunningham, and Kirk (1941) employed their glass electrode (Fig.
70, page 184 ) for the electrometric titration, and they also used an
indicator method in which the titration is carried out in an open drop
but differing little from the procedure of Weil.

LIPID

A clever microtitration technique for the measurement of lipid in
quantities of the order of 10 fig., suited for the determination on 1
mg. of tissue, with a precision of about 1% was developed by
Schmidt-Nielsen (1942). The method is based on saponification of
the lipid in a sealed capillary tube by alcoholic alkali in the presence
of toluol, liberation of free fatty acid by the addition of an excess
of mineral acid, solution of the fatty acid in the toluol phase, and
titration of an aliquot of the toluol layer after the toluol has been
evaporated.

Schmidt-Nielsen Method for Lipid

SPECIAL REAGENTS

Pure Toluol.

1 N Potassium Hydroxide in Absolute Alcohol (aldehyde free).

0.5 N Hydrochloric Acid.

0.01% Thymol Blue in Absolute Alcohol.

0.02 N Alcoholic Tetramethylammonium Hydroxide.

PROCEDURE

1. Pipette the lipid sample in 20 lA. toluol into the bottom of a
thin-walled capillary tube having an internal diameter of about
1.8 mm. and a length of about 50 mm. Clamp the pipette vertically
and raise the capillary tube, placed in a holder, on a mechanical
stand so that the tip of the pipette will not touch the sides of the
tube as it is raised. Use constriction pipettes for all pipettings.
Exercise care to avoid evaporation of toluol in steps 1-6.

2. In the same way pipette 2.5 p\. alcoholic potassium hydroxide
into the toluol soln. and immediately seal the opening of the capil-
lary.

292 TITRIMETRIC METHODS

3. Set aside overnight at room temperature or heat to over 80°
for 10 min. to effect saponification. Keep the material at the one
end of the tube so that none will be lost when the other end is
subsequently cut off to introduce the next reagent.

4. Centrifuge to drive down the droplets of toluol which con-
dense on the upper walls of the capillary, open the empty end of the
tube with a diamond point, and pipette an excess of 0.5 N hydro-
chloric acid down into the tube ( 13 fA. will suffice in most instances) .

5. Reseal the open end of the capillary at once and thoroughly
mix the contents by shaking. The special centrifuging apparatus
which repeatedly turns the tubes (Fig. 63, page 178) has been
especially designed for mixing liquids in capillary tubes.

6. Separate the toluol and water phases by centrifuging, open the
capillary, and pipette an aliquot of about 13 //,!. of the toluol layer
into a titration vessel with a standard ground mouth (Fig. 86) .

7. Evaporate the toluol in a desiccator furnished with paraffin
chips at about 30 mm. mercury pressure.

8. Dissolve the fatty acid in 100 fA. 0.01% alcoholic thymol
blue, add a stirring "flea," and titrate with the standard tetra-
methylammonium hydroxide using the set-up shown in Figure 86
to exclude atmospheric carbon dioxide. Match the color to that of
a faint green color standard prepared by titrating 100 jxl. alcoholic
thymol blue to a pure yellow color and then adding 5 /xl. 0.04%
bromophenol blue, which has been rendered blue with a little sodium
hydroxide.

9. Run a blank titration on 100 fA. alcoholic thymol blue.

EXTRACTION AND FRACTIONATION OF LIPIDS

To enable the extraction and fractionation of the lipids in quanti-
ties of tissue of the order of 1 mg., so that analytical methods
might be applied to the material, Schmidt-Nielsen (1944b) elabo-
rated appropriate procedures. The tissue lipids are treated with
alkali and the unsaponifiable fraction is extracted with toluol.
Fatty acids are liberated from the aqueous portion by the addition
of mineral acid and then they are extracted with toluol.

LIPID EXTRACTION AND FRACTIONATION 293

Schmidt-Nielsen Method for Extraction
and Fractionation of Lipids

SPECIAL REAGENTS

Pure Toluol.

1 N Potassium Hydroxide in Absolute Alcohol (aldehyde free).

4 N Hydrochloric Acid ( chlorine free) .

PROCEDURE

A. Unsaponifiable Fraction

1. Place the tissue in the bottom of a capillary tube (1.8-2 mm.
internal diameter, 50 mm. long, and sealed at one end). Two micro-
tome sections each 25 /x thick and 5 mm. in diameter weighing
approximately 1 mg. may be used. It is convenient to employ the
cryostat microtoming technique (page 427), in which the sections
are allowed to curl up on the knife edge and are transferred into
the capillary tube kept cold in the cryostat.

2. Pipette 2.5 fA. alcoholic potassium hydroxide into the tube
with the sample and follow with the addition of 20 /xl. toluol. Seal
the open end of the tube at once. Follow pipetting directions on page
291.

3. Saponify by heating the tube to 80-100° for 2 min. Centrifuge
to throw down droplets of toluol that condense along the tube.

4. Open the tube, introduce 16 fA. water, and reseal.

5. Mix the liquids thoroughly by placing in the mixing centri-
fuge (page 178).

6. Separate the emulsion into two layers by centrifuging at
3500 R.P.M. or faster, after heating the capillary tube, the ordinary
centrifuge tube into which it is placed, and the metal centrifuge
tube holder in an oven at 110°. If the separation is incomplete,
reheat and recentrifuge.

7. Open the tube and pipette a 16 fA. aliquot of the toluol phase
into a paraffined reaction tube (page 169). Evaporate off the toluol
in a desiccator over paraffin shavings at a pressure of 20 mm.
mercury.

294 TITRIMETRIC METHODS

B. Saponifiable Fraction

1. Evaporate the toluol left in the aqueous phase from step A7
in the desiccator, as above. Should the iodine number subsequently
be determined on the saponifiable fraction it would be necessary to
correct it for the presence of the remaining unsaponifiable lipid.
This would require that the iodine number of the unsaponifiable
fraction be determined also. It is imjiossible to obtain complete
separation of the two phases by pipetting.

2. Add 2.5 fi 1.4 iV hydrochloric acid and 20 /x toluol, and seal
the capillary tube as before.

3. Mix as in step ^5 and separate the layers by centrifuging
without heating.

4. Remove an aliquot of the toluol phase as in step A7. If the
sample is to be used for titration of fatty acids, transfer the aliquot
to a titration vessel with the standard ground mouth (Fig. 86). If
the iodine number is to be determined, transfer to a paraffined
reaction tube. Evaporate the toluol as in step A7.

IODINE NUMBER OF LIPIDS

Schmidt-Nielsen ( 1944a) employed the principle of Kaufmann
(1926) to develop a method having a precision of about 1% for
the determination of the bromine-combining power of lipids in
samples of the order of 10 fig. Unsaturated bonds in the lipid are
saturated with bromine and the excess bromine is titrated iodometri-
cally. The iodine number can be calculated from the titration value
if the amount of lipid is also measured (page 291). Schmidt-Nielsen's
innovation of using glycol monobutyl ether (butyl Cellosolve)
for the fat solvent has the advantage that the low vapor pressure
of the liquid obviates difficulties that would result in a micro method
from evaporation. As the solvent is also miscible with water, it has
the additional advantage of enabling the titration to be performed
in a single liquid phase.

Kretchmer, Holman, and Burr (1946) employed a similar method
for 10-100 fig. samples of lipid in chloroform solution, using 0.05 A^
pyridine sulfate dibromide in glacial acetic acid as the brominating
agent. At least 20 min. was required for completion of the reaction.

IODINE NUMBER OF LIPIDS 295

Schmidt-Nielsen Method for Iodine Nunil>er

SPECIAL REAGENTS

Cellosolve (glycol monobutyl ether). Redistill and store in the
cold.

1 % Potassium Iodide Solution. Store in the dark.

0.1% Soluble Starch Solution. Stir up 0.1 g. in 10 ml. cold water
and pour, with stirring, into 90 ml. water at 76°. Store at 0°.
Vs or ^/e N Bromine Solution. (May be made weaker.) Saturate
methyl alcohol with sodium bromide (about 13 g./lOO ml.);
filter and add 10% more methyl alcohol. Dissolve the bromine
in this soln. Store in the dark. Transfer some into a bottle with
a small opening to minimize the evaporation of alcohol while being
used.

0.05 N Sodium Thiosidfate. Stabilize by adding 0.2 g. sodium
carbonate per liter.

Pure Toluol.

PROCEDURE

1. Pipette the sample dissolved in toluol into the bottom of a
paraffined reaction tube (page 169).

2. Evaporate the toluol in a desiccator over paraffin shavings at
a pressure of 20 mm. mercury.

3. Introduce a stirring "flea" and pipette 5 /A. Cellosolve into
the bottom of the tube.

4. Place a seal of the potassium iodide soln. (about 45—50 ^1.)
across the lumen of the tube about one third of the tube length above
the bottom, and follow with a similar seal of the starch soln. about
one third from the top. The paraffin coating prevents the seals from
running. Place the seals by holding the tube horizontally. Introduce
the tip of the pipette to the proper position and have it just touch
the wall. Blow out the liquid while rotating the tube. Finally cap
the tube.

5. Use the "flea" to dissolve the lipid in the Cellosolve. Some
creeping of the lipid occurs during the evaporation of toluol and
all this lipid must be incorporated in the Cellosolve. The paraffin
coating is insoluble in Cellosolve.

6. Introduce 2.5 [A. bromine soln. into the bottom of the tube,

296 TITRIMETRIC METHODS

from a pipette fixed vertically, by raising the tube on a mechanical
stand so that the tip and shaft of the pipette pass through the two
seals without touching the sides of the tube at any point.

7. After 5 min. the bromination is complete; then use the "flea"
to draw the potassium iodide seal down into the liquid in the bottom
of the tube and mix them.

8. Raise the tube on a stand so that the tip of the burette
passes through the starch seal and dips into the bottom liquid
without touching the sides of the tube. Add thiosulfate from the
burette until the yellow color just disappears. Remove the tube, and
with a quick snap of the hand throw the starch seal down into the
bottom liquid. Replace the tube in the titration position and con-
tinue the titration to the end point. Use a previously titrated sample
as a colorless control to facilitate judgment of the end point.

9. Run blank titrations on the bromine without a lipid sample.

REDUCING SUGARS

The iodometric method for the estimation of reducing sugars was
adapted by Linderstr0m-Lang and Holter (1933a) to the micro
scale. Their procedure has a precision of about 0.06 lA. of 0.05 N
thiosulfate, which corresponds to 0.25 /xg. glucose. Holter and Doyle
( 1938) introduced a few alterations in concentration and volume of
the reagents and employed a reaction tube with a narrow neck
(Fig. 38, page 167), which enabled an increase in precision to
0.04 lA. of 0.02 N thiosulfate.

Heck, Brown, and Kirk ( 1937) adapted the cerimetric method
to the determination of reducing sugars with an average accuracy
of about ±1%. The cerimetric method was also used by Lewy
(1946) for the determination of glucuronic acid. For a colorimetric
method see page 210.

Holter and Doyle Modification of Linderstr0ni-Lang
and Holter Iodometric Method for Reducing Sugars

SPECIAL REAGENTS

04 M Carbonate Buffer (pH 10.2) .

0.1 M Iodine in Potassium Iodide. Store in a black bottle connected
by a siphon arm to a 6 fA. automatic pipette (Fig. 60, page 175).
1.2 M Sulfuric Acid.

REDUCING SUGARS 297

0.5% Soluble Starch.

0.02 N Sodium Thiosulfate.

PROCEDURE

1. Pipette 7 lA. of the 0-1.5% sugar soln. into the bottom of a
paraffined reaction vessel (page 169).

2. Add a stirring "fiea" and pipette 10 jul. carbonate buffer into
the tube.

3. Mix the buffer and sugar soln. and add 6 /xl. iodine soln. from
the automatic pipette, whose tip is below the surface of the liquid
in the tube to prevent evaporation of iodine.

4. Directly afterward pipette 10 fxl. 1.2 M sulfuric acid into the
tube to form a seal across the narrow neck of the tube, and then
place another seal of about 10 lA. starch soln. above the acid seal
(Fig. 38, page 167). These seals prevent loss of iodine.

5. Stir the reaction mixture and let stand for 20 min.

6. Cap the tube and centrifuge for 1 min. at about 1500 R.P.M.
to collapse the seals and cause them to mix with the reaction liquid.

7. Titrate the iodine with 0.02 A^ thiosulfate, using a micro-
burette in which mercury is not in contact with the thiosulfate.

8. Run a control in which water is substituted for sugar soln.

Heck, Brown, and Kirk Cerimetric Method for Reducing Sugars

SPECIAL REAGENTS

14% Sodium Carbonate. Prepare from anhydrous salt.

1.5% Potassium Ferricyanide.

10% (by vol.) Suljuric Acid.

0.01 N Ceric Sulfate.

0.01 N Ferrous Ammonium Sulfate in 0.1 N Sulfuric Acid. Store

as directed on page 275.
0.0025 M Phenanthroline Ferrous Sulfate. Prepare as described

on page 275.

PROCEDURE

1. Measure soln. of sample containing 1-12 [xg. glucose, or that
quantity of other sugar with equivalent reducing power, into a tube
(3 mm. inside diameter, 35 mm. long) calibrated at 200 ix\.*
* It would be well to simplify this step as indicated on page 165.

298 TITRIMETRIC METHODS

2. Add about 40 fd. carbonate soln. and the same vol. ferricya-
nide soln.

3. Add water to the calibration mark* and place in boiling water
for 5 min.

4. Transfer to a porcelain titration dish with a pipette and use
successive rinsings with water to insure a quantitative transfer.*

5. Add 50 ixl. 10% sulfuric acid and a little phenanthroline
indicator.

6. Titrate with eerie sulfate to the first permanent grass-green
color.

7. Run control by substituting water for the sugar soln. 1 [xg.
glucose requires 2.556 /xl. 0.01 A^ eerie sulfate.

note: For the deproteinization of blood, Heck et al. pipette sample into
calibrated tube having a vol. about ten times that of the sample. The
pipette is then rinsed into the tube,* a vol. of 10% copper sulfate
(CuSOi.SHaO) equal to that of the sample is added, and while stirring
a similar vol. of 10% sodium tungstate in 2% sodium carbonate is added
slowly. The soln. is made up to the calibration mark with water,* stirred,
centrifuged, and the supernatant is drawn off and used.

GLYCOGEN

Heatley ( 1935) described a micro determination of glycogen
based on the isolation of glycogen from tissue, acid hydrolysis, and
estimation of the reducing sugar formed by the method of Linder-
str0m-Lang and Holter (1933a). The procedure of Heatley may be
applied to amounts of tissue of the order of 1 mg. The maximum
error is less than ±2 fig. glycogen corresponding to ±0.3 jul. 0.05 N
thiosulfate. The loss of about 1.5% which occurs during the pre-
cipitation of the glycogen is approximately compensated by stand-
ardizing the reagents through a control estimation of a glycogen
solution of known strength. Subsequently Heatley and Lindahl
(1937) extended the method to include the separation of desmo-
and lyoglycogen, according to the principles given by Willstatter
and Rohdewald (1934), and the measurement of the two forms
individually. For a colorimetric method see page 247.

*It would be well to simplify these steps as indicated on page 165.

GLYCOGEN 299

Heatley Method for Total Glycogen

SPECIAL REAGENTS

All of the reagents used for the determination of reducing sugars

(page 296) are required, in addition to:

30% Potassium Hydroxide.

Absolute Alcohol.

0.6 N Hydrochloric Acid.

0.1 N Hydrochloric Acid.

0.05% Thymol Blue.

1 N Sodium Hydroxide.

Paraffin.

PROCEDURE

1. Place the tissue sample, which may be fixed in absolute
alcohol to stop enzyme action, in about 35 ju,l. 30% potassium hy-
droxide in a reaction tube.

2. Cap the tube and heat in a steam bath for 20 min.

3. With the pipette used for the alkali, add 35 /xl. water and then
105 ju,l. absolute alcohol.

4. Mix well with the aid of a platinum wire.

5. Immerse the tube in a steam bath for only a few sec. to
flocculate the glycogen without having the contents boil out.

6. Centrifuge at 3500 R.P.M. for 15 min. and pipette off and
discard the clear supernatant.

7. Wash the precipitate with about 100 /A. absolute alcohol;
centrifuge and again discard the supernatant.

8. Repeat the washing process once more and remove the last
traces of alcohol by placing the tube in a steam bath for a few min.

9. With about 35 ^1. 0.6 N hydrochloric acid wash down the
deposit on the upper walls of the tube.

10. Add a 20-30 mg. piece of paraffin and place the tube in the
steam bath for 2.5 hr.

11. While the tube is still hot, invert and rotate it to spread an
even film of paraffin over the upper walls of the tube.

12. Add a "flea" and use it to break through the thin film of
paraffin on the surface of the liquid.

300 TITRIMETRIC METHODS

13. Add a little thymol blue soln. and bring to a bluish-grey
color with 1 A'' sodium hydroxide. If the end point is passed, bring
to the proper color with 0.1 N hydrochloric acid.

14. Set up a control by neutralizing 35 fxi. 0.6 N hydrochloric
acid to thymol blue.

15. Determine the glucose by the method of Linderstr0m-Lang
and Holter (page 269).

Hcatley and Lindahl Method for Desmo- and Lyoglycogen

SPECIAL REAGENTS

The same as in the preceding method for total glycogen.

PROCEDURE

1. Place the tissue in a little distilled water in a tared reaction
tube cooled by ice.

2. Plunge the tube into a current of steam, and shake it at
intervals for 10-15 min. while in the steam. A mechanical shaker
is recommended.

3. Centrifuge, and transfer the supernatant to another tube.

4. Add another portion of distilled water to the residue and
again heat and shake as before.

5. Centrifuge, and add the supernatant to that previously
drawn off.

6. Make one or two more extractions in the same way and
evaporate the water from both tubes. Dry the tubes at 100° for
2 hr. and reweigh. The sum of the increases in weight of the tubes
represents the dry weight of the tissue sample taken. The extracted
material contains the desmoglycogen and the material in the super-
natant is the lyoglycogen.

7. Add 35 [A. 30% potassium hydroxide to the material in each
tube and proceed with the glycogen determination as previously
described.

ASCORBIC ACID

The method introduced by Tillmans and associates for the deter-
mination of ascorbic acid by titration with 2,6-dichlorophenoIindo-
phenol was applied by Glick ( 1935b) to measurements on microtome
sections of tissue. This micro method has a reproducibility of ±0.1
/Ltg. ascorbic acid. Since the publication of the procedure, the use of

ASCORBIC ACID 301

metaphosphoric acid has replaced other acids for extraction because
of the greater stability of the vitamin in this medium. Hence, in the
procedure to be given, metaphosphoric acid rather than the acetic
acid originally employed will be indicated. Ponting (1943) found
that oxalic acid may also be used. Ascorbic acid can be titrated with
the dye, and dehydroascorbic acid cannot be measured unless it is
reduced to ascorbic acid by an agent such as hydrogen sulfide. This
reduction is made preferably in a 3% metaphosphoric acid extract
buffered to pH 3.5 to 3.7 with citrate (Bessey, 1938) . In macro work
the hydrogen sulfide is passed through the solution for about 15
min., and after standing at room temperature for 2 hr., the sulfide
is removed by passing wet nitrogen through the liquid for 45-60 min.
An application of the procedure to micro work would require the ex-
perimental determination of the proper time interval for each of these
steps. For the colorimetric determination, see page 245.

Glick Method for Ascorbic Acid

SPECIAL REAGENTS

Saturated Sodium 2,6-Dichlorophenolindophenol Solution (sodium
2,6-dichlorobenzenoneindophenol) . Shake a few mg. with 100 ml.
warm distilled water. Cool, filter, and store in a refrigerator. The
soln. may be used for about 10 days. [A stable soln. prepared in
dioxane was described by Stone ( 1940) : Distill the dioxane, dis-
carding the first and last 10%. Stir a weighed quantity of the dye
into the dioxane (about 0.4% equals 0.01 M) and add glacial
acetic acid to 1% of the volume. Stir well for about 15 min., and
filter through dry No. 42 Whatman filter paper. During titration
the dioxane soln. used should not exceed 10% of the total volume.]

2-3% Metaphosphoric Acid. Store in a refrigerator and prepare
fresh every few days.

Color Standard. Prepare a rose Bengal soln. ( about 1 part per mil-
lion) having a pink tinge when placed in a reaction tube and com-
pared with a similar vessel containing water.

Standard Ascorbic Acid Solution. Prepare a fresh 0.1% soln. of
crystalline L-ascorbic acid in 2-3% metaphosphoric acid.

PROCEDURE

1. Standardize the dye soln. against freshly prepared ascorbic
acid soln. under the titration conditions employed for the unknown.

302 TITRIMETRIC METHODS

2. Place 50 fA. of the metaphosphoric acid soln. in a 250 /xl. tube
and add the sample (care must be exercised to prevent oxidation of
the material prior to extraction with the acid). For work on tissue
sections, the practice has been to freeze the tissue solid immediately
upon removal from the source, and fresh-frozen sections (20-40 /x
thick, 4-5 mm. diameter) are prepared and transferred individually
as they are cut to separate tubes containing the acid extractant. If
titration cannot be made at once, as in the case of serial-section
titrations, the tubes may be kept in a freezing mixture until time for
titration.

3. Titrate with the standardized dye soln. using a microburette
in which the mercury does not come into contact with the dye soln.
The end point is reached when the color of the imknown persisting
for 5 sec. matches that of the color standard placed beside it. Employ
active stirring throughout the titration.

AMYLASE

The Linderstr0m-Lang and Holter (1933a) method (page 296)
for the determination of reducing sugars was used by Linderstr0m-
Lang and Engel ( 1937) for the measurement of amylase activity in
tissue sections of barley. Holter and Doyle (1938) employed the
method for studies of amylase in protozoa.

In the barley work the enzyme was extracted with M/15 phos-
phate buffer (pH 5.3) and 7-8 ix\. extract was added to a similar vol-
ume of substrate solution consisting of 1.5% soluble starch in phos-
phate buffer of the same pH. After an appropriate digestion period
at 40°, the increase in reducing sugar was estimated. In much the
same manner the protozoan enzyme was measured. The phosphate
buffer (pH 6.0) extract was added to a 2% starch substrate con-
taining 2 % sodium chloride.

The relative proportions of enzyme extract and substrate solution,
the length of the digestion period, and the pH of the buffer must be
dictated by the requirements of the particular case.

PROTEOLYTIC ENZYMES

Titrimetric methods for the determination of proteolytic enzymes
in small samples of biological material, such as microtome sections

PROTEOLYTIC ENZYMES 303

of tissue, have been developed by the Carlsberg Laboratory in-
vestigators. Linderstr0m-Lang and Holter (1932) described a
method for dipeptidase based on the acetone titration of amino
groups. AVith DL-alanylglycine as the substrate, the procedure has a
])recision of about 0.08 /A. of 0.05 A^ hydrochloric acid corresponding
to 5.6 X 10"^ mg. amino nitrogen. Holter and Linderstr0m-Lang
( 1932 1 extended the method to include proteases of both the peptic
and catheptic type. The maximum deviation in this case amounts
to about 0.16 ix\. 0.05 N acid or 1.2 X 10"^ mg. amino nitrogen. By
substitution of the appropriate substrate, the acetone titration micro
method has also been used for aminopolypeptidase, carboxypoly-
peptidase and prolinepeptidase by S0eborg-Ohlsen (1941) and
others. Buffers need not be added in these measurements, since the
substrates have sufficient buffer capacity of their own.

The alcohol titration for carboxyl groups was adapted by Linder-
str0m-Lang and Duspiva ( 1936) for the alkalimetric measurement
of protease; their method has a precision of about 0.3 ix\. 0.05 N
alkali. Because of precipitation, solutions more acid than pH 8 can-
not be titrated by this procedure.

Weil ( 1936) described a formol titration for the determination of
tryptic activity on a similar scale, which he reported to be repro-
ducible to ±0.1 fA. 0.05 N alkali.

The dilatometric method for peptidase is described on page 417.

Linderstr^m-Lang and Holter Acidimetric Acetone Method
for Proteolytic Enzymes

SPECIAL REAGENTS

Enzyme Extraction Mediujn. 30 ml. 88% glycerol + 5 ml. M/15
primary potassium phosphate -f- 5 ml. M/15 secondary sodium
phosphate made up with water to 100 ml.

Acetone Containing 2 Milligram Per Cent Naphthyl Red.

0.05 N 90% Alcoholic Hydrochloric Acid.

Substrates. The peptidase substrates are 0.2 M solns. of the racemic
peptides containing sodium hydroxide to give the desired pH, e.g.,
0.036 M alkali gives a pH of 7.4 to 0.2 M alanylglycine. An excep-
tion is leucylglycine which must be used in 0.18 M soln. because
of its sparing solubility.
Dipeptidase. Alanylglycine or leucylglycine.

304 TITRIMETRIC METHODS

Aminopoliipeptidase. Alanylglycylglycine or leucylglycylglycyl-
glycine; also glycylglycyl-D-alanine (Levy and Palmer, 1943).
Carboxypeptidase. Benzoylleucylglycine.
Prolinepeptidase. Glycyl-L-proline.

Cathepsin. 4% edestin in 0.008 A'" hydrochloric acid, pH 4.4.
Pepsin. 4% edestin in 0.056 N hydrochloric acid, pH 2.1.

PROCEDURE

1. Pipette 7/xl. enzyme extraction medium into a 0.25 ml. reaction
tube. Place tissue section in the liquid, and introduce a mixing
"flea."

2. After standing for 1-2 hr. at room temperature for enzyme ex-
traction, 7 ixl. substrate soln. is pipetted in. The liquid is then mixed.

3. Cap the tube and suspend in thermostat for the chosen diges-
tion period.

4. Set up a control experiment by carefully placing the substrate
soln. on the side of the tube as a separate drop not touching the
enzyme in the bottom. Since there is a tendency for the side drop
to run down into the bottom drop, hold the tube in a horizontal
position, cap, and suspend in the thermostat in a horizontal position.

5. Stop the reaction, after removing from the thermostat, by
pipetting in 30 /xl 0.05 N alcoholic acid and mixing. The automatic
pipette shown in Figure 60 (page 175) is especially useful in this step.

6. Add 150 111. acetone-naphthyl red soln. and titrate the yellow
liquid with 0.05 N alcoholic hydrochloric acid to an orange end point.
Place 200 [A. of a standard orange soln. in a reaction tube and mount
this beside the titration tube so that the end points may be brought
to the same color tone.

Linderstr0m-Lang and Duspiva Alkalimelric Alcohol Method
for Proteolytic Enzymes

SPECIAL REAGENTS

Enzyme Extraction Medium. Glycerol-phosphate soln. prepared as

for preceding method.
Absolute Alcohol Containing 0.05% Thymol Blue.
0.05 N 90% Alcoholic Tetramethylammonium Hydroxide.

PROTEOLYTIC ENZYMES

505

End Point Color Standard. Mix methyl green, fuchsin, and picric
acid to obtain the bluish-green color obtained at the titration end
point.

Substrate, (a) Dissolve 12 g. casein in 8 ml. 1 A?" ammonium hy-
droxide in the presence of 0.05 ml. octyl alcohol and make up to
100 ml. with water, (b) Prepare buffer soln. by mixing 2 N am-
monium hydroxide with 2 A'' ammonium chloride in the ratio a/b
given in the following table, (c) Prepare a buffer-pH correction
soln. for the casein by mixing 1 ml. of the chosen buffer soln.
with the corresponding number of milliliters of alkali as in-
dicated in Table VII and make up to 10 ml. with water. The so-
dium hydroxide may be used instead of the ammonium hydroxide
to avoid high concentrations of ammonia, (d) Before the experi-
ment mix 1 ml. of the 12% casein soln. with 1 ml. of the buffer-
pH correction soln.

TABLE VII
Composition of Ammonia Buffers for Various pH Values

pH

a/b

Alkali, ml.

18°

40°

2 N NHiOH

8 N NHiOH

2 N NaOH

8.58

8.0

1/8

0.22

—

0.20

9.19

8.7

1/2

0.48

—

0.32

9.80

9.3

2/1

1.56

—

0.52

10.10

9.6

4/1

3.00

—

0.60

10.40

9.9

8/1

5.96

—

0.66

11.00

10.5

52/1

—

6.17

0.76

PROCEDURE

1-4. These steps are the same as in the preceding method. How-
ever, since the absorption of carbon dioxide by the alkaline reaction
mixture must be avoided, the tubes are capped with soda lime stop-
pers. (Fig. 47, page 171).

5. Stop the reaction by pipetting in 130 [A. alcohol-thymol blue
soln.

6. Titrate to the bluish-green end point with the 0.05 A^ tetra-
methylammonium hydroxide, matching the color to the color stand-
ard. Protect from carbon dioxide during the titration by using the
paraffin-oiled glass bead arrangement (Fig. 85, page 257), or the
soda lime "desiccator" (Fig. 86, page 258).

306 TITRIMETRIC METHODS

Weil Forniol Method for Tryptic Activity

SPECIAL REAGENTS

Enterokinase Solution. Prepared according to Waldschmidt-Leitz,
(1924). Dehydrate and defat scrapings of hog duodenum mucosa
with acetone, acetone-ether, and ether in this order. Grind and
sieve the dried material, and extract it at 30° for 2 hr. with 50 ml.
0.04 iV ammonium hydroxide per gram solid. Centrifuge, and con-
centrate the supernatant by evai)oration at about 35°.

Veronal Buffer (pH 8.4). Add 8.23 ml. M sodium diethylbarbi-
turate to 1.77 ml. 0.1 A'" hydrochloric acid.

Substrate Solution. Bring 4% casein (Hammarsten) to pH 8.4 with
1 N sodium hydroxide.

Formol Solution. Neutralize 4 ml. 40% formaldehyde, to which 2
ml. 0.1% alcoholic phenolphthalein is added, with 0.1 N sodium
hydroxide to the first pink color, and make up to 25 ml. with dis-
tilled water. Prepare fresh before use.

0.05 N^ Tetramethylammonium Hydroxide.

PROCEDURE

1. Place tissue section or enzyme preparation in 7 fA. entero-
kinase soln. in a reaction tube and let stand for 1 hr. at room
temperature.

2. Add 7 ;ul. Veronal buffer and 7 /xl. substrate soln.

3. Close the vessel with a soda lime tube (Fig. 47, page 171) and
place in thermostat for the required digestion period.

4. Set up a control in which the buffer and substrate are placed
on the side of the tube as a drop not touching the enzyme drop in
the bottom of the tube. Place horizontally in thermostat to prevent
the drops from touching.

5. Add 1000 fA. formol soln. and titrate with 0.05 N tetramethyl-
ammonium hydroxide to a marked red color matching against a
standard. Protect from carbon dioxide in the air during the titration
by one of the arrangements shown in Figs. 85 and 86, (pp. 257, 258).

ARGINASE

Two methods for the micro estimation of arginase were reported
by Linderstr0m-Lang, Weil, and Holter ( 1935) . The hydrolysis of
the very strong base arginine yields the fairly strong base ornithine

AfeGlNASE 507

&tid the very weak base urea. By titration to a pH at which the
guanido group of arginine is completely ionized and the e-amino
group of ornithine (and urea) is not ionized, the cleavage of arginine
can be followed as the increase in the quantity of base required to'
bring the reaction mixture to this pH. The increase in the amount
of base will be one equivalent for each mole arginine hydrolyzed.-
This principle was employed by Linderstr0m-Lang et ctl. The propef
pH w^as obtained by titration in acetone-alcohol with thymol blue'
indicator. This method has the advantage of convenience and ra-
pidity; however, as the authors pointed out, it cannot be used if a
large amount of urease is present because the subsequent action of
the enzyme on the urea will increase the titration over the theoretical
value of one equivalent per mole arginine. The urease activity is rela-
tively low at the pH optimum of arginase, but the factor becomes ap-
preciable nevertheless if high concentrations of urease are present.
The titration of the ornithine set free is only about 95% of the theo-
retical, but this can be corrected if necessary by applying the factor
1.05 to the results.

The other method reported is based on the measurement of the
ammonia formed when urease is allowed to act on urea set free by
the action of the arginase. One volume of 0.10 A'' base in this method
corresponds to one volume of 0.05 N base in the acetone-alcohol
titration method.

Linderstr0in-Lang, Weil, and Holier Methods for Arginase

A. Urease Method
SPECIAL REAGENTS

Substrate Solution. (0.1 M arginine, pH 9.5). Dissolve 0.2104 g.

arginine hydrochloride in 1.335 ml. 0.5 N sodium hydroxide and

water to make a final vol. of 10 ml.
Urease-Buffer Solution. Prepare 100 ml. of a soln. containing 7 g.

urease (Squibb) and 35 ml. 0.5 M phosphate buffer, pH 6.8 (mix

equal vol. primary and secondary phosphate) .
40% Sodium Hydroxide.
0.3 N Hydrochloric Acid containing 10 ml. 0.04% bromocresol

purple/50 ml.
0.1 N Sodium Borate.

308 TITRIMETRIC METHODS

PROCEDURE

1. Pipette 7 /xl. enzyme soln. and 7 (A. substrate into the bottom
of the reaction tube, add a "flea," and mix. Close the tube with a
rubber stopper and allow the reaction to proceed for a suitable time
at an appropriate temperature. (Note that the type of tube origi-
nally used. Fig. 40, page 167, and the accompanying technique have
been replaced by the method of Briiel et al., 1946.)

2. Place the vessel in boiling water for about 15 min., cool, add
20 ^1. urease-buffer soln., and stir with the "flea."

3. After allowing the vessel to stand 1 hr. at 20°, measure the
ammonia formed using the method of Briiel et al. ( 1946) (page 283) .

B. Acetone-Alcohol Titration Method

SPECIAL REAGENTS

Substrate Solution. Same as for urease method.

Acetone- Alcohol-Indicator Mixture. Prepare 100 ml. by adding a

mixture of equal vol. acetone and absolute alcohol to 5 ml, of 0.1%

alcoholic thymol blue.
0.05 N Tetramethylammonium Hydroxide in 90% alcohol.
End Point Color Standard. See page 292, step 8.

PROCEDURE

1. Pipette 7 /xl. enzyme soln. and 7 /xl. substrate into the bottom
of a simple 250 [xl. reaction tube (Fig. 35, page 167). Add a "flea,"
stir, stopper with a soda lime tube (Fig. 47, page 171), and allow
the reaction to proceed at an appropriate temperature for a suitable
period.

2. Stop the reaction by adding 150 /xl. acetone-alcohol-indicator
mixture.

3. Titrate with the tetramethylammonium hydroxide to the shade
of the greenish end point color standard. Use one of the devices to ex-
clude carbon dioxide during the titration (Figs. 85, 86, pp. 257, 258).

ESTERASES AND LIPASES

An acidimetric micro method for esterase was described by Glick
(1934) ; this method was later applied to lipase by changing the sub-
strate (Glick and Biskind, 1935). The method is based on the con-

ESTERASES AND LIPASES 309

tinuous neutralization of the acid liberated by an alkaline buffer
medium. The loss in alkalinity of the buffer is then measured by
titration with acid to pH 6.5, the magnitude of the titration being
inversely proportional to the amount of substrate hydrolyzed. The
greatest deviation in the determination is equivalent to about 0.14
,al. 0.05 N acid.

An adaptation was also made for the determination of cholin-
esterase (Glick, 1938). The sensitivity of this method is equivalent
to the hydrolysis of 1 X 10"^ mole of ester. The titration is carried to
an end point of pH 6.2. Sawyer (1943) reported a sharper end point
and an increase in sensitivity when bromocresol purple is substi-
tuted for bromothymol blue and the end point is taken at pH 5.9.
For a gasometric method see page 393.

Glick Acidimetric Method for Esterase and Lipase

SPECIAL REAGENTS

Enzyme Extraction Medium. 30% glycerol.

Buffered Substrates. Esterase: 1% methyl butyrate in a buffer
soln. 0.1 A^ to sodium hydroxide and 0.4 A'' to glycine, pH 8.7 at
40°; prepare fresh before use. Lipase: Shake thoroughly a drop
of tributyrin in a few ml. of the buffer soln. above; let stand for
about 1 hr. in a refrigerator to allow the larger droplets to settle
and use the supernatant homogeneous emulsion, or filter through
paper and use the filtrate; prepare fresh before use.

Phenol-Indicator Solution. 10 ml. 2% phenol -[-1.5 ml. 0.04%
bromothymol blue.

0.05 N Hydrochloric Acid.

End Point Color Standard (pH 6.5). 3.1 ml. M/15 secondary
sodium phosphate -j- 6.9 ml. M/15 primary potassium phosphate
-}- 1 ml. 0.04% bromothymol blue.

PROCEDURE

1-4. These steps are carried out in the same fashion as the cor-
responding ones in the procedure for proteolytic enzymes (page 304).

5. Stop the reaction by adding 50 fA. phenol-indicator soln. with
a constriction pipette.

6. Titrate with 0.05 A'' hydrochloric acid, matching the greenish
color to that of the end point color standard. Take a uniform time

310 TITRIMETRIC METHODS

for each titration since a gradual bluing of the color occurs when the
soln. stands at a pH near neutrality.

Glick Acidinietric Method for Cholinesterase

SPECIAL REAGENTS

Enzyme Extraction Medium. 30% glycerol.

Buffered Substrate Solution. 0.4% acetylcholine chloride in veronal

buffer, pH 8.0 (7.15 ml. 0.1 M sodium diethylbarbiturate + 2.85

ml. 0.1 M hydrochloric acid.) Dissolve the substrate in the buffer

just before use.
Eserine-Indicator Solution. Add 10 ml. 0.1% eserine sulfate to 1.5

ml. 0.04% bromothymol blue, or, according to Sawyer (1943), to

1.5 ml. 0.04% bromocresol purple.
0.05 N Hydrochloric Acid.
End Point Color Standard (jdH 6.2 or 5.9). 10 ml. i¥/15 phosphate

buffer -\- 1 ml. 0.04% bromothymol blue or bromocresol purple.

PROCEDURE

Identical with that in the preceding method with the substitution
of the special reagents required for this enzyme.

CATALASE

Holter and Linderstr0m-Lang (1936) and Holter and Doyle
( 1938) described an adaptation to the micro level of the iodometric
catalase method of Stern (1932). The estimation of the decomposi-
tion of hydrogen peroxide by the enzyme is made by thiosulfate
titration of the iodine formed by oxidation of iodide placed in the re-
action mixture. The precision of the titration is 0.06 /a1. 0.02 N thio-
sulfate, which is equivalent to the decomposition of 0.02 /xg. hydrogen
peroxide.

Holter and Doyle Method for Catalase

SPECIAL APPARATUS

A 15 ix\. constriction pipette for the substrate is surrounded by a
water jacket, as shown in Figure 54 (page 173). Circulate ice water
through the jacket to keep the soln. cold during the pipetting.

CATALASE 311

SPECIAL REAGENTS

Substrate Solution. 0.01 M hydrogen peroxide in 0.03 A^ phosphate

buffer, pH 7.0. Prepare and keep at 0°.
Molybdic-Sulfuric Acid Solution. Make up 10 ml. saturated molyb-

dic acid to 100 ml. with 33% sulfuric acid.
2% Potassium Iodide.

0.2% Soluble Starch in 0.5% Potassium Iodide.
0.02 N Sodium Thiosulfate.

PROCEDURE

1. Pipette 5-7 fA. enzyme soln. into a reaction tube (Fig. 38, page
167) coated with ceresine (page 169). Add a mixing "flea," and cool
to 0°.

2. Add 15 fA. substrate soln. from the cooled pipette. Mix, and
keep the liquid at 0°.

3. After a 1-2 hr. reaction period at 0°, add 10 /xl. molybdic-
sulfuric acid soln. and 10 /xl. 2% potassium iodide.

4. Immediately after adding the iodide, place a seal of 20 /^l.
starch soln. across the lumen of the tube about 5 mm. above the sur-
face of the reaction mixture to prevent loss of iodine.

5. After 3 min., pass the tip of the microburette through the
starch film into the liquid below, and titrate with 0.02 N thiosulfate.
After most of the thiosulfate has been added, draw the starch film
down into the mixture and finish the titration.

IV. GASOMETRIC TECHNIQUES

As in the usual macro techniques, both volumetric and manometric
methods have been employed for studies on single cells and small
cellular aggregates of a well-defined nature. While some are simply
refinements of the more macro methods which have been modified
for the use of small quantities, other methods involve principles
hitherto not applied to gasometric measurements, e.g., Cartesian
diver manometry.

No mention will be made of the Warburg or Barcroft apparatus
not only because they are adapted to measurements of a relatively
macro order, but also because they have been thoroughly treated in
previously published works such as those of Dixon ( 1943) and Um-
breite^aL (1945).

A. VOLUMETRIC

The gasometric techniques of histo- and cytochemical interest
which have been based on volumetry have been developed chiefly for
respiration studies. In addition to these, gas analysis techniques
devised for use with very small volumes of blood will also be in-
cluded, since they may prove useful in some investigations.

•The advantage of volumetric over manometric apparatus for gaso-
metric measurements depends largely on the fact that the former
enables direct measurement of gas volume without the necessity of
determining the volume of the apparatus. Furthermore, volumetric
apparatus permits experimentation at constant pressure, which may
be advantageous when a large proportion of liquid is present. How-
ever, the determination of the volume of gasometric apparatus is
no great problem, and the choice of the technique need not be based
on this factor. The particular nature and magnitude of the gas
changes to be considered, combined with the availability of the ap-
paratus and the personal tastes of the experimenter, will be more
cogent factors in the selection of a technique.

313

314 GASOMETRIC-VOLUMETRIC METHODS

1. Capillary Respironietry

The technique of vokimetric capillary respirometry is based on the
measurement of changes in the gas volume in a capillary tube con-
nected to a chamber in which the respiring sample has been placed.
The position of an index droplet placed across the lumen of the
capillary serves as an indication of the volume at any given moment.

The history of the use and development of capillary respirometers
has been documented in a review by Tobias (1943) and need not
be presented here. In the following a description will be given only
of a few of the more modern designs adapted to measurements on the
histo- or cytochemical level.

The differential instruments consist essentially of two chambers
connected by a capillary tube containing an index droplet. One
chamber contains the biological material, medium, and reagents,
and the other only the medium and reagents. Changes in the amount
of gas which result from the biological action are followed by meas-
urements of the displacement of the index droplet. In the closed sys-
tem used, barometric fluctuations are without effect. When both
chambers, drilled in a block of heat-conducting material, are of
nearly the same size, temperature variations are of significance only
in so far as they influence the rate of the reactions measured, and
changes occasioned by the medium alone are cancelled. However,
in this case, the displacement of the index droplet is only about half
that obtained if the end of the capillary were open to the air. By
employing a compensating chamber which is very large with respect
to the reaction chamber, the movement of the index droplet in the
differential instrument can be made to approximate that in the open
type ; but more careful temperature control will be required, since a
given variation in temperature will take longer to exert its full effect
on the larger chamber.

(a) Cunningham-Bar th-Kirk Differential Respirometer

Cunningham and Kirk ( 1940) described a differential respirometer
with chambers of approximately equal size, which has the advantages
of enabling substitution of different capillary tubes, alteration of
chamber volume, filling of the chambers with various gas mixtures
as desired, and mixing of solutions during an experiment by means
of an electromagnetic "flea" (page 179). Since the symmetrical

CAPILLAKY RESPIROMETRY

315

chambers are bored in a solid brass block, uniform temperature dis-
tribution is insured, and thermostasis is required only to control the
rate of the reaction to be measured. The apparatus is adapted to

LJ

(i)

T\

Irj-

r

-IS

^1

1"

^

L__

1
1

^r

^^

Fig. 95. Construction diagram of differential microrespirometer.
Lengths are given in inches.
From Barth and Kirk (1942)

measuring gas changes of the order of 0.1-10 /xl. per hour with a read-
ing accuracy down to 0.001 /xL, depending on the capillary diameter.
In an experiment on the respiration of single pupae of Drosophila
nielanogaster the mean deviation from the mean of the six organisms
whose oxygen consumption was measured was 4 X 10"^ ju.1. per minute
or an average of 2.4%. Barth and Kirk (1942) subsequently simpli-
fied the construction of the apparatus and rendered it more adapt-
able to multiple determinations; it is this apparatus which will be
described (available from Microchemical Specialties Co.)

Apparatus. A diagram of the Barth-Kirk respirometer is given
in Figure 95: The main block {A) is made from a 3 in. length of
rectangular brass ( 1^/4 X % in. ). On one narrow face two chambers

316 GASOMETRIC-VOLUMETRIC METHODS

are drilled symmetrically P/ie in. apart and Vs in. diameter, and a
machined rim is made around each chamber. Threaded bolts are
placed at each end of the block as shown and a flat head plate (B)
of Lucite (Vs in. thick) is drilled to fit over the bolts. The head plate
has the same dimensions as the top of the block and has a 1 mm.
hole drilled over the center of each chamber. A glass capillary tube

(D) with a bore 0.1-1.0 mm. and a length of 1% in. is flattened on
one side by grinding to a width of Ys to Vie in. and the ground sur-
face is polished on a smooth stone to remove the frosting. Two holes
separated by l^^/ie in. are drilled through the flat side into the capil-
lary lumen ; a blunt 27 gauge hypodermic needle may be used to drill
the holes. Rubber gaskets (C), perforated by a short length of 27
gauge needle, are used to seal the capillary tube to the head plate.
The capillary is cleaned with chromic acid mixture and the ends are
plugged with paraffin and sealed by warming. A channel brass strip

(E) is drilled to fit over the bolts so that by means of thin knurled
nuts (F) it may be used to hold the capillary tube, head plate, and
chamber block tightly together. The bottom of the channel brass is
slotted over the entire length of the capillary to press only on the
sides of the latter. The screws have to be slowly and evenly tightened
to prevent the index droplet from being forced into one of the vertical
connecting holes. All the surfaces to be sealed are coated with stop-
cock grease.

A paper scale may be placed under the center part of the capillary
to enable reading of the displacement of the index droplet, or a low-
power microscope fitted with an ocular micrometer may be used. The
index droplet consists of kerosene which has been treated for several
days with concentrated sulfuric acid and then stored over pellets of
sodium hydroxide in a closed vessel in order to remove unsaturated
compounds which might lead to resin formation. [Tobias and
Gerard, 1941, claim that isodecane (2,7-dimethyloctane) is the
index fluid of choice for use in small capillaries.]

The size of the chambers may be varied by filling them with
paraffin to any chosen level. The surface of the paraffin is then
smoothed out with the flat end of a metal rod. Both chambers are ad-
justed to about the some volume. A disc of Lucite with one depres-
sion to hold the biological material and another to hold the droplet
of alkali used to absorb carbon dioxide is placed in each chamber.

Both chambers are charged in exactly the same manner with the

CAPILLARY RESPIROMETRY 317

exception that the biological material is placed only in one of them.
The index droplet is placed on the capillary by introducing a little
kerosene into one of the drilled holes, and removing the excess by
absorbing it into the end of a hardwood toothpick. Then the droplet
is forced to the proper position and the capillary is clamped in place.

The assembled apparatus is tested for leaks by warming one end
with the hand. When the hand is removed, the index droplet will re-
turn to its original position if no leaks are present.

Calibration. The volumes of the chambers are determined, after
greasing the metal to prevent amalgamation, by filling with mercury,
squeezing out the excess by pressing a glass plate over the chamber,
and weighing the mercury left. The volumes of the Lucite discs are
also determined by weighing them and dividing by the specific grav-
ity of the Lucite. The volumes of the capillary posts and the capillary
tubes are calculated from measurements of the dimensions of the
holes.

Corrections for the volume of kerosene adhering to the capillary
walls were found to be 0.45 ± 0.10% for a 0.57 mm. diameter capil-
lary, and 0.42 ± 0.07 for a 0.22 mm. capillary, in experiments by
Cunningham and Kirk (1940). The corrections were determined by
measuring the length of a kerosene droplet before and after it was
made to move along the capillary for a known distance. The volume
of liquid on the walls, calculated from this measurement, divided by
the total volume of the capillary traversed by the droplet times 100
gives the percentage error.

Calculations. Cunningham and Kirk (1940) gave the deriva-
tions of the formulae which may be used to calculate changes in the
amount of gas in the apparatus. The change in the gas volume
(AFp„) at the initial pressure (Po), which equals the barometric
pressure at the time the apparatus is sealed, is given by the following
expression for the case of a reaction liberating carbon dioxide, as in
the interaction of acid and bicarbonate in the reaction chamber:

/ Vr. \

AVp, = nAd

-Yc - Ad/

where n is the ratio of the total volume of the two chambers to the
volume of the compensation chamber, A the effective cross-sectional
area of the capillary (actual area minus 0.4% to correct for the film
of kerosene which coats the wall), d the displacement of the index

318 GASOMETRIC-VOLUMETRIC METHODS

droplet and Vc the volume of gas in the compensation chamber and
capillary up to the index droplet. When Ad is negligibly small with
respect to Vc, the expression may be reduced to:

AVp„ = nAd

Conversion of (ATpJ to standard conditions is carried out in the
usual manner.

The preceding equations do not take into account the volume of
gas which may be dissolved in the liquid. For carbon dioxide the
following relation obtains at temperature t:

CO2 (dissolved) = (,, ^^\)j PraC02,-Vln

Wr + Ad/

where Vr is the volume of gas in the reaction chamber plus the
volume in the capillary up to the index droplet. P/ is the total gas

pressure in the reaction chamber, a C02< is the solubility coefficient
of carbon dioxide for the liquid at temperature t and one atmosphere
pressure of carbon dioxide (vol. gas at S.T.P./vol. liquid), and VIr
is the volume of liquid in the reaction chamber.

If the precision of the measurements merits corrections for dis-
solved oxygen and nitrogen in the liquid, additional formulae may
be applied. Thus the volume of oxygen or nitrogen forced into solu-
tion by compression of gas in the compensation chamber during the
experiment equals:

AO2 (dissolved) = a O^^-Vk-Po, (

P_f
Po

1 (a)

AN2 (dissolved) = a N2 -R-Pn. (^^ - 1 ) (&)

t

vPo

where a O21 and a N2t are the solubility coefficients of the respective
gases in the liquid at temperature t, Vic is the volume of liquid in

the compensation chamber, P02 and P^j are the initial partial pres-
sures of the respective gases (in calculating these values one should
not forget to consider the aqueous tension), and P/ is the final
pressure of the system. These equations are based on the fact that
the partial pressures of the gases in the compensation chamber have
increased to (P//Po) times their original values. Since the partial
pressures of the gases in the reaction chamber have decreased to

CAPILLARY RESPIROMETRY 319

{Vr — Ad)/VB times their original values, the volume of oxygen or
nitrogen released from solution in the reaction chamber during the
experiment equals:

AO2 (dissolved) = aOo^-F/«.Po, y^yj^^"^ - 1) (c)

AN2 (dissolved) = aN2,-y/«-PN, y^^f^ - 1) (^)

These last quantities (c and d) are negative, but they tend to give
rise to error in the same direction as the preceding ones (a and b) .
All four A values are added, and the sum is subtracted from AFp„.
Correction for dissolved carbon dioxide is made by adding the
volume that has dissolved to the value for AVp^.

In the case of respiration experiments the total pressure of the
system decreases. Hence:

AFpo = TiAd

V.

c

.V, + Ad^

Since only oxygen is removed during the respiration, the partial
pressures of oxygen and nitrogen do not remain equal in the two
chambers. In the compensation chamber, the partial pressures of
both gases are reduced to ( TV [T^c + -A-dl ) times their original value.
On the other hand, the partial pressure of nitrogen in the reaction
chamber has been increased to {Vr/IVr — Ad"]) times its initial
value, while the partial pressure of oxygen has decreased to:

( yn \

\Vr - Ad + nAdJ °'

Substitution of these factors for the partial pressure changes in the
preceding formulae for the volumes of oxygen and nitrogen dissolved
will permit the corrections to be applied.

(6) Cunningham-Kirk Open-Tube Respirometer

The capillary tube respirometer of Kalmus (1928) consists of an
enlargement at one end of a capillary tube to serve as a reaction
vessel while the other end of the capillary is open to the air. The
position of the meniscus of liquid in the capillary is used to indicate

320

GASOMETRIC-VOLUMETRIC METHODS

gas volume changes. The original technique of Kalmus has been the
object of considerable criticism, but Cunningham and Kirk (1942)

Type A Type B

Fig. 96. Left : two capillary respirometers, shown with
protective jackets of glass tubing. Right: respirometer
assembly with microscope for observing shift of meniscus.
From Cunningham and Kirk (1942)

have instituted improvements and they have chosen this open-tube
instrument in preference to differential types for studies requiring a
very high degree of sensitivity, e.g., measurements of the respira-
tion of a single Paramecium. The sensitivity obtained by Cunning-
ham and Kirk was 5 X 10"^ /^l- While the absolute accuracy is
doubtful, the accuracy for relative values was given as at least
dzl5% for the instruments used. It should be pointed out that the
technical difficulties involved in the use of this type of respirometer
are very great, and these will, no doubt, seriously limit its applica-
tion.

Diagrams of the instruments are given in Figure 96; their dimen-
sions follow:

Dimension

Type A

Type B

Volume of liquid phase

Volume of gas phase

Diameter of respiration chamber.

Diameter of manometer

Length of manometer

3.0 (A.

13.0 (i\.

0.5 lA.

0.5 ill.

0.56 mm.

0.56 mm

0.08 mm.

0.08 mm

200 mm.

200 mm

CAPILLARY RESPIROMETRV 321

Double Dewar flasks are used to maintain constant temperature
for the period of measurement (1-2 hr.l. The outer flask is sur-
rounded by a 1 in. layer of cotton packing; it has all. capacity
and is used three fourths filled with water. The inner flask has a 4
oz. capacity and is two thirds filled with water.

The respirometer tube is cleaned by filling it with cleaning solu-
tion and after 30 min. it is rinsed with distilled water, followed by
0.01 M sodium bicarbonate, and finally distilled water again. It is
then dried by drawing filtered air through it, and is allowed to stand
overnight to eliminate any temperature differences within the glass.

A respiration experiment on a Paramecium is carried out as fol-
lows:

1. Fill the tube to within 4 mm. of the chamber end with 0.5 N
sodium hydroxide by filling the cup at the top of the tube with al-
kali and allowing the tube to fill by gravity. When the solution
reaches the proper position pour out that remaining in the cup.

2. Place a droplet of tap water containing the organism over
the end of the respirometer chamber and force about 2 mm. of this
tap water into the 0.56 mm. capillary by inserting a needle through
the drop into the capillary and then withdrawing it,

3. Observe the end of the tube until the Paramecium enters it
and then quickly wipe off the surface tap water and seal the end of
the capillary with a piece of vaselined cover slip.

4. Fill the cup around the upper part of the tube with the sodium
hydroxide soln. and place the respirometer in the Dewar flasks.

5. After 1 hr. remove the alkali from the cup with a fine pipette
and plug the top of the cup with cotton.

6. After 1 hr. more focus the microscope (magnification lOOX)
on the meniscus of the liquid in the capillary and record the rate of
fall of the meniscus. During these measurements, note the temper-
ature and barometric pressure. If these vary sufficiently to result in
significant errors, discard the experiment.

The errors introduced by changes in (1) temperature, {£} baro-
metric pressure, (3) height of the liquid in the tube, (4) volume on
dilution of the alkali soln., (5) volume of the dissolved gas phase with
change in partial pressure, (6) bore of the capillary, and (7) surface
tension of the liquid in the capillary, have been discussed by Cun-
ningham and Kirk (1942) . The original paper of these authors should

322

GASOMETRIC-VOLUMETRIC METHODS

be consulted for the determination of each of these errors. For the
two instruments described by them, the algebraic sum of those errors
which are constant and independent of the presence of the organism
in the respirometer equalled —1.6 X 10-* fA./hr. for the type A, and

Fig. 97. Glass stopper with

respirometer tubes attached.

Fnrm Tobias (1943)

Fig. 98. Complete Tobias-Gerard

respirometer assembly.

From Tobias (191,3)

—4.2 X lO"'* ij\./\\v. for the type B instrument. The summation of
the errors which are dependent on the presence of the organism
amounted to 0.147 v for the type A, and 0.407 v for the type B in-
strument, where v is the observed value for the volume of oxygen
measured.

(c) Tobias-Gerard Respirometer

Tobias and Gerard (1941) employed the principle, used earlier
by Gerard and Hartline (1934), of placing a small capillary reac-

CAPILLARY RESPIROMETRY 323

tion chamber in a large compensation vessel which is sealed off to
make the system independent of barometric changes. Volume
changes within the capillary chamber are so small in comparison to
the volmiie of the compensation vessel that these changes can be
considered completely undamped, and the arrangement makes for
great stability in measurement. An improvement initiated by Tobias
and Gerard in this type of respirometer is the use of ten or more of
the capillaries at the same time in a single compensation vessel.
Gas volume changes of the order of 0.0005 to 0.001 microliter per
minute can be followed at minute intervals with individual readings
varying from the mean by about 8%. For five to ten minute inter-
vals the accuracy is about 2%.

Apparatus. The respirometer is made by sealing a 90 mm.
length of 0.2 mm. glass capillary tubing into a 30-40 mm. length of
1.2-1.5 mm. thin-walled glass tubing with DeKhotinsky cement. The
wider tube serves as the reaction chamber while the capillary con-
tains the index droplet by means of which gas vol. changes are
measured. It has been found that isodecane (2,7-dimethyloctane)
is superior to other index fluids for small capillaries. Bits of filter
paper soaked with acid or alkali, and separated by dry paper guards,
may be used in the reaction chamber to absorb certain gases during
the reaction. The absorbing materials and the tissue are placed in
the reaction tube. The open end of the latter is then sealed with
plasticine. The isodecane droplet is placed in the capillary and the
respirometer is mounted on a central thick-walled capillary tube
connected to the glass stopper of the compensation vessel (Figs.
97, 98) . A number of the respirometers may be mounted around the
central tube as shown. The glass stopper bearing the respirometers
is then fitted into the compensation chamber, which is supported
horizontally. The entire glass unit is submerged in a thermostat
bath. By rotating the stopper, individual capillaries can be brought
into the field of the horizontal micrometer microscope employed to
determine the positions of the index droplets. At the end of an ex-
periment the small capillary is broken off and the diameter of the
lumen at the end is measured with an ocular micrometer.

Certain mechanical improvements have been introduced (Tobias,
1943) : "Permanent, heavy-walled capillary units, expanded at one
end to accommodate tissue and absorbing reagents, replace the fine
capillaries. A small ground-glass cap instead of plasticine closes the

324 GASOMETRIC-VOLUMETRIC METHODS

chamber. The units are mounted in a vertical bank inside a brass-
glass box. Brass clips into which the respirometers fit are rotatable
from the outside of the box by means of a metal arm connected
thereto by a packed joint. Since the tissue being studied clings to the
roof of the expanded end of the unit, a droplet of substrate to be
added may be placed on the floor along with a small lead shot coated
with celloidin and paraffin. Rotation of a single capillary at any
time then causes the mixing ball to drag the reagent around to the
tissue. Greasing of the cap-respirometer joint makes it transparent
and thus allows visual observation of the tissue being studied."

(d) Scholander Micrometer Burette Differential Respirometers

Scholander ( 1942a j adapted his micrometer burette (page 262)
to the measurement of small gas changes in volumetric respirometers
of his design. One of the respirometers was sensitive to about 0.3 fA.
per hour and another to about 0.01 /xl. per hour. Greater refinement
would be achieved by reducing the dimensions of the micrometer
spindle and the glass parts. In a subsequent publication Scholander
and Edwards (1942) described a larger respiration chamber for an
apparatus designed for measurements on aquatic organisms such as
sand crabs, dragonfly larvae, or water plants. This instrument was
sensitive to within 1 ixl. per hour.

The principle of the apparatus lies in the maintenance of con-
stant gas volume in a reaction vessel by the addition of oxygen
from a micrometer burette to replace that used up by the respira-
tion. The carbon dioxide evolved is absorbed by alkali. The pressure
in the reaction vessel is balanced against that in a compensation
vessel.

Respirometer Sensitive to 0.3 Microliter per Hour. This
apparatus (Fig. 99) consists of a micrometer burette (1), storage
bulb for oxygen (2), respiratory chamber (3), manometer (4), and
a compensation vessel (5). The capillary bore is 1 mm. The frame
holding the apparatus is held by a rod which passes through a hole
(7), and it rests on the water bath at 6. The biological material
in 3 is separated from the carbon dioxide absorbent, which is in the
bottom of the same vessel. A ground joint, or a syringe needle
pushed through a rubber stopper as shown in 3, may be used to
connect the respiratory chamber with the rest of the apparatus.

CAPILLARY RESPIROMETRY

325

Brodie fluid is placed in the manometer and water is used in the
compensation vessel. A layer of water is also used to cover the
mercury in 2.

The chamber {3) is attached, while both stopcocks are open to the
air. At the start of an experiment the stopcocks are set as shown in

Fig. 99. Volumetric respirometer,

sensitive to about 0.3 yX. per hour.

From Scholander (1942a)

Fig. 100. Volumetric respirometer,

sensitive to about 0.01 fi\. per hour.

From Scholander (1942a)

the figure ; during the experiment the micrometer is used to keep the
menisci level in the manometer tube by replacing the oxygen as it
is consumed. In order to prevent contamination of the oxygen in 2,
the stopcock attached to the oxygen storage vessel is kept closed
except when the micrometer adjustment is made.

Respirometer Sensitive to 0.01 Microliter per Hour. This
apparatus (Fig. 100) consists of a micrometer whose spindle has
been replaced by a Vie in. diameter drill rod which passes through
a fiber washer with an air-tight fit into the glass tube {2) . The glass
part is held in the micrometer frame. Oxygen is stored at 3\ the
respiratory chamber fits on the side arm at 4; the indicator drop
is at 5, and the compensating chamber at 6. The capillary bore is
about 0.25 mm. The micrometer is screwed to a heavy brass foot as
illustrated.

The apparatus is first filled with mercury and carefully freed from
air bubbles. The reservoir [3) is filled with oxygen after a little
water is drawn in to cover the mercury. The respiration chamber is
then connected; it contains a loop of platinum wire to hold the
sample in a drop of medium while the carbon dioxide absorbent is

326 GASOMETRIC-VOLUMETRIC METHODS

placed on the bottom of the vessel, or the absorbent may be placed
in the loop and the sample on the bottom.

An index drop of 2% Turgitol, or other wetting agent, is intro-
duced through the compensation chamber with a hypodermic needle.
Then a drop of water is placed in the compensation chamber, and
finally the opening is greased and sealed with a fiat piece of glass.
The instrument is operated in a manner similar to that employed
with the larger apparatus.

2. Gas Analysis

(a) Scholander Micrometer Burette Gas Analyzer

Scholander (1942b) employed his micrometer burette to develop
an apparatus by means of which 10 /xl. of a gas mixture may be
analyzed for its various components with an accuracy of about
0.1% of the total sample. Specific absorbents for each component
are used, and the volume change resulting from the absorption of
each gas is measured.

Apparatus. The apparatus is shown in Figure 101 (available
from 0. Hebel, Edward Martin Biological Laboratory, Swarthmore
College) . The micrometer spindle is replaced by a ^/le in. drill rod
(2) , which displaces about 50 /xl. by its full traverse. Each microm-
eter scale division corresponds to approximately 0.02 /xl. and esti-
mates are safely made to 0.005 /xl. The capillary bore is around 0.25
mm. and the bulb (3) has a capacity of about 50 /xl. The absorption
chamber (1) has a bore no greater than 2.5 mm. The fine line (4) is
used for reading. The drill rod must fit perfectly tight through a
fiber washer (5) and the mercury vessel must be completely freed
from air. The waterjacket surrounding the burette is clamped so
that it can be tilted by the handle (6) .

Scholander uses separate 2 ml. syringes to hold the respective
liquids required, i.e., mercury, a manometer liquid of 2% Turgitol
(wetting agent), 0.5 N sulfuric acid, and gas absorbents such as
0.25 A'' potassium hydroxide which has been shaken well with air
(for carbon dioxide) and hydrosulfite (for oxygen). (The latter is
prepared by adding a mixture of 10 parts hydrosulfite and 1 part
sodium anthraquinone-/?-sulfonate to a small test tube filled with
0.25 N potassium hydroxide, containing a drop of mercury for mix-

GAS ANALYSIS

327

ing, until a dark red solution is obtained. The full tube is closed,
taking care to exclude air bubbles, shaken, and finally stored in a
syringe with a bubble of nitrogen.) Each syringe is connected by
stiff rubber tubing to a drawn-out glass tube. The orifice of the glass

wmm

\

i

Fig. 102. Steps in the
introduction of the sample
into the 10 fil. gas analyzer.
From Scholander (1942h)

Fig. 101. Analyzer for 10 lA. of gas mixture.
From Scholander (1942h)

tip should not be fine enough to cause pressure to develop in the
syringe" during delivery, since this would force air or nitrogen into
solution and result in low absorption values. A glass tip connected
to suction is also required. Syringe needles, if not finer than 20
gage, may be substituted for the glass tips.

The transfer pipette (Fig. 102) is convenient for handhng the gas
sample {&). It consists of 2-2.5 mm. bore glass tube drawn out at
one end and connected to a piece of rubber tubing at the other. One
end of the rubber tube is plugged with a piece of glass rod. Mercury
is used to confine the sample as in ^. A transparent cap with a cen-
tral hole fits over the absorption chamber to guide the tip of the
pipette.

Manipulation. As an example of the procedure to be followed,
consider the analysis of a mixture of carbon dioxide, oxygen, and
nitrogen :

1. Rinse the absorption chamber and bulb with 0.5 N sulfuric
acid and move the mercury up to the absorption chamber.

328 GASOMETRIC-VOLUiMETKIC METHODS

2. Introduce the gas sample (about 10 jul.) into the absorption
chamber following the steps shown in Figure 102. Guide the tip of
the pipette so that it does not touch the wall of the chamber during
the transfer.

3. Draw the mercury back into the capillary until the meniscus
is at the mark, and then take the first micrometer reading.

4. Draw the gas into the capillary, and after it a little mercury.

5. Suck away the mercury left in the absorption chamber and
replace it with manometer fluid.

6. Expel the residual mercury in the capillary into the manom-
eter fluid contained in the absorption chamber. Draw in a little of
the fluid, and suck away that remaining in the chamber. Leave only
2 mm. of the fluid in the capillary.

7. Bring the gas-fluid meniscus to the mark and take a second
reading. The volume of the sample is obtained from the difference
between the two readings.

8. Move the manometer fluid out into the absorption chamber
again and fill the chamber with the potassium hydroxide solution td
join the fluid without enclosing any air bubbles.

9. Move the gas sample out into the alkali in the chamber until
the gas bubble breaks loose. Then move the bubble back and forth
by tipping the instrument.

10. Draw the bubble back into the capillary and take a third
reading when the gas-mercury meniscus reaches the mark.

11. When the last of the gas has been drawn into the entrance
of the capillary, replace the alkali by manometer fluid and then
suck all of the fluid away except a short drop in the capillary.

12. Move the fluid-gas meniscus to the mark and take a fourth
reading. The volume of the sample minus the carbon dioxide is
obtained from the difference between the third and fourth readings.

13. Repeat steps 8-12 using the hydrosulflte solution to absorb
the oxygen. Allow 2 min. for the oxygen to be absorbed. The gas
remaining after this absorption is entirely nitrogen.

(6) Berg Simplified Gas Analyzer

Using the principle of the Scholander micrometer-burette gas
analyzer. Berg (1946) employed the simplified apparatus illustrated
in Figure 103. The thermometer tubing is 6 in. long and has a bore
of 0.11 mm. The water jacket surrounding it affords protection

GAS ANALYSIS . 329

against temperature change during the analysis. A saturated solution
of lithium chloride, preferably equilibrated with gas of approxi-
mately the same composition as the sample to be analyzed, is used
to fill the apparatus. The gas sample is introduced into the absorp-
tion chamber while the instrument is held vertically with the cham-
ber end down. By opening the screw clamp on the rubber tubing
attached to the capillary, the gas is drawn into the latter and the
volume is determined by noting the length of the gas column and
multiplying it by the cross-sectional area (0.0095 mm.-). The liquid
in the absorption chamber is then replaced by the gas-absorbing
solution (lithium chloride containing potassium hydroxide for car-
bon dioxide, and alkaline pyrogallol for oxygen) and the gas is
forced out into the absorption chamber. The unabsorbed portion of
the gas is then drawn back into the capillary and its volume is

Absorption chamber Clamp Glass rod

/^_^^. , ...,, ,, ■"''■'■'"'"'^'1^^ I Fig. 103. Simplified

f^ ' . J.; j'n ^ ^^^ analyzer.

^ ^ .... .-..^Z y ij' /"' ' From Berg (1946)

Thermometer tubing ^ Water lacket ^ Pressure tubing

measured. To be sure of complete absorption the gas is re-expelled
into the absorption chamber and after a number of seconds redrawn
into the capillary. This process is repeated until no change in the
volume of the residual gas is observed.

The small bore of the capillary enables the analysis of samples in
the range 0.4-1 fA. Under the most favorable conditions the experi-
mental error can be reduced to <0.3%. The chief source of error is
the diffusion of the gas into the lithium chloride solution and hence
the need for saturating this solution with gas of approximately the
same composition as that of the sample to be analyzed. There might
be an advantage in using mercury as the analyzer fluid in a manner
similar to that employed by Scholander with the micrometer-burette
apparatus.

(c) Scholander-Ronghton Syringe Gas Analyzer

The syringe gas analyzer, first described by Scholander and
Roughton (1942), has been used primarily for the estimation of
gases in blood samples of the order of one drop. Analysis can also
be performed on other fluids, and, in general, the apparatus may be
used for the analysis of components in gas mixtures. As applied to

330

GASOMETRIC-VOLUMETRIC METHODS

blood gases, oxygen and carbon monoxide can be determined in 40
fx\. blood samples with an accuracy of 0.15 to 0.20 volume per cent,
nitrogen in 120 fA. samples to 0.05 volume per cent, and carbon
dioxide in 13 [A. samples to about 1 volume per cent. The time
required for a determination is 6-10 min. In these methods the blood
is drawn into a capillary attached to a 1 ml. tuberculin syringe, the
gas is liberated by a suitable reagent, the gas volume is noted,
specific absorbents are used to remove the gases separately, and the
volume change produced by the removal of each component is
recorded.

20

30

Fig. 104. Shaking of syringe

and extraction of gas.

From Roughton and Scholander (1943)

Fig. 105. A and B, syringe showing the tech-
nique for absorption of the carbon dioxide used
for extraction. C, temperature equihbration of
gas bubble in capillary before the reading is
made. From Roughton and Scholander (1943)

The apparatus, which was slightly modified by Roughton and
Scholander (1943), consists of a 1 ml. Pyrex tuberculin syringe with
an arresting clip on the plunger to prevent its slipping, and with a
standard bore precision 0.5 mm. Pyrex capillary fused to its nozzle
(Fig. 105). The top of the capillary is expanded to a cylindrical cup
of about 1.5 cm. length and 2.5 mm. bore. The capillary, which is

GAS ANALYSIS AND OXYGEN 331

7-8 cm. long, is graduated into thirty divisions, each 2 mm. long.
A detachable rubber cup of about 1 ml. capacity may be fitted over
the cylindrical glass cup.

The sampling pipette consists of a piece of thin-walled glass tub-
ing having a 1-1.5 mm. bore, and it is drawn out to a tip which is
ground smooth in order to permit a snug fit into the bottom of the
glass cup. The mark on the pipette is placed so that the volume from
the mark to the tip is equal to 100 divisions on the capillary, i.e.,
39.3 fA. The capillary and the pipette can be calibrated with a
microburette or by mercury weighing. (Interchangeable pipettes and
syringes are supplied by Mr. J. D. Graham, Department of Physi-
ology, University of Pennsylvania Medical School.)

Collecting Blood Samples. Citrated blood samples may be
collected directly in the blood pipettes from finger pricks as de-
scribed by Scholander (1942c). A short tube, 15 mm. long, flanged
out at one end is held on the finger tip by a rubber band which
presses the flange against the skin. A few crystals of citrate are
placed in the tube; a tourniquet is wound around the finger toward
the tip. The finger is pricked through the tube using a spring blood
lance, and blood is quickly drawn into the pipette from the bottom
of the tube.

The blood may be collected anaerobically in syringes and trans-
ferred to the pipettes. The transfer is made by setting the pipette on
a table with the tip protruding about 1 in. over the edge, pressing
the tip into the opening of the syringe nozzle while the syringe is
held horizontally, and then slowly filling the pipette by screwing
in the plunger (Roughton and Scholander, 1943).

OXYGEN

Roughton and Scholander (1943) described the oxygen method in
which an excess of carbon dioxide is used to extract the oxygen, the
carbon dioxide is then absorbed in alkali, and the volume of the
residual gas is measured in the capillary before and after the oxygen
is absorbed by alkaline pyrogallol.

Roughton and Scholander Method for Oxygen
SPECIAL REAGENTS

Caprylic Alcohol.

Ferricyanide Solution. Dissolve 12.5 g. potassium ferricyanide, 3 g.

332

GASOMETRIC-VOLUMETRIC METHODS

potassium bicarbonate, and 0.5 g. saponin in distilled water and
make up to 50 ml. Do not use the soln. after 3 days.

Acetate Buffer. Dissolve 70 g. sodium acetate (NaC2H302.3H20)
in 100 g. distilled water and add 15 ml. glacial acetic acid.

i5% Urea (used as cleaning solution).

10% Sodium Hydroxide.

Pyrogallol Solution. Add 15 g. powdered pyrogallol to 100 ml. 20%
sodium hydroxide under a 2 cm. layer of oil. Dissolve the pyro-
gallol under the oil by stirring with a glass rod.

PROCEDURE

1. With the syringe analyzer in a vertical position and the
plunger pushed up, fill the cup with ferricyanide soln. Draw the soln.
down to the bottom of the syringe, push it back up into the cup, and
suck it out. Repeat twice with fresh lots of ferricyanide, taking care
to avoid trapping air bubbles. Leave the dead space in the syringe
full of ferricyanide. Use no grease or oil in the syringe.

Fig. 106. Transfer of blood

from pipette directly to capillary.

From Roughton and Scholander (1943)

Fig. 107. Further details
of the transfer of the blood
from the pipette to the
capillary. From Roughtoii
and Scholander (1943)

2. Fill the glass cup to the mark with ferricyanide soln. and
draw it down to the bottom of the cup.

3. Place a drop of caprylic alcohol on the bottom of the cup.

OXYGEN 333

4. Draw the sample from the pipette, which has been filled to
the mark (about 40 fA.) into the capillary as shown in Fig. 106.
Pull out the plunger gradually so that the sample is slowly and
evenly drawn in followed by an air bubble of about 1 mm. length
(A and B, Fig. 107) . The air bubble prevents the blood from getting
back up into the pipette. If the pipette tip is properly ground and
the correct pressure applied during the transfer, no appreciable
quantity of caprylic alcohol will be drawn into the capillary.

5. Remove the pipette quickly, and expel the air bubble {C, Fig.
107) through the caprylic alcohol, using a fine wire if necessary or
tapping the capillary.

6. Draw in a column of the caprylic alcohol two divisions in
length onto the top of the blood and suck out the rest of the
caprylic alcohol from the cup.

7. Fill the cup to the mark with acetate buffer and draw it down
to the bottom of the cup.

8. Immediately fill the cup to the top with 45% urea and close
firmly with the finger.

9. Shake the closed apparatus vigorously for 2 min. in a hori-
zontal position, gradually pulling out the plunger as the gases are
evolved to keep the gas pressure in the syringe about 1 atmosphere.
About 0.75 ml. is usually evolved (Fig. 104) . The amount of carbon
dioxide evolved may be varied by changing the strength of the
bicarbonate in the ferricyanide soln.

10. Carefully release the finger while adjusting the plunger to
keep the gas meniscus in the capillary. Let a little urea soln. run
down into the capillary, and allow it to remain there until the walls
are perfectly clean.

11. Remove three fourths of the urea soln. from the cup. Fit the
rubber cap over the glass cup, and fill it with 10% sodium hydroxide
without trapping air bubbles {A, Fig. 105).

12. Draw a little of the alkali into the syringe. As the carbon
dioxide is absorbed more alkali will be sucked in until only a small
bubble of the other gases will remain {B, Fig. 105). The absorption
requires a few sec, and, before it is complete, carefully move the
residual bubble into the capillary to prevent reabsorption of oxygen.

13. Remove the rubber cup. Empty the glass cup, and set the
capillary in a beaker of water at room temperature for 30 sec. (C,
Fig. 105).

334 GASOMETRIC-VOLUMETRIC METHODS

14. Remove from the water, dry by lightly wiping, taking care
not to handle the capillary, and read the gas vol. (V'l) in divisions.

15. Fill the glass cup with pyrogallol soln. and absorb the oxygen
by pulling the gas down to the bottom of the capillary and back
again a few times. Finally move the gas bubble veiy slowly up to
the top part of the capillary. Equilibrate the temperature, and again
read the vol. (72) in divisions. When V2 is only a few divisions,
the second temperature equilibration may be omitted.

16. Run a blank on the reagents by omitting the sample.

17. Calculate the oxygen content from the formula:

Oxygen = {\\ - V2 - C)f

where C is the blank correction for oxygen in the reagents, and / is
the correction factor for temperature, aqueous vapor tension, and
barometric pressure. C amounts to 1.0 to 1.1 volume per cent at
room temperature as a rule ; / may be obtained from the usual tables
such as that given by Peters and Van Slyke (1932, page 129, Table
15).

CARBON MONOXIDE

Scholander and Roughton (1943a) described three applications
of the syringe analyzer to the measurement of carbon monoxide in
blood: a general method for saturation ranging from 0-100% car-
bon monoxide hemoglobin, a method for the measurement of both
oxygen and carbon monoxide on the same sample of blood, and a
method sufficiently precise for the determination of blood volume in
which the carbon monoxide level is held below 2 volume per cent.
Only the first of these three will be described.

Scholander and Roughton Method for Carbon Monoxide

SPECIAL REAGENTS

Winkler Solution. Place 20 g. cuprous chloride, 25 g. ammonium
chloride, and 75 g. water in a bottle just large enough to contain
them. Stopper the bottle, shake with as little air as possible, and
allow the precipitate to settle. Put a coil of copper wire in the
soln. and cover the liquid with a layer of paraffin oil. The reagent
becomes almost colorless after a while.

CAEBON MONOXIDE AND NITROGEN 335

Other Reagents. The other reagents required are those listed for
the oxygen method (page 331) with the exception of the 10%
sodium hydroxide. The distilled water is aerated.

PROCEDURE

1-10. Same as for oxygen (pages 332-333).

11. Substitute pyrogallol soln. for the 10% sodium hydroxide,
but otherwise same as for oxygen (page 333).

12-14. Same as for oxygen (pp. 333-4). Read vol. of bubble (Fi).

15. Flush the glass cup clean with water and leave filled. Quickly
pull about three fourths of the water in the cup down into the
syringe to form a layer over the blood mixture. Then immediately
run the bubble with the clean water below it up into the top of the
capillary.

16. Empty the water from the glass cup and fill it with Winkler
soln.

17. Incline the syringe so that the cup points downward at a
slight angle from the horizontal. Cautiously screw in the plunger to
drive the gas bubble out into the glass cup where it can rest near
the junction of the capillary and the cup. Suck in the Winkler soln.
behind the bubble to half fill the capillary, and rotate the instru-
ment gently for a few sec. to complete the absorption of the carbon
monoxide. Then turn the syringe to the vertical position with the cup
down and suck the gas bubble back into the capillary. Measure the
gas vol. (72) in the usual fashion, and calculate the carbon mon-
oxide content from the formula:

Carbon monoxide = (Fj — Fg)/

where / is the correction factor (page 334),

NITROGEN

The adaptation of the syringe analyzer to the measurement of
nitrogen in fluids was made by Edwards, Scholander, and Roughton
(1943). Their method calls for a pipette with two marks, calibrated
to deliver not only the usual volume (page 331) but also three
times this. The principle of the method depends on extraction of
all the nitrogen and part of the oxygen from the sample with carbon
dioxide, absorption of the oxygen and carbon dioxide with alkaline
hydrosulfite, and measurement of the nitrogen which remains.

336 GASOMETRIC-VOLUMETRIC METHODS

Edwards. Scholander, and Roughton Method for Nitrogen

SPECIAL REAGENTS

Bicarbonate Solution. Dissolve 11 g. potassium bicarbonate in 100
g. water.

Acid Phosphate Biiffer (about 5 M). Dissolve 95 g. sodium dihy-
drogen phosphate (NaH2P04.H20) in 100 g. warm water.

Hydrosulfite Solution. To 50 ml. of 20% potassium hydroxide add
15 g. of a mixture of 10 parts sodium hydrosulfite and 1 part
sodium anthraquinone-^-sulfonate. Store well stoppered in con-
tact with as little air as possible.

45% Urea.

Caprylic Alcohol.

Aerated Distilled Water.

PROCEDURE

1. Same as for oxygen (page 332) substituting the bicarbonate
soln. for the ferricyanide.

2. Dry the glass cups with cotton or filter paper and place a drop
of caprylic alcohol in the bottom of the cup without trapping air
bubbles.

3-5. Same as steps 4-6 in oxygen method (page 333). Take care
to prevent caprylic alcohol from being drawn down into the capil-
lary with the sample. A 120 ;ul. sample is used.

6-9. Same as steps 7-10 in oxygen method (see page 333)
substituting the acid phosphate bufi'er for the acetate. Draw the
urea soln. down to the bottom of the capillary but do not let it enter
the syringe barrel.

10. Holding the syringe vertically, attach the rubber cap, and
add about 1 ml. hydrosulfite soln. without trapping air bubbles in
the glass cup.

11. Draw a little hydrosulfite into the syringe; the vacvmra
created by the gas absorption will draw in the rest of the soln.
required for the complete absorption of the carbon dioxide and
oxygen.

12. Push the bubble up into the lower part of the capillary;
suck out the hydrosulfite ,from the rubber cap and detach the latter.
Fill the glass cup with water, and draw three fourths of it down over
the bubble.

NITROGEN AND CARBON DIOXIDE 337

13. Push the bubble up into the clean capillary very gently.
Equilibrate the temperature and read the volume (Fi) in divisions
(steps 13, 14, pages 333 and 334).

14. Run a blank on the reagents by substituting aerated distilled
water for the blood; however, use one third the vol. of water, i.e.,
to the first mark on the pipette (about 40 i-d.) . The blank then
equals V — (a//) where V is the uncorrected nitrogen reading, a is
the solubility of atmospheric nitrogen in water at room temperature
in volume per cent (at 22°, a = 1.2), and / is the correction factor
(page 334). The authors of the method found the blai\k to be con-
stant and of the order of 1.3 to 1.5 units on the capillary, depending
on the instrument used.

15. Calculate the nitrogen content of the blood from the formula:

Nitrogen = (T\ - C) (//3)
where C is the blank correction for the nitrogen in the reagents.

CARBON DIOXIDE*

Certain changes in the syringe analyzer technique were intro-
duced by Scholander and Roughton (1943b) to enable the estimation
of carbon dioxide. The gases are vacuum extracted from the sample
mixed with acid buffer, and the gas volume is measured before and
after absorption with alkali.

This determination requires a rubber-tipped wooden plug, made
by dipping the end of a round toothpick in rubber latex, leaving
a drop on the tip, and drying it, tip downward, at a moderate
temperature in an oven (A, Fig. 108). A spacer for holding out the
syringe plunger in a fixed position (B, Fig. 108) is also required. It
consists of a piece of light sheet metal, about 1.5 cm. wide and 5.5
cm. long, folded into a V-shaped channel. The length of the spacer
allows a gas space of about 0.75 ml. in the syringe.

Srholander and Roughton Method for Carhon Dioxide

SPECIAL REAGENTS

Carbon Dio.ride-Free Distiller] Water. Boil the water after adding

a drop of sulfuric acid.
Caprylic Alcohol.

* See Bibliography Appendix, Ref . 52.

338 GASOMETRIC-VOLUMETRIC METHODS

Acid Phosphate Buffer. Same as for nitrogen (page 336). Carbon

dioxide has a low solubility in this soln.
10% Sodium Hydroxide.
Glycerol. ,

PROCEDURE

1. Lubricate the rear part of the dry plunger with a few streaks
of glycerol and place it in the syringe barrel which is moist with
water,

2. Fill the glass cup with distilled water, draw it one fourth down
the barrel, ind expel it through the cup. Leave the dead space in
the syringe full of the water without trapping any air bubbles.

3. Pull out the plunger slightly so that the water meniscus at
the bottom of the cup is lowered 1-2 mm. down into the capillary.

4. Hold the pipette against the opening of the capillary, trapping
a small air bubble, and draw the sample (13 [A.) down very slowly
into the capillary with the air separating it from the water.

5a. With the meniscus of the sample at the 30 or 35 mark, detach
the pipette and suck out the sample from the cup. Slowly move the
upper meniscus of the sample to the zero mark, and read the amount
of sample b (page 340) in divisions.

5b. As an alternative procedure, move the sample down to a
mark made at 33.3 divisions. Adjust the upper meniscus exactly to
the zero mark with a fine suction tip, keeping the lower meniscus
at the 33.3 mark. (In this way one third the normal pipette load is
used and the carbon dioxide vol. has only to be multiplied by 3 to
give the carbon dioxide content in volumes per cent after correc-
tions for temperature, etc.)

6. Deposit a drop of caprylic alcohol on the bottom of the cup
and eject the bubble of air above the sample through the caprylic
alcohol with the aid of a piece of fine wire if necessary.

7. Same as step 6 in the oxygen method (page 333) .

8. Fill the glass cup to the mark with acid phosphate and draw
it down very slowly until the upper meniscus is 2 mm. below the
bottom of the cup.

9. Moisten the rubber end of the wooden plug with phosphate
buffer and, with a few drops adhering to it, insert it in the bottom
of the cup, trapping a small air bubble.

10. With the plug resting loosely against the bottom of the cup,
gently move the air bubble up until it touches the rubber tip. Then

CARBON DIOXIDE

339

press the plug against the bottom of the cup, leaving the bubble in
contact with the rubber {A, Fig. 108) . Keep the capillary closed in
this way with the left hand throughout steps 11-17.

11. Place fresh glycerol in the plunger bearing.

12. Place one end of the metal spacer around the plunger under
the plunger head, and hold it there with the other end sticking out
at an angle. Keep the cup end of the syringe pointing upwards at a
slant through the steps 13-16.

(s>

i

Fig. 108. A, Rubberized wooden plug for
vacuum-sealing the glass cup. B, spacer for keep-
ing the plunger extended in a fixed position dur-
ing the vacuum extraction. From Scholander and
Roughtnn (1943b)

13. Slowly move the plunger out so that the fluid meniscus
under the stopper moves down the capillary very slowly. When the
capillary and its opening into the barrel are drained of fluid, slowly
draw the plunger out and move the free end of the spacer in until it
fits against the syringe barrel {B, Fig. 108) .

14. Add glycerol to the plunger bearing.

15. Shake the syringe for 2 min. with the cup end up to prevent
fluid from blocking the opening to the capillary. Should the capillary
become occluded, clear it by warming the capillary with the hand.
Should foaming occur, release the plunger and draw it out again.

16a. With the capillary free from fluid, pull out the plunger to
allow the spacer to fall out. Allow the plunger to rise rather rapidly
until the lower meniscus is inside the capillary and the gas pressure
is atmospheric.

16b. Should fluid bridge the opening to the capillary while the
plunger is let in, adjust the speed of the plunger so that the bridge
moves slowly up the capillary to enable proper drainage to occur.

17. Remove the plug and move the upper meniscus slowly and
evenly to the zero mark when the gas bubble is at atmospheric
pressure.

340 GASOMETRIC-VOLUMETRIC METHODS

18. Equilibrate the temperature and read the vol. (Fi) in divi-
sions as for oxygen (steps 13-14, pages 333-4). If liquid is in the
capillary, subtract the length of the liquid column from the total
reading.

19. Fill the cup with water and draw three fourths of it into the
syringe. Return the bubble to the capillary with water beneath it.

20. Fill the cup with 10% sodium hydroxide, with the cup point-
ing downward and the bubble expelled into the alkali, some of which
is drawn into the capillary as soon as the bubble is free.

21. To complete the carbon dioxide absorption, rotate the instru-
ment a few times with the cup pointing slightly downward. Then
with the instrument held vertically, cup down, suck the gas bubble
back into the capillary.

22. Again adjust the temperature and take a gas vol. reading
(¥2).

23. Calculate the carbon dioxide content in volume per cent from
the equation:

Carbon dioxide = (T^ — T'ol/dOO/b);'

where h is the vol. of sample (step 5a I, / is the gas correction factor
(page 334), and i is an empirical factor correcting for reabsorption
and incomplete extraction. For measurements on blood, i = 1.015.

3. Membrane Interferometer Volumetry

A sensitive indicator of pressure change which is based on the
principle of the interferometer was developed by Tobias (1942).
While the instrument has not yet been employed in biological
experimentation, a brief description will be given nevertheless, since
its further development and exploitation may be stimulated by so
doing. The instrument responds to pressure changes of the order of
0.05 mm. water, or about 0.004 mm. mercury. It consists of a pol-
ished microscope slide on which an ordinary cover slip (0.1 mm.
thick), having a hole ground through it about 2 mm. in dian:ieter,
is fastened with heavy shellac. A film of collodion is placed over the
cover slip and then another cover slip with a hole drilled through it
is fitted on top so that the holes of both cover slips coincide. Shellac
is also used to fix the positions of the film and the top cover slip.
The assembly is shown in Figure 109. When placed on the stage of
a low-power microscope, and illuminated with monochromatic light.

MEMBRANE -INTERFEROMETER VOLUMETRY

341

displacement of the film with respect to the glass slide can be fol-
lowed by recording the magnitude of the shift of the interference
fringes produced. A movable ocular micrometer may be used to
measure this shift.

Microscope -

nj

Mirror

(D

Source

Shellac

Membrane

Microscope slide

Cover slips

Fig. 109. Membrane inter-
ferometer manometer.
Froyn Tobias (1942)

Adjustable
diaphragm

Fig. 110. Detail of membrane inter-
ferometer manometer, showing that
fringes maj^ be obtained by reflected
light by shifting source, diaphragm,
and mirror from below to above the
interferometer.

Frum Tohins (19.^2)

The sensitivity of the instrument may be increased by employing
shorter wavelengths of light. Tobias used the green mercury line at
546 m/i, after the light was passed through a Corning filter. The use
of a diaphragm to narrow the light beam lends sharpness to the
fringes. By placing a glass slide over the top cover slip, a differen-
tial manometer with closed chambers on each side of the membrane
is formed (as in Fig. 110). One of the chambers may be used as a
reaction vessel, or, as water may condense on the membrane, it
might be preferable to make connection between the chamber under
the film and a reaction chamber ground out of the slide a short
distance away by cutting out a segment in the cover slip to form a
capillary channel between the two.

Drilling Holes in Cover Slips. A pile of ten to twelve cover
slips are mounted in pitch on a heavy glass plate. Round holes are
drilled with a carborundum or silica pencil bit operated in a hand
drill using water as the lubricant. The pitch is removed by soaking
in xylol for a day. The serrated edges are no handicap, but they can
be rounded off by coating with several layers of a plastic.

Preparation of Membranes. The collodion films are made
from a mixture containing 25 ml. celloidin {Merck), 25 ml. amyl

342 GASOMETRIC-MANOMETRIC METHODS

acetate, and 0.5 ml. n-butyl phthalate. The thinnest fihn that Tobias
used and measured was about 20 m/t thick, although thinner films
proved adequate.

B. MANOMETRIC

Of the two manometric techniques which will be considered here,
Cartesian diver and optical lever manometry, the former holds the
more prominent position. The relative simplicity of the apparatus
and the great sensitivity and high precision of which it is capable
make the diver technique one of choice for many gasometric studies
on the histo- and cytochemical level. The diver has already been
applied to the gasometric determination of isolated enzymes and
other metabolic catalysts as well as to respirometry. Future devel-
opments can be expected to exploit extensively the many possibili-
ties of this versatile technique.

1. Cartesian Diver Manometry

The application of the Cartesian diver to the measurement of the
volume changes in small volumes of gas was conceived and first
elaborated by Linderstr0m-Lang (1937b). A short discussion of the
technique appeared in a subsequent report by Linderstr0m-Lang
and Glick (1-938). Certain technical changes were employed by
Boell, Needham, and Rogers (1939) and Boell, Koch, and Needham
(1939) in their noteworthy investigations dealing with the respira-
tion and anaerobic glycolysis of regions of the amphibian gastrula.
Other technical modifications were suggested by Rocher (1942,
1943). Theoretical aspects of Cartesian diver micromanometry have
been given a complete treatment by Linderstr0m-Lang (1942, 1943).
Following a study by Linderstr0m-Lang and Holter (1942) of the
diffusion of gases through liquid seals in the diver, Holter (1943)
presented a finely delineated description of diver technique which
contains a full complement of precise detail, and includes the refine-
ments and innovations subsequently introduced. Zeuthen (1943)
designed divers of even smaller capacities and worked out the details
of their use.

The principle of the diver technique lies in the fact that any
change in the amount of the gas in the diver, which is used as a

CARTESIAN DIVER MANOMETRY

343

reaction vessel, requires a corresponding charge in the pressure
necessary to hold the gas volume constant so that the diver will
remain submerged at a fixed level in the flotation medium surround-
ing it. Thus the pressure changes become measures of the changes in
the amount of gas in the diver.

Divers having gas volumes of 1-10 pi. are referred to as "^uL-
divers" by Holter (1943), and he calls the methods which employ
them "/i,l. -methods." This nomenclature has obvious advantages over
the use of ambiguous terms such as "ultramicro," "submicro," etc.

Fig. 111. Schematic drawing of
measuring apparatus and diver.

Apparatus :

A, rubber tubing

B, coarse screw

C, fine screw

D, manometer

E, flotation vessel

F, circular mark

G, connecting manifold
H, three-way tap

J, rubber pressure tubing
K, ground-glass joint.

Diver :

a, bottom drop

b, neck seal

c, mouth seal

d, gas phase.
From Holter (1943)

^^=^

(a) Microliter Diver Technique'^

A diagram of the diver and apparatus is shown in Figure 111
(diver equipment may be obtained from E. Petersen, Carlsberg
Laboratory). The reaction mixture (a) is placed in the bottom of
the diver, and gas evolution or absorption changes the amount of gas
confined under the neck seal (b). Gas changes cause the diver to
rise or sink in the flotation medium in the vessels (E), and the water
manometer (D) is adjusted to vary the air pressure over the surface
of the flotation medium in order to bring the divers to the mark
(F) chosen as the equilibrium position. In this fashion the diver is
used as a constant-volume gasometer.

* See Bibliography Appendix, Ref. 46.

344 GASOMETRIC-MANOMETRIC METHODS

Since the precision of the jjressure measurement is about 1 nun.
water, which corresponds to 0.0001 of the total pressure of 1 atmos-
phere, and the total volume of the type of diver commonly used is
about 10 /xl., the accuracy of the measurement of the gas volume in
the diver is about 0.001 fx\. Holter has pointed out that for profitable
work with these divers, the volume changes to be measured should
be of the order of 0.01 ix\. per hour.

(1) TEMPERATURE CONTROL

It is advantageous to carry out work with divers in a room the
temperature of which is not very different from that of the thermo-
stat used with the diver apparatus. While not absolutely necessary,
additional advantages are obtained if the room has an approxi-
mately constant temperature, since only the flotation vessels and the
air bottle, which is connected to one end of the manometer, can be
conveniently submerged in the thermostat. The manometer itself
and the connecting tubing are subject to variations in the room
temperature. Rocher (1942,1943) employed the arrangement shown
in Figure 112. The manometers are submerged in the thermostat, and
a cathetometer is used to take their readings.

The thermostat used for the apparatus in the Carlsberg Labora-
tory is maintained at a temperature 2° above that of the room,
which is kept at 21°, and the thermostat is regulated with a precision
of 0.01°. The stirrer for the thermostat water must run very quietly
and it should not be attached to the thermostat itself in order to
prevent its vibrations from being transmitted to the diver. A 40 watt
lamp may be used as the heating element for a thermostat of the
size used at the Carlsberg Laboratory (58 X 30 X 30 cm.). The
light from the lamp should be shaded from the eyes of the operator.
High-capacity heaters are to be avoided, since they continue to
supply heat for a time after the current is cut off. Lamps are by far
the best heaters.

Since the diver must be observed during the experiment, it is
necessary that the front wall of the thermostat be made of glass,
and the lighting must also be considered. A ground-glass plate,
sufficiently large to form a background for all the flotation vessels
may be placed immediately behind the thermostat, where it can be

MICROLITER DIVER TECHNIQUE

345

illuminated in a manner which will nut heat the bath. This requires
that the back of the thermostat also be made of glass. Local heat-
ing must be avoided at all costs. Hence, if it is necessary to have
strong illumination on the divers, as is the case when observations
are to be made of organisms placed in the diver for study, a special
arrangement for submerging the light source in the thermostat will
probably have to be made. For this purpose Holter employed lamps
placed in a trough made of ground-glass plates cemented together.
The trough was lowered into the thermostat and filled with water to
the water level of the thermostat. The water in the trough was
cooled by a cooling coil to keep the temperature a few tenths of one
degree below that of the thermostat.

Fig. 112. Arrangement of Cartesian
diver apparatus.

<S, Syringe to vary pressure

Mu, mercury manometer

AIw, water manometer

B, air bottle
From Rocher (1943)

Convection currents in the flotation vessel, which may result from
local heating, can be detected by pipetting a colored solution of the
same density as that of the medium into the flotation vessel to form
a colored layer in the bottom third. Any convection movement will
produce colored streaks in the medium.

(2) FLOTATION VESSELS

A flotation vessel and its clamp are illustrated in Figure 113. A
standard-taper ground-glass joint is used to connect the vessel
through rubber pressure tubing to the manifold which leads to the
manometer. All parts not surrounded by thermostat water, here as
elsewhere in the apparatus, must be made small and thick-walled to
reduce the air space. The angle of the conical bottom of the vessel
should not be too small, since it is well to reduce the surface of
contact between the end of the diver tail and the bottom of the
vessel to a minimum. Otherwise there is a tendency for the tail to
stick to the bottom.

346

GASOMETRIC-MANOMETRIC METHODS

The distance between the top of the diver, when the diver is at
the bottom of the vessel, and the circular mark (H) on the vessel
should be only a few mm. in order to minimize the movement of the
neck seals. The hydrostatic pressure on the diver should be kept
small; accordingly, the level of the medium should not be higher
than G in Figure 113.

Fig. 113. Flotation vessel.

A, ground-glass joint

B, brass block

C, supporting rod (in cross section)

D, brass rod

E, brass cuff

F, wing screws

G, surface of flotation medium
H, circular mark.

From Holier (1943)

The flotation vessels are mounted by means of B (Fig. 113) on a
brass rod running the length of the thermostat.

Observations of the diver may be facilitated by using a magnify-
ing glass or a low-power horizontal microscope mounted so that it
can be moved sidewise on a rail fastened to the outside of the
thermostat. With this arrangement the microscope can be shifted
for use with divers in the different flotation vessels. If the eyepiece
has a horizontal cross hair, this can serve to establish the equilib-
rium position of the diver; in this case the circular marks on the
vessels are superfluous.

(3) THE FLOTATION MEDIUM

The flotation medium originally used by the Carlsberg Laboratory
workers was an almost saturated solution of ammonium sulfate, and
the Cambridge group employed an 11 A^ solution of lithium chloride.

MICROLITER DIVER TECHNIQUE 347

An investigation of various solutions which might be used as media
led Holter (1943) to the choice of a sodium nitrate-sodium chloride
solution. As Holter pointed out, a proper medium should have low
viscosity, chemical stability, transparency, biological innocuity,
glass-wetting properties, low and reproducible surface tension, and
very little gas solubility. Holter's medium has proved satisfactory
over a long period of use.

Preparation of Holter Medium. Dissolve 27.2 g. sodium
nitrate, 13.7 g. sodium chloride, and 0.2 g sodium taurocholate in
59.1 ml. distilled water containing 3 drops of 0.1 N hydrochloric
acid. To a sample of the soln. add bromocresol purple and then 0.01
N hydrochloric acid to bring the color to that of the indicator at pH
5.8 to 6.0 in salt-free soln. From the quantity of acid required, cal-
culate the amount of 0.1 A" hydrochloric acid to add to the main
portion of the soln. to bring it to the same pH. Add the calculated
amount and filter the medium through lintless paper, refiltering the
first part of the filtrate. Any cloudiness appearing upon the addition
of the acid is due to precipitation of taurocholic acid and should
soon disappear. Check the density of the medium at the proper
temperature with an accuracy of 0.1%; a Mohr-Westphal balance
may be used.

When the gas in the diver is air and no glass stoppers are used in
the-neck seals to inhibit diffusion (see page 368) the medium should
be saturated with air as described on page 374. When other gases
are used in the diver, the flotation medium need not be saturated
with these gases but the diffusion losses may be impeded by mouth
seals and glass stoppers.

Dust is apt to contaminate the flotation medium after some time ;
hence, it is occasionally necessary to combine the contents of the
vessels, refilter the solution, check the density, and refill the vessels
with a pipette. The pipette is used to avoid getting the soln. on the
neck of the vessel where it might crystallize.

(4) THE CONNECTING MANIFOLD

The flotation vessels are connected to the manometer by means
of a thick-walled glass manifold tube (G, Fig. Ill), carrying about

348 GASOMETRIC-MANOMETRIC METHODS

eight side tubes which communicate with the flotation vessels
through pieces of rubber pressure tubing about 15 cm. long, and the
standard-taper joints (A'), which are slightly greased with vaseline.
Each of the side tubes (5 cm. long and 5 cm. apart) has a glass stop-
cock to enable one flotation vessel at a time to be connected to the
manometer. The three-way tap (H) enables either the flotation
vessels or the manometer to be connected to the air.

(5) THE MANOMETER

The glass manometer tube has an inside diameter of about 2 mm. ;
each arm is around 150 cm. long. The manometer fluid is adjusted
by means of the pressure regulator illustrated in Figure 114. This
pressure regulator has been found to be more practical than the
syringes originally used, chiefly because the latter tend to leak and
also give rise to air bubbles in the manometer tube. The pressure
regulator is made of a piece of rubber tubing, 30 cm. long, 2 mm. in
wall thickness, and 11 mm. in inside diameter, which lies in a metal
trough {F) . The pressure screws are held by bridges (K) fastened to
the trough. The metal cylinder (D) used for coarse adjustment is
15 cm. long and the width of the trough. The pitch of screw B
causes a 1 mm. motion of D for each half-turn, corresponding to a
pressure change of about 25 cm. in the manometer. The pitch of the
fine screw (C) is about half that of B and a half-turn results in a
pressure change of about 1 cm. The ends of the tubing (A) are
closed with rubber stoppers. The reservoir (G) has a 20 ml. capacity.
L and M are three-way taps, FI and J stopcocks, and E a metal shoe.

Manometer Fluid. Brodie solution is used as the manometer
fluid. It consists of 23 g. sodium chloride and 5 g. sodium tauro-
cholate dissolved in 500 ml. distilled water. Dyes may be used to
color the solution if desired, and it is well to store the solution over
a few crystals of thymol.

The manometer is filled by drawing the solution up through J into
G. The levels in the arms are then adjusted so that both menisci are
about at eye level between the screws B and C.

Air Bottle. In order to render the measurement of the equi-
librium pressure independent of barometric changes and, to a large
degree, of temperature changes in the thermostat, one end of the
manometer is connected by thick-walled capillary tubing to an air

MICROLITER DIVER TECHNIQUE

349

Fig. 114. Manometer and pressure regulator. At left: detail
of pressure regulator. From Holter (1943)

Fig. 115. Cartesian diver assembly in thermostat.
Fro)n Holter (1943)

350 GASOMETRIC-MANOMETRIC METHODS

bottle coini)letely submerged and held to the bottom of the thermo-
stat by lead weights (Fig. 115). The vohmie of the air bottle should
be as large as possible (2-4 1.).

(6) THE MICROLITER DIVER

Dimensions of the Diver. The size and shape of different types
of diver are shown in Figure 116. The "standard" divers are most
generally employed; they have the dimensions:

Inside neck diameter 0.8 to 0.9 mm.

Neck wall thickness 0.1 mm.

Neck length 8-10 ram

Bulb diameter 2-3 mm.

Tail length 5-8 mm.

Total volume 10-12 Mi-
Neck volume about 5 /ul .

Total weight 20-30 mg.

These dimensions have been found to be preferable to those previ-
ously used, chiefly because the longer and narrower neck enables
reduction in the diffusion of gas through the seals. Regardless of how
large the divers are made, the inside neck diameter should not exceed
1-1.5 mm., although seals can be placed conveniently in necks up to
2 mm. On the other hand, the diameter should not be less than 0.6
mm., because with smaller diameters the difficulty of placing neck
seals is increased, due to the greater surface tension. It may not be
possible to go down to 0.6 mm. in some cases because the pipettes
which must be used may be too large to enter the neck. Usually the
smallest neck widths have to be 0.65-0.75 mm.

In order to determine the minimum neck length, allow 1.5 mm.
between the seals, which, with their menisci, require about 1.5 mm.
each. The lowest seal must be about 1 mm. from the lower end of
the neck.

Cylindrical divers {C, Fig. 116) are capable of the greatest re-
duction in size (down to a total vol. of about 3/a1.) and the conical
divers (D) furnish the greatest accuracy by a combination of a
relatively small gas volume with a reaction mixture having a rela-
tively large surface. The standard (^4) and long-necked {B) divers
are also shown in the figure.

Making the Diver. Divers may be made according to the
method of Boell, Needham, and Rogers (1939) or as described by
Holter (1943). The technique of the former may be applied to

MICROLITER DIVER TECHXlQUE

351

relatively thick-walled divers having a total volume >10 ix\., al-
though this method involves a lack of economy of the carefully
selected and calibrated capillary glass tubing from which the divers
are made. Holter's method is more economical and it is far superior
for divers having a total volume <10 jjI.

GO

6

B

o

V

0

z\

D

o

5 mm.

Fig. 116. Different types of divers.
From Halter' (1943)

The following table has been given by Holter (1943, page 420)
to serve as a guide for the selection of glass capillaries suitable for
making divers :

Inside diameter of capillary
(width of diver neck), mm.

Wall thickness,
mm.

0.65 to 0.75.

0.75 to 0.85.

0.85 to 1.0 .

>1.0 .

.0.08 to 0.09
.0.09 to 0.10
.0.10 to 0.12
.0.12 to 0.15

The capillaries should be made from a stock of glass tubing hav-
ing a ratio of 1:10 between the inside diameter and the wall thick-
ness. The density of the glass should be measured, since this value
enters into the calibration of the divers. The waste glass left after
drawing the capillaries should be saved and used for making the
diver tails. Entrapped air bubbles, which may be visible only as
streaks, sometimes occur in the glass tubing. It is important that
the glass used for divers has no entrapped air, which would change
the density of the diver; accordingly, the glass should be tested by
fusing one end of the capillary into a ball and examining for air
bubbles.

352

GASOMETRIC-MANOMETRIC METHODS

To make a diver by the method of Boell et al.:

1, Fuse one end of a suitable length of capillary tubing and draw
out a solid tail as indicated in steps A-C, Figure 117. The tail must
be light enough not to lose its alignment when the bulb is being
blown.

A

A

Ki^

B

0 1 2 3 4 5 mm.

D

D ^^^v^'^^^^'■

Fig. 117. Making of diver

according to Boell, Needham, and Rogers.

Fro7n Holler (1943)

0 12 3 4 5 mm.

I 1 \ ] I I

Fig. 118. Making of diver
according to Holter.
From Holter (1943)

2. Thicken the walls above the tail (step D, Fig. 117).

3. Seal the open end and heat carefully over a micro flame
while rotating the tube so that a bulb will blow itself (step E, Fig.
117). The size of the bulb will be determined by the vol. and tem-
perature of the air in the sealed unit.

4. Cut off the neck at the length desired.
To make a diver by the method of Holter:

1. Connect a long thin- walled piece of rubber tubing to one end
of the capillary chosen for the proper inside and outside diameters.

2. Fuse the other end of the capillary in a micro flame to form
a glass drop {A, Fig. 118) or, to economize on capillaries, use waste
glass to form a drop and then seal it to the end of the capillary.
Take care that no air bubbles are entrapped in the glass. The size
of the drop will determine the size of the bulb.

3. Rotate the capillary and hold it horizontally while heating
the drop in the flame. Carefully blow a bulb, moving the end out
of the flame so that the heat is applied only to the thick-walled
portion of the growing bulb (B, C, D, Fig. 118).

MICROLITER DI\TER TECHNIQUE 353

4. Fuse the end of a glass thread to form a drop, and fuse the
drop to the bottom of the bulb by heating the drop to a bright red
heat and the very bottom of the bulb to a medium red heat, and
then joining the two with a slight pressure.

5. Remove from the flame and pull out the glass tail at once
{E, Fig. 118). If the bulb is too hot, its shape will be spoiled while
pulling the tail; if too cold, the glass drop will not fuse into the
bottom and the tail will break off. Test the strength of the fusion
by applying a lateral pressure to the tail.

6. Cut off the excess at the neck and tail with a diamond point.
To prepare a conical diver [D, Fig. 116) heat only the side walls

of the bulb after the tail has been fused on, and collapse and draw
into the conical shape.

Cleaning the Diver. Holter ( 1943, page 429) has recommended
the following procedure for cleaning the diver after an experiment:

1. Rinse the outside of the diver with distilled water.

2. Remove the neck seals separately with soft filter paper cut
into short narrow strips and rolled between the fingers to form tight
smooth rolls. Take particular care to remove the paraffin oil as
completely as possible.

3. With a finely drawn pipette having a 2-3 ml. capacity, fill
the diver with glass-distilled water and blow about 2 ml. water
through the diver in a brisk stream. Stop before air gets into the
diver so that filling with the next liquid will be easier.

4. Repeat the washing with acetone, toluol, acetone, and twice
with glass-distilled water in the order given. Leave the toluol in the
diver for a few min. before replacing it with acetone. If the diver
had a neck ring of wax, rinse two to three times with toluol.

If the diver is unusually dirty, treat as described and then fill it
with freshly prepared 1% potassium permanganate in cone, sul-
furic acid, and let stand overnight. Replace the solution with water,
wash with strong hydrochloric acid, and flush well with glass-dis-
tilled water.

5. Blow air through the diver to remove as much water as
possible after the final washing.

6. Dry the outside of the diver with filter paper and place in an
oven at 120°.

7. Holding the diver by the tail with forceps, heat only the
neck of the diver by moving it slowly two to three times through

354 GASOMETRIC-MANOMETRIC METHODS

a micro flame. The flame should acquire a faint sodium tinge.

This process can be replaced by heating for about 20 min. at
400° in a furnace. It is necessary to heat the diver neck to insure
proper moistening of the neck by the seals.

8. Handle the clean divers only with clean forceps (cork-
tipped forceps are recommended). Clean divers should never be
touched with the fingers. Store the divers in stoppered glass tubes.

Afljusting the Weight of the Diver. The weight of the diver
must be adjusted so that when it is finally charged for an experi-
ment the equilibrium pressure will approximate the barometric
pressure. The desired weight of the diver may be calculated from
the formula:

9d =

1 — {<t>m/<t>gl)

where Qd is the desired weight of the diver, Fu- the aqueous volume
in the bottom and neck of the charged diver, V,. the total diver
volume, Voii the vol. of the paraffin oil seal, Vm the vol. of the
mouth seal, </>„„ </>(„7, 0„, 4)gi the densities of the medium, paraffin
oil, aqueous charge, and glass, respectively. The total volume [Vt)
is measured by weighing the diver to 0.1 mg., first empty, and then
filled with water. The densities of the liquids may be determined
with a Mohr-Westphal balance, and the glass density by weighing
2-5 g. of the glass in air and in water to 1 mg. If air bubbles are
present in the glass of the diver, the density can be obtained by the
flotation method. Holter has given the following proportions of
ethylene dibromide and bromoform to make flotation liquids of the
densities indicated:

Ethylene dibromide, Bromoform, j22.50

ml. ml. °40

20 5 2.314

15 10 2.460

15 15 2.529

10 20 2.643

Of course it is possible that different grades of the two compounds
will give mixtures with different densities. The following example of
the calculation of the desired weight of a diver has been taken from
Holter (1943, page 427) : "A diver made from Jena glass of specific
gravity 2.40 weighs 12.9 mg. empty and 23.6 mg. filled with water.

MICROL-ITER DIVER TECHNIQUE 355

It is to be used for the determination of the respiration intensity
of infusoria and is to contain the following charge: at bottom: 0.8
fA. of the infusoria culture medium; in the neck: 0.7 jxl. N/IO NaOH,
0.7 ^1. paraffin oil, and a mouth seal 2 mm. long. The inside diameter
of the diver's neck is 0.75 mm., its area 0.445 mm.-, hence the volume
of the mouth seal 0.89 [A. 0^ the density of the bottom drop and the
.V/10 NaOH is practically = 1, 0,,, = 0.87, 0,„ = 1.325, F, = 23.6-
12.9 = 10.7 lA.

10.7 X 1.325 - 0.7 X 0.87 - 1.5 X 1 - 0.89 X 1.325

yo =

1 - (1.325/2.40:
10.89

0.448

= 24.3 mg.

The weight of the diver may be adjusted by cutting off part of
the tail or fusing more glass onto it. If too heavy, a little more of
the tail is cut off than necessary and the diver is brought to the
proper weight, within 0.1-0.2 mg., by fusing on measured lengths
of glass thread of known diameter. Finally, the threads are fused
into a ball at the end of the tail, and the diver is dried and weighed
to 0.05 mg.

The weight of the diver is checked by charging it with the
quantities of the liquids chosen in the calculation, and determining
its equilibrium pressure. If properly prepared, the equilibrium
pressure will not deviate more than 20 cm. from the barometric
pressure.

Devices for Holding the Diver. The diver is charged by raising
it on a mechanical stand so that the tip of the pipette, which is
stationary, can be brought exactly to the position desired and the
contents of the pipette discharged at the proper place in the diver.
Special diver holders have been described by Holter which facilitate
the pipetting manipulations.

For divers with short necks (^ 1 mm. in width) the simple
rubber tubing holder, shown in Figure 119, may be used. For divers
with narrow necks, the centering with respect to the stem of the
pipette is more difficult, and, accordingly, Holter devised a clamp
stand which can be fixed relative to the pipette so that repeated
centering is not required. The clamp stand is shown in Figiu'es 120

356

GASOMETRIC-MANOMETRIC METHODS

and 121. The tongue on the left in Figure 121 moves on a horizontal
axis and presses the neck of the diver into a vertical V-shaped
groove in the head of the stationary post on the right. The edge of
the tongue is curved so that it touches the diver only at one point.
The tension needed to hold the tongue against the diver is derived
from a light spring which presses against the handle of the tongue

3 cm.

Fig.' ll'9. Diver support made of pressure tubing.
From Hotter (1943)

(Fig. 120). The entire clamp is fitted on the top of a rack and
pinion stand, which enables the diver to be raised or lowered evenly.
The base of the rack and pinion stand has leveling screws so that
the inclination of the diver can be controlled. A mirror fitted to the
side of the stand enables the operator to observe the centering of
the pipette from the side, and a magnifying glass attached to the
front of the stand or used in a head band permits the operator to
observe more easily from the front.

The diver is placed in the flotation vessel and removed from it
by means of a loop at the end of a piece of stainless steel wire
about 15 cm. long. The loop is made a little smaller than the diver
bulb so that the diver can rest on the loop with its tail hanging
down through the hole, and the loop is bent at right angles to the
wire so that the diver is upright when the wire is held vertically.

(7) THE PIPETTES

The pipettes used with the divers are straight glass capillaries
designed to measure 0.2-2 /xl. volumes with an accuracy of 1%.
They are filled and emptied by mouth through a piece of rubber
tubing. Four types of pipettes are used with microliter divers.

MICROLITER DIVER TECHNIQUE

357

Type 1 Pipette. This form of pipette (Fig. 122) is intended for
measuring dilute aqueous solutions, particularly for neck seals. The
pipettes are drawn from thermometer tubing having a bore of 0.2-
0.3 mm. The tubing is heated and the capillary widened into a bulb

Fig. 120. Clamp stand for divers. From Holier (1943)

having a wall thickness about half the diameter of the bulb. A
slight pull is applied to give the shape in A, Figure 122. At the
point indicated by the arrow, heat is applied with a micro flame
and the tubing is pulled out to give the pipette {B, Fig. 122). The

358

Fig. 121. Detail of clamp stand.
Fro77i Holler (1943)

I

10

I

15

20 mm.

r

:B

Fig. 122. Making of type 1 pipette, two successive stages.
From Holier (1943)

MICROLITEE DIVER TECHNIQUE 359

inside diameter of the tip of the pipette should be about 0.07-0.12
mm., and the outside diameter about 0.15-0.25 mm., depending on
the size of the diver employed. The delivery end of the pipette should
be strictly cylindrical for about 1 cm. from the end, or, even better,
slightly trumpet shaped at the tip. This form minimizes the tend-
ency of the liquid to creep up on the outside.

Type 2 Pipette. This pipette resembles the type 1 ; however, it
is intended for viscous liquids, particularly for paraffin oil. The
orifice at the tip must have a diameter no less than 0.15-0.18 mm.
and the glass wall thickness must be about 0.04 mm. In making the
pipette, the ratio of wall thickness to diameter of the bulb {A, Fig.
122) should be about 1:4, and the pipette is drawn to a tip with an
outside diameter of 0.2-0.25 mm. Paraffin oil has no tendency to
creep so that the exact shape of the tip is not important.

Type 3 Pipette.* This form of pipette is known as a "braking
pipette," and is intended for the transfer of cells and pieces of tissue
together with known amounts of liquid. The pipette (Fig. 123) is
drawn out at its upper end to a hair-thin tip so that the rate of
filling and emptying will be determined by the rate at which air
can pass through the very fine tube or ''brake" (E) . The outside
diameter of the pipette {A, Fig. 123) is 0.3-0.5 mm. The cork (B)
is smoothly cut in half lengthwise and a fine longitudinal groove
is made in the cut surface to hold the pipette. The rubber tubing
(D) is used for the operation of the -pipette by mouth. C in the
figure indicates the jacket tube.

In preparing the pipette, the upper end is drawn out long and
with a gradual tapering. Small portions are broken off with a fine
forceps until the rate of filling and discharge of the pipette is suit-
able. To test this, dip the mouth of the pipette into water and
observe the rate at which the water rises by capillarity. A rate of
about 1-5 mm./sec. has been found to be about right. The dimen-
sions of the delivery end of the pipette will be determined primarily
by the size of the object to be transferred. Whenever possible, taper
the mouth of the pipette, since it is difficult to reproduce the size
of the last droplet remaining in the mouth after delivery.

When in operation, should the braking tip of the pipette become
filled with condensed water which cannot be dislodged by sucking
or blowing, apply a slight excess pressure to the jacket tube and

*See Bibliography Appendix, Ref. 47.

360

GASOMETRIC-MANOMETRIC METHODS

close off the rubber tubing with a pinchcock. Heat the jacket tube
cautiously with a gas flame in the region of the braking tip; this
will cause the water in the tip to evaporate and the warm air will
clear the fine channel. If the tip becomes filled with a salt solution,
it will usually be necessary to draw a new tip. Obviously, it is
desirable that the pipette be rather long when first made.

D

A

E

B

Fig. 123.

Fig. 124.

Fig. 125.

Fig. 123. Braking pipette. From Holier (1943)
Fig. 124. Ball-tipped pipette. From Holler (1943)

Fig. 125. Ball joint for holding pipettes. A, Supporting discs; B, metal
springs; C, supporting rail. From Holler (1943)

Type 4 Pipette. The ball-tipped pipette (Fig. 124) is used to
deposit a drop of liquid at a definite place in a very narrow diver
neck (<0.7 mm. diameter) or in a fine capillary. The stem attached
to the ball has an outer diameter of 0.15-0.25 mm. and the ball
itself a diameter of 0.2-0.4 mm.

The pipette is made by drawing out a glass capillary from a piece
of tubing of medium wall thickness, sealing the tip by fusing, and
connecting the pipette with rubber pressure tubing to a source of
compressed air at 0.5 to 2 atmospheres (depending on the size of the
ball to be made). The sealed tip is brought close to a micro flame;
when the glass becomes soft enough it blows out into a ball which
perforates at the softest point.

MICROLITER DIVER TECHNIQUE 361

Devices for Holding Pipettes. In order to allow convenient
adjustment of pipettes, clamps fitted to universal ball joints were
used by Holter (Fig. 125). The clamps were mounted on a horizontal
bar, under which a ground-glass plate was fixed and illuminated
from behind to furnish favorable light for the pipetting. Pipettes
which must be removed from their clamps to obtain the sample, or
for any other reason, should be marked so that they can be re-
clamped at exactly the same place in order to avoid the necessity of
recentering them with respect to the diver neck when the diver is
held in its clamp stand. When it is desirable to use several pipettes
in quick succession with the same diver, the pipettes must all be
aligned to the diver, and the diver stand must rest on a surface
smooth enough to enable it to be pushed under the various pipettes
without losing the alignment. A level glass plate on the top of the
table is recommended.

Calibration of Pipettes. Pipettes of types 1 and 3 are calibrated
by iodometric (1 N potassium iodate) or acidimetric (1 N potas-
sium hydroxide) microtitration of a pipetted vol. of liquid using
0.01-0.02 A^ standard solutions for the titration. Type 2 pipettes
are calibrated by weighing paraffin oil delivered from the pipette
directly on to the pan of a microbalance.

Calibration markings on types 1, 2, and 4 pipettes are placed far
up on the pipette in the form of strips of millimeter paper. If the
pipettes are made from a broken thermometer, the graduations on
the glass will serve. A thin-walled and expanded portion of the
pipette, 5-10 mm. long, should lie between the fine stem and the
graduated part (Fig. 125). Type 3 pipettes require only one mark,
since they are used for complete discharge. No very satisfactory
means has been found for placing the mark when it must be so
near the end as to enter the diver neck during pipetting. Loops of
hair, held by a tiny bit of picein or glass cement, are usually too
bulky. Glass ink covered with lacquer is soon worn off. The Carls-
berg Laboratory group prefer no markings at all, but, after the
pipette has been filled according to judgment under a binocular
microscope, the length of the liquid column in the pipette is meas-
ured to 0.1 mm. by means of a micrometer scale. The pipette is
emptied into the bottom of the diver, and if the volume was mis-
judged, the difference is made up by correspondingly changing the
volume of one of the neck seals or the mouth seal. When the pipette

362 GASOMETRIC-MANOMETRIC METHODS

is narrow, the calibration mark will come high enough on the stem
to avoid contact with the diver, and in this case a hair loop or a
glass ink mark may be used.

(8) FILLING THE DIVER

The Bottom Drop. According to the calculations of Linder-
str0m-Lang (1943), the general rule follows that the thickness of
the layer of the bottom drop of reaction mixture must not exceed
0.5 mm. if the rate of gas diffusion through the drop is not to become
the limiting factor for the rate of the gas exchange to be measured.
This refers to gas changes in the range of those to be expected in
the usual biological systems (of the order of 0.01 ix\./hr.). For all
other cases, the equations given by Linderstr0m-Lang (1943) should
be applied to check the conditions.

It is essential that the bottom drop spread to reduce the thick-
ness of the liquid layer. The cleanliness of the diver is an important
factor. If the reaction mixture does not wet the glass, recourse may
be had to one of the following devices:

1. Addition of a surface-active substance, such as 0.1% sodium
taurocholate, provided that the substance will not interfere with
the reaction.

2. A very short centrifuging, or whirling by the arm while
holding the diver (the diver must be held with cloth or filter paper
to prevent the fingers from touching it) .

3. If centrifuging is not sufficient, and it is permissible in the
experiment, moistening the wall of the diver bulb before the bottom
drop is added by filling the bulb to within 1 mm. of the neck with
a suitable liquid and then sucking it out as completely as possible.
It will be necessary to correct the volume of the bottom drop for
any of the liquid not removed. When the moistened part is less
than 2 mm. from the lowest neck seal there is danger from creeping;
this may be obviated by a wax ring, described on page 363.

In order to avoid injuring cells, or damaging the tip of the braking
pipette used to transfer them by having the tip touch the glass
bottom of the diver, a small amount of a suitable liquid (about
0.1 [xl.], termed a "forerunner" by Holter, is placed into the bottom
of the diver first. The "forerunner" is introduced with a slender
flexible pipette which will not break if it touches the bottom of the
diver. It is necessary to avoid evaporation of the "forerunner" by

MICR0L.1TER. DIVER TECHNIQUE 363

keeping the diver in a moist chamber until time for adding the main
bottom drop.

Should the biological object be so large that it cannot be trans-
ferred into the diver by means of a pipette, the diver is filled com-
pletely with the bottom drop solution, the diver is lowered into a
dish containing the object, and the object is led into the mouth of
the diver. Then the diver is placed upright and the object sinks
to the bottom. The excess solution is pipetted out, the outside of the
diver is dried and small rolls of lintless filter paper are used to dry
the inside of the diver neck, and finally the weight of the remaining
liquid plus the object is determined after a neck seal of paraffin
oil has been placed to prevent evaporation.

To improve the visualization of the object in the bottom drop
during the pipetting, a drop of water is used to make a small piece
of cover slip adhere to the diver bulb after the diver has been
placed in the clamp stand. When the diver is in the thermostat, the
best view of the object may be had by using a cylindrical diver
or placing the object in a neck seal if this is possible.

In order to remove an object from a diver without injury, the
neck of the diver is first rinsed as described on pages 364-365 ; then
the whole diver is filled with a suitable liquid, the diver is placed
in the dish intended for the object, and the object is allowed to
float or sink out of the diver. A slender pipette filled with the liquid
may be used to flush the object out of the diver.

Wax Rings. There is great danger that the bottom drop will
creep up and be drawn into the capillary space between the stem
of the pipette and the diver neck when very small divers are used.
Proper centering of the pipette may not be sufficient to avoid the
difficulty. In such a case, a ring of beeswax about 0.5 mm. wide
is placed at the base of the diver neck. This will also prevent the
neck seal from creeping down.

To place the wax ring an electrically heated platinum loop (Fig.
126) is used. When the diameter of the neck is greater than 1 mm.,
carry out the following operation by hand:

1. Heat the loop and fill it with wax.

2. Allow to cool and place in the diver neck.

3. Apply the ring where desired by heating to melt the wax and
rotating the loop to form a ring.

4. Move the loop away from the glass wall and let it cool.

364 GASOMETRIC-MANOMETRIC METHODS

5. Remove the loop from the neck.

For divers with smaller necks, the loop apparatus is clamped
horizontally, the diver is held horizontally in a diver clamp, and
the wax ring is placed by rotating the diver smoothly about its
long axis. A dissecting microscope assists in the observation and
control of the operation.

Neck Seals. Neck seals are placed by bringing the end of the
pipette (type 1 or 2) into the neck so that the pipette stem is care-
fully centered. Then, by blowing cautiously a free-hanging drop
is formed which slowly grows {A, Fig. 127) until it touches the
neck wall (B) and finally forms the seal (C). If the centering is
inexact the liquid may spread between the pipette and the neck wall.

With wide necks it may be necessary to add more liquid to form
the seal than is desired in the final seal. Should this be the case, the
tip of the pipette is raised from the bottom meniscus to the interior
of the seal, and the excess liquid is drawn up into the pipette. Hence,
pipettes used for neck seals must either have two marks, the vol-
ume between them being that desired in the final seal, or they must
have calibrated cylindrical bores such as are obtained when the
pipettes are made from broken thermometers.

When coarse-tipped pipettes are employed or the divers have very
narrow necks, care must be exercised in removing the pipette from
the seal to prevent pulling up the liquid. The pipette should be
drawn out of the seal with a jerk. Similarly, when the ball-tipped
pipette is used, the ball should be removed from the drop deposited
by carefully moving the ball until it is just underneath the surface
of the drop, and then pulling it out with a jerk.

When it is necessary to exchange neck seals without disturbing
the bottom drop, the seals are first carefully removed with small
rolls of filter paper, soft paper rolls being used to absorb most of
the liquid; hard lintless paper rolls which almost fill the neck are
used to effect the final cleaning and drying. The neck is then filled
with water by bringing the tip of the pipette to a point about 1 mm.
below the bottom neck seal and blowing out the water very care-
fully. Once the neck is filled, the cautious blowing is continued to
rinse out the neck with a slow stream of water. Finally the water is
removed from the neck and filter paper is used for drying. There
is danger of evaporation of the bottom drop as long as there is no
oil seal present, therefore the exchange of seals must take place

MICROLITER DIVER TECHNIQUE

365

as rapidly as possible. When it is especially important to guard
against evaporation of the bottom drop, the operations should be
performed in a moist chamber.

0 1 2 3 4 5 cm.

Q

D

0

A B

\ /

3 mm.

Fig. 126. Platinum loops for placing
wax rings. Above, general appearance;
below, detail. A, glass tube; B, 0.5 mm.
platinum wire; C, loop of 0.2 mm. plati-
num wire; d, diver. B and C are welded Fig. 127. Formation of neck seal;
together. Current needed is 2-6 amp. three successive stages.

From Holler (1943) From Holier (1943)

In general, the neck seals should not be less than 0.5 mm. in
length, i.e., the distance between the apexes of the opposing menisci,
and there should be at least 1 mm. of dry glass between the seals to
prevent their mixing. Oil seals should be about 0.5 mm. long and
other seals about 1 mm. The distance between the oil seal and the
mouth seal should always be as small as possible in order to keep
the volume of the air space between them as small a fraction of
the total gas volume as possible ( Linderstr0m-Lang, 1943).

Aqueous Neck Seals. To insure proper wetting of the neck wall
by an aqueous seal, it should contain 0.1% sodium taurocholate
if the compound can be used in the experiment. The aqueous neck
seal should also be isotonic with the bottom drop to avoid distilla-
tion. In respiration work a seal of 0.1 A'' sodium hydroxide may be
used, since the difference in vapor tension between this solution
and water is small enough to be unimportant.

Should liquid be drawn along the diver neck accidentally, when
placing the aqueous seal, and if it is not feasible to repeat the filling,
the moistened area may be dried with filter paper. It is particularly
important that no moisture be present on the neck above the oil
seal since the water would distill into the mouth seal during the
experiment.

366 GASOMETRIC-MANOMETRIC METHODS

The Paraffin Oil Seal. Highly refined colorless paraffin oil
having a viscosity of about 20 Engler degrees at 20° and a density
of about 0.87 will suffice for most purposes. Of many liquids tested,
this has been found to be the best for sealing off the reacting sys-
tem from the outside.

Seals Containing Living Organisms. In order to place a living
object in a seal without damage, it is first necessary to place a seal
of the aqueous medium in the neck and then introduce the object
into the seal with a braking pipette.

The Mouth Seal. The mouth seal serves to prevent the loss of
gas from the diver, and it enables an adjustment of the equilibrium
pressure of the diver to a chosen value. In general, the length of
the seal must be several mm. to prevent significant gas loss, and
the length must be adjusted with an accuracy of 0.1 mm. to obtain
an equilibrium pressure within about 5-10 cm. of the desired value.
The length of the mouth seal is determined by measuring the dis-
tance between its lower meniscus and the diver mouth by means of
a traveling microscope (Fig. 128). The mouth seal is made as
follows:

1. After placing the oil seal, bring the tip of an "air pipette"
(a braking pipette with a finely drawn delivery tip having an inside
diameter of about 100 /a and an outside diameter of about 150 jx)
into the diver neck to a point about 1 mm. below the position at
which the lower meniscus of the mouth seal is to be placed.

2. Adjust the measuring microscope until the horizontal cross
hair in the eyepiece coincides with the top point on the rim of the
diver's mouth. Then lower the microscope by means of the mi-
crometer screw until the cross hair comes to the point where the
bottom meniscus of the seal is to'be.

3. Place a seal of diluted flotation medium around the stem of
the braking pipette at the mouth of the diver with a fine hand
pipette. (The diluted medium is employed to avoid crystallization
in the mouth seal. It is prepared by diluting the ordinary medium
with an equal volume of water and adding sodium taurocholate to
0.1 to 0.2 % to insure that the menisci will be properly formed.)

4. Apply suction to the braking pipette to draw the seal down
until the bottom meniscus reaches a position about 0.5 mm. below
the microscope cross hair; then raise the seal by blowing until the

MICROLITER DIVER TECHNIQUE 367

meniscus comes to rest at the position of the microscope cross hair.
The appearance of the seal at this stage should be that shown in
Figure 128. (By drawing the seal below the final position, the glass
walls will be moistened and the lower meniscus will not be distorted.
The upper meniscus should also be well developed and for this
reason the seal should not reach the rim of the neck. When the two
menisci equally oppose one another's surface tension, the mouth
seal will stay in place.)

5. Lower the diver stand quickly to remove the "air pipette"
from the diver, and fill up the mouth seal to the brim. Immediately
remove the solution which partly fills the air pipette when it is
taken out of the seal, and rinse with distilled water to prevent
clogging.

Fig. 128. Placing of mouth seal.
Frnm Holter (1943)

When the diver is submerged, the diluted medium in the seal is
displaced by the undiluted medium in the flotation vessel. This
displacement will be complete in a few min. with mouth seals 1-1.5
mm. long and 1 mm. in diameter. Should it be necessary to establish
the initial equilibrium pressure more quickly, and if the mouth seal
must be long and narrow, the equalization of the concentration of
the liquid in the seal must be speeded by displacing the liquid with
the flotation medium by rinsing. The rinsing process is carried out
by first drawing up into a glass tube, which has been drawn out to
a capillary about 10 mm. long and 0.2 mm. inside diameter, about 1
ml. of medium. Then the tip of the capillary is introduced into the
mouth seal and the medium in the tube is blown in to displace that
in the seal.

If crystallization occurs in the mouth seal, equalization of the
diver will be delayed for hours, and displacement of the liquid by
rinsing may not help since crystals sometimes hide in the meniscus
edges and escape the rinsing current.

The length of the mouth seal and the diver charge must be chosen
to give the diver the weight required for it to have an initial
equilibrium pressure of about 1 atmosphere. The weight may be

368 GASOMETRIC-MANOMETRIC METHODS

adjusted by changing the length of the mouth seal. Should it be
necessary to change the volume of other liquids in the diver, the
equilibrium pressure can be kept constant by changing the volume
of the mouth seal according to the relationship:

1 vol. aqueous soln. (d = 1) = 0.76 vol. medium (d = 1.325)
1 vol. paraffin oil (d = 0.87) = 0.64 vol. medium (d = 1.325)

In order to replace the mouth seal with one of another vol., the
diver is removed from the flotation medium, rinsed on the outside
with distilled water, and dried with cloth or filter paper. Then the
mouth seal is removed with a pipette or filter paper rolls, and
replaced with a seal of water extending into the diver deeper than
the previous seal. This water seal is flushed with fresh water by
the rinsing technique (page 367). The water is removed, the
mouth is dried with hard filter paper, and the new seal of medium is
introduced.

The length of the mouth seal may also be corrected by diluting
it with water, removing the excess with a roll of filter paper, and
then adjusting the position by means of an "air pipette" in a manner
similar to that employed when dealing with divers in anaerobic
experiments (page 373).

Seal Stoppers. Linderstr0m-Lang and Holter (1942) pointed
out that the diffusion of gas through seals can be reduced consider-
ably by placing glass stoppers into the seal. The stopper consists
of a bit of glass rod having a diameter about 50-80 fi less than
that of the inside of the diver neck. The length of the stopper will
depend on the magnitude of the effect desired, but in general they
are used 2-3 mm. long.

The solid stopper is the easiest to make but its weight may be
too great, making it more feasible to use hollow stoppers. Solid
stoppers can only be used with divers having a thin-walled neck
and a low center of gravity, since otherwise they render the diver
top-heavy. The hollow stoppers are made under a dissecting micro-
scope from a glass capillary of the proper outside diameter by fus-
ing one end shut in a micro flame, cutting off the capillary 0.5 mm.
longer than the stopper is to be, and fusing the open end shut. The
hot sealed end begins to bulge out due to the internal pressure

MICROLITER BIYER TECHNIQUE 369

of the heated air, so it is necessary to remove it from the flame be-
fore the enlargement exceeds 10-15 fi.

A stopper is placed in an oil seal by introducing it into the diver
with fine forceps and dropping it into the oil seal. A bubble, usually
found under the stopper, is removed and the stopper is completely
submerged at the same time by pushing the stopper down with the
tip of the oil pipette until the bubble bursts at the lower meniscus.
To remove a stopper from an oil seal, the mouth seal is first re-
moved, and the mouth of the diver is rinsed with water and dried
with filter paper rolls. Then the whole space over the oil seal is filled
with oil and the diver is inverted into oil to let the stopper fall out.

A stopper is placed in the mouth seal by the following procedure:

1. Place the fully charged diver in the flotation vessel where it
will remain at the surface because it has been calibrated to bear a
stopper and is therefore too light.

2. Rinse the mouth seal with medium (page 367).

3. Pick up the stopper with the forceps especially designed to
handle it (Fig. 129) and place the tip of the stopper which protrudes
from the forceps into the mouth of the diver. Linderstr0m-Lang
and Holter (1942) describe the forceps as follows:

5 cm.

Fig. 129. Forceps for placing glass stoppers.
From Holter (1943)

"It consists of a piece of thin metal tube, 15 cm. long and 2-5 mm. outside
diameter, which encloses a metal pin, 1.5 mm in diameter, 2 cm. longer than
the tubing and ending in a knob. On its lower end the tube carries two narrow
sheets of springy metal, 2 mm. wide and 2 cm. long, which are soldered on to
the outside of the tube in such a fashion as to protrude 15. mm. over the mouth
of the tube, parallel to its axis. By a slight bend the sheets are made to meet
at their ends, thus forming a pair of tweezers. Between the knob at the upper
end of the pin and some kind of button near the upper end of the tube lies
a spring which, when at rest, keeps the pin partially lifted out of the tube. By
pressing the knob of the pin down one forces the tweezers apart, thus releasing
any object which they have held."

370

GASOMETRIC-MANOMETRIC METHODS

4. Slowly push the diver, with the stopper partway in its mouth,
down to the bottom of the flotation vessel.

5. Release the forceps and push the stopper deeper into the
mouth with the tip of the forceps. The diver will not rise if the
charge has been properly calculated. However if the diver should
rise, push the stopper deeper into the mouth.

Hollow stoppers must not have a specific gravity less than that
of the medium or they will float out of the mouth seal. A stopper
is removed from a mouth seal by inverting the diver, dipping its
mouth into water, and letting the stopper fall out.

-^M-

A~

a

A~.

D

u o

a b

I

Fig. 130. Various arrangements for respiration measurements.

From Holler (1943)

In the figure, 1 shows the cylindrical diver for use when the main require-
ment is small volume : a, gas exchange between R and gas phase sufficient, also
when the surface of R is small; b, the gas exchange requires a large surface of
R (only possible in case of oi-ganisms that can rest on an air meniscus). 2 shows
the flask-shaped diver for use when the main requirement is a large surface
of R and when air support is not tolerated by the organism : c, relatively large
gas space; d, gas space as small as possible; the neck diameter and the length
of M depend on the importance of preventing the gas loss by diffusion. In 3 is
seen the glass stopper for use when the main requirement is that the diver be
as gas-tight as possible : e, glass stopper in 0 ; /, glass stopper in M.

The type of diver used and the arrangement of the seals will be
determined by the particular experiment to be performed. The
following basic set-ups (Fig. 130) were given by Holter (1943) for
a simple measurement of respiration which requires only one aqueous
neck seal (sodium hydroxide to absorb carbon dioxide): "Sugges-

MICROLITER DIVER TECHNIQUE 371

tions regarding the choice of diver type and arrangement of seals
under different experimental conditions. Assmned: A diver charge
like that used in the measurement of respiration, comprising a
reaction mixture (R) containing the organism, an absorption seal
(A), a seal of paraffin oil (0), a mouth seal (M). The gas phase
consists of air."

(9) FILLING THE DIVER UNDER ANAEROBIC CONDITIONS

In their studies on anaerobic respiration in the amphibian embryo,
Boell, Needham, and Rogers (1939) employed an apparatus which
permitted charging the diver in a nitrogen atmosphere. Holter (1943)
described an apparatus which has the advantage of permitting
small and narrow-necked divers to be filled more easily under
anaerobic conditions. The apparatus of Boell et al. is shown in
Figure 131, and that of Holter in Figure 132.

In Holter's apparatus (Fig. 132) the mercury vessel is mounted
on a stand with a rack and pinion so that it can be moved up and
down. E shows the base plate, F the diver clamp, and G the mercury
vessel. The glass tube {K) must be wide enough to allow the diver
to be moved horizontally to bring it under the various pipettes.
The gas enters through tubes C and D and goes out through the
bubble counter (L) . In the arrangement shown, pipette A is used for
an aqueous solution such as sodium hydroxide, pipette B for the oil
seal, and C to introduce the gas into the diver. The latter pipette
is drawn to an outside diameter of 0.3-0.5 mm. at the tip. The
pipettes must be drawn so that the fine stems are precisely coaxial
with the main tubes ; only when the stems are parallel to one another
will they all fit into the diver. The upper tubes of A, B, and C are
heated and cemented together with picein, and, if the pipette stems
are parallel, the tube D is joined to the group and they are all
sealed air-tight with picein or wax into a wide hole in the rubber
stopper. Rubber tubing is connected to the upper ends of the tubes.
The liquids which are to be pipetted are placed in small tubes drawn
out at the mouth to form a stem thin enough to be held by the diver
clamp.

372

Fig. 131. Apparatus for Fig. 132. Apparatus for

filling divers anaerobically. filling divers under

From Boell, Needham, and Rogers (1939) anaerobic conditions.

A is the filling chamber. "It bears a glass universal From Holler (1943)
joint, B, through the upper shank of which (C) a tube

D ending in an almost capillary point is fixed by means of the rubber con-
nexion, E. The pipette D is vaselined so as to be movable vertically as well as
from side to side. At its upper end it carries a three-way tap, F, and a teat, G.
The whole chamber is closed by the rubber stopper, P, through which an entry
tube (H) brings in the gas mixture (previously purified in the usual way by
passage over metallic copper in an electrically heated furnace). The entry tube
also has a three-way tap, /, so that part of the stream can be diverted to pass
through the pipette D. The exit tube is represented at K. We found it con-
venient to insert a wash-bottle between the electric furnace and the entry tube
//, so as to ensure that the entering gas mixture was not unduly dry, and also
a mercury trap in case of excess pressure. Inside the filling chamber there is
placed a diver-carrier (L) constructed of a small cardboard box and match-
sticks (also shown in the cross section and in perspective). This carrier bears
a wire arrangement to support two very small bowls, one for oil and one for
lithium chloride* (M and A''). These must be in such a position that they can
easily be reached by the movable pipette."

* Lithium chloride solution was used as the flotation medium.

MICROLITER DIVER TECHNIQUE 373

The gas is saturated with water vapor of the proper tension by-
passing through wash bottles of salt solution. The flow of gas
through C and D is controlled by pinchcocks, and bubble counters
are inserted. The gas stream through C is regulated to about one
bubble per second, and through D to about 2-3 bubbles per second.

The manipulations are carried out as follows:

1. Place the bottom drop in the diver before placing the diver in
the anaerobic filling apparatus.

2. Bring the diver into the apparatus and allow 5 min. for the
gas in the diver to become equalized with that in the chamber (K) .

3. During the latter part of the 5 minute period, bring the tip of
C into the bulb of the diver and allow the gas stream to flush
through the diver about 15 seconds.

4. Place the neck seals while the diver is in the apparatus, and
the mouth seal after the diver has been removed from the apparatus.
The diver is now ready for measurement.

In cases in which the exclusion of air is particularly important,
as in experiments on anaerobic metabolism of tissue having a low
oxygen consumption, place a drop of the diluted medium into the
mouth of the diver under anaerobic conditions using an extra
pipette for the purpose; this pipette need not be calibrated in any
way. Then remove the diver from the apparatus and draw the mouth
seal down into the neck by means of an "air pipette" (page 366)
as follows:

(a) To prevent air from getting into the diver, pass the gas
through the "air pipette" until the moment the pipette is to be used.

(6) The instant the "air pipette" is disconnected from the gas,
connect it to the tube which the operator holds in his mouth and
lower the tip of the pipette into the mouth seal. (A three-way stop-
cock connecting the air pipette with either the gas or the operator's
mouth facilitates the operation.)

(c) Blow down the liquid which rises in the pipette by capillar-
ity until the tip is completely, or almost completely, emptied. Then
pierce the lower meniscus of the seal with the tip and extend it into
the gas space under the seal.

(d) Suck out gas until the seal has been lowered to the desired
position,

(e) Withdraw the pipette and rinse it at once to prevent clogging
by evaporation and crystallization.

374 . GASOMETRIC-MANOMETRIC METHODS

A test of the anaerobiosos may be made, as suggested by Boell,
Needham, and Rogers (1939), by adding leucomethylene blue into
a bottom drop under nitrogen and then sealing the diver. The drop
will remain colorless for many hours if the filling has been properly
conducted.

(10) MEASUREMENT

Saturating the Flotation Medium with Gas. The flotation
medium should be saturated with air, unless, in a special case, it is
considered necessary to saturate it with another gas. The air satura-
tion should be carried out at least 1 hr. before introducing the diver
in order to allow time for all of the bubbles to disappear. The aera-
tion need not be performed before every experiment unless the
experiments are conducted at pressures deviating considerably from
the normal. The saturation is carried out by removing the flotation
vessel from the thermostat, sealing the mouth of the vessel with
the finger (not a stopper, since this would increase the air pressure
when inserted) , and shaking until the entire liquid is filled with air
bubbles. The vessel is replaced in the thermostat, and at 5 min. in-
tervals, the process is repeated twice.

If the medium is to be saturated with a gas other than air, the
gas should be washed first in some of the medium or a salt solution
of the same vapor tension and then passed through the medium to
be used in a stream of fine bubbles.

When the ground-glass joint is fitted to the top of the vessel, the
latter should be open to the outside through the manifold stopcock
and the tap (M), Fig. 114 (page 349). This avoids suddenly com-
pressing the air in the flotation vessel, which, if a diver is present,
will cause undesirable displacements of neck seals.

The measurement is conducted by the following procedure:

1. Connect the air bottle with the outside for a moment through
the threeway tap (L, Fig. 114), to equalize the pressure with that of
the atmosphere.

2. Place the filled diver in the flotation vessel by either dropping
it in, or, preferably, by lowering it in on the wire loop. Should air
bubbles be attached to the diver, remove them with the wire loop or
lift the diver out of the medium for a moment. If the mouth seal has
been made correctly, an air bubble will not form at the mouth of the
diver.

MICROLITER DR^R TECHNIQUE 375

3. With stopcock M (Fig. 114) open to the atmosphere, place the
ground-glass joint on the vessel and then turn the stopcock to con-
nect the vessel to the manometer.

4. Adjust the manometer to make the diver float, and see
whether the equilibrium pressure falls in the desired range. With
practice, one can judge the equilibrium pressure by the rate at
which the diver sinks. The position of the diver at the equilibrium
pressure is arbitrarily taken as that at which the top of the upper
air meniscus bordering the mouth seal coincides with the mark on
the vessel or the cross hair in the observing microscope.

5. Again check to make sure no air bubbles are on the surface of
the diver.

6. Rinse the mouth seal of the diver with medium as described
on page 367 if small effects are to be measured and especially if
narrow-neck divers with long seals are used. This need not be done
if large changes in gas vol. are to be measured.

7. Use the manometer to place each flotation vessel vmder the
pressure at which the divers are to be kept between measurements,
i.e., "the basic pressure." This pressure should be high enough to
prevent the diver from rising and touching the flotation vessel.
(Holter found that a pressure 3-5 cm. over the equilibrium pressure
of the diver at the time is usually sufficient if the room tempera-
ture does not vary much.)

8. At the proper times, make measurements of the equilibrium
pressure in the following manner:

(a) Adjust the manometer to the basic pressure before opening
the stopcock of the flotation vessel.

(b) Slowly reduce the pressure until the diver begins to rise.
Should the diver stick to the bottom so that it does not rise even
at a pressure of about 10 cm. less than the equilibrium pressure,
tap the neck of the flotation vessel lightly. Do not apply more
negative pressure to make the diver rise, since this may cause too
great a displacement of the neck seals. Tapping too hard may also
disturb the seals.

(c) Regulate the pressure so that the diver remains at the
equilibrium position for at least 10 sec, and then read the manom-
eter to 0.5 mm.

(d) Repeat the adjustment and reading. If the diver does not
change position during the time required for manometer readings

376 GASOMETRIC-MANOMETRIC METHODS

(about 30 sec), displace it by means of the fine screw on the
pressure regulator.

(e) If the two readings check, take the time of the second
adjustment within 1 min. as the time of reading.

note: Observe the shape of the menisci of the seals during measurement.
If they do not wet the neck properly they are apt to be deformed and the
accuracy of the measurement may suffer. It is sometimes helpful to vary
the pressure quickly to about 10 cm. on each side of the equilibrium value
to cause the menisci to move over the glass in their immediate locality
and thus wet it. Of course it is necessary as a rule to employ control divers
in each experiment. The use of the air bottle renders therm obarometer
divers superfluous.

(11) REVIEW OF THE EXPERIMENTAL PROCEDURE

As an example of the steps in an actual experiment, Holter (1943,
page 460) gave the procedure to be followed in a measurement of
the respiration rate of echinoderm eggs. The experiment described
was carried out in air, and two experimental divers and one control
diver were used. The following is Holter's own description (page
references, however, refer to this book) :

Choosing the Proper Size and Type of Diver. Since the assumed oxygen
consumption is large (about 5 X 10"'' /xl./hr.), the diver need not be smaller
than "standard size." The total gas volume of these divers being about 10 fj.\.,
the oxygen consumption will correspond to a change in equilibrium pressure of
about 50 mm. per hour. As the oxygen used in the reaction drop is supplied only
by diffusion, we need a reaction drop having a large surface and forming a
thin liquid layer (this excludes filling a in the schedule on page 000. Naked
echinoderm eggs do not stand direct contact with the surface of water (ex-
cludes filling b). No glass stopper is needed (excludes filhng e and /) since the
experiment is to be of short duration (2-3 hours) and since the pressure changes
are expected to be so large that a slight gas loss by diffusion through the diver's
mouth plays no appreciable role. On account of the magnitude of the antici-
pated effect we choose among fillings c and d the former, which also by its
better utiHzation of the area of the diver bottom, gives the largest surface of
the bottom drop.

Manipulations :

(1) If the flotation medium has not been in use for a long time the flota-
tion vessel is, not later than 1 hour before the beginning of the experiment,
shaken with air (page 374).

(2) The air bottle in the thermostat is connected to the outside air (page
374).

MICROLITER DIVER TECHNIQUE 377

(3) The divers to be used are heated (page 353).

(4) Each diver receives 0.1 ix\. of sea water as "forerunner" (page 362) ; if
this does not satisfactorily moisten the inside wall of the diver an amount of
sea water sufficient for such moistening is introduced into the diver bulb and
sucked out again until 0.1 imI. is left (page 362). Until ready for use the divers
are kept in an atmosphere saturated with sea water.

From this point on the description of the manipulations applies
to only one diver.

(5) The centering of the pipettes (page 361) for sodium hydroxide, paraffin
oil, and air (for placing the mouth seal) is checked and the two former pipettes
are filled. Of these the sodium hydroxide pipette is filled last, and it is to be
borne in mind that a small portion of the contents evaporates during the period
before pipetting off. NaOH isotonic with sea water! (page 365).

(6) Immediately before picking up the organism in the braking pipette (page
359) the diver is mounted in the clamp stand (page 357).

(7) The organism is picked up, and the water which enters the pipette
along with it is either adjusted to the mark or its length is measured (page
361).

(8) The braking pipette is mounted in its holder (page 361), the diver placed
under it, the centering checked, and the pipette is emptied into the forerunner.

(9) The sodium hydroxide seal and the oil seal are placed in the diver neck
(page 365).

If it is desired to check the contents of the diver under the microscope in
order to determine whether the organism has suffered by the pipetting it is
best done at this point, after the placing the oil seal.

(10) The mouth seal is placed (page 366).

(11) The diver is transferred to the thermostat, introduced into the flota-
tion vessel (page 374), and freed from any air bubbles that may stick to its
outside wall (page 374).

(12) The diver mouth is rinsed (page 375).

(13) The air bottle is shut against the outside air and manometer measure-
ments are started (page 375). Each measurement lasts about 3 minutes.

(14) Upon completion of the manometric measurements the diver is re-
moved from the thermostat and rinsed, whereupon the condition of the organ-
ism is checked under the microscope, best by submerging the entire diver in
water.

(15) The neck seals are removed, the diver neck rinsed (page 364) and the
organism removed from the diver (page 363).

(16) The diver is cleaned (page 353).
(17 The diver is dried (page 353).

It takes about 10-15 minutes to perform the manipulations 6-12. In a series
of experiments, therefore, this is the time interval between the initial ma-
nometer measurements of the two subsequent divers.

378 GASOMETRIC-MANOMETRIC METHODS

(12) CALCULATIONS

The Diver Constant. A constant must be determined for each
diver which represents the total gas volmne (V) in the diver when
the latter is at its equilibrium position at a given pressure (P) and
temperature (t). V may be measured in two ways:

(1) By subtracting the total volume of the liquids in the diver
from the total volume of the diver itself, and then correcting the
gas volume obtained at barometric pressure and room temperature
to P and t. The total volume of the diver is determined by weighing
or filling from a burette.

(2) By calculating V according to the formula:

y _ gp + Voil((}oil + Vw4>w — Voil<i>M — Vw4>M — igD4>Af/<t>gl)

where g, v, and 4> represent weight, volume, and density, respectively,
and subscripts D, oil, w, M, and gl refer to diver, oil, aqueous
phase, medium, and glass, respectively. This formula is derived from
the expression which is based on the fact that, at equilibrium, the
density of the diver unit equals that of the flotation medium :

'W

_ gp + Voil(t>oil + Vw(l>i,

y + Voii + v,r + gp/<i>ei

In practice, the first method is less exact than the second for
divers of the type described, because of the difficulty of measuring
exactly the volume of mouth seals and the total volume of small
divers by filling with a liquid. The chief reason for this is the poorly
defined surface of the liquid at the mouth.

According to Lindestr0m-Lang (1943), the errors of the values
in the formula for V which would result in a 1% error in V are:

Voii 33%

Vu, 50%

<t>,i 1%

<t>^f 0.5%

<i>oii, <t>w 12%

For divers having a total volume of <5 [A., an accuracy of 0.02
mg. in igr,) may be desirable, while for larger divers, an accuracy
of 0.1 mg. is sufficient as a rule.

MICROLITER DI\^R TECHNIQUE 379

Holier (1943, page 464) has explained that the calculation may
be simplified as follows: "Assuming that we are always using the
same medium (which is generally true) and that it has the density
(f)M, then the quantity go — {Qd 4>M/<i>ai) is a constant charac-
teristic for each diver and may be calculated once for all and
recorded in the diver inventory. Since, moreover, the same stock of
paraffin oil will be used in all experiments it is likewise possible
once for all to calculate the values of Vou 4>ou— ou <f>M and to plot
them as a function of Vou- The same applies to the volume of the
aqueous solutions, the densities of which in most cases deviate so
little from 1 that they may be ignored. With these two curves
drawn the whole calculation of V becomes a matter of reading the
values of V^ — Vw(J)m and Vou (pou — Vou (t>M from the curves, adding
them to Qd — (gD4>M/(t>oi) and dividing b}- the value of c^a/ which
is also known once for all."

Change in Gas Volume. The pressure change (Ap) is read
directly on the manometer. The equilibrium pressure (P) is actually
the manometric pressure (p), plus the barometric pressure at the
moment the air bottle was sealed, plus the hydrostatic pressure of
the medium over the diver, plus capillary pressure at the bound-
ary between the mouth seal and the gas in the diver neck. When
calculating the change in equilibrium pressure (aP), this quantity
may be made equal to (Ap) since all the other factors are essentially
constant. The surface tension factor is eliminated by using sodium
taurocholate in the medium. The relation Ap = aP requires the
correction :

P = p (1 + 1000 A/y/)

where A is the cross-sectional area of the bore of the manometer
tube, F/ the vol. of the air bottle, and 1000 the barometric pressure
in cm. of Brodie soln. For the dimensions of the manometer tube
and air bottle given previously (pages 348-350) the correction
amounts to about 1%.

When no correction is required for the solubility of the gas in the
liquids in the diver, the following relationship holds:

AV = FAP/Po

where Po is the normal pressure (1000 cm. Brodie soln.). However,
the effect of the gas solubility is appreciable in many instances, and

380 GASOMETRIC-MANOMETRIC METHODS

Holter (1943, page 466) has chosen the following cases from Linder-
str0m-Lang (1943) for which formulas for AV are given which are
simplified and adequate for practical purposes :

Case 1. The solubility of all gases present is low. This case comprises all ex-
periments in air, Na or O2 wherein CO2 does not occur or in which all the CL
formed disappears completely (respiration experiments in solutions without
carbonate buffers, in which the CO2 produced is absorbed by alkali). (Linder-
str0m-Lang 1943, page 369).

In this case we find for all experimental procedures which may come into
consideration in practice:

AV = VAP/Po

Case 2. The gas to be measured is soluble in the liquids of the diver charge
(CO2). The amount of this gas is small (^5% of the total gas) in proportion
to the other gases. The latter are sparingly soluble and do not change in quan-
tity. This case includes inter alia all experiments which are based on the cir-
cumstance that acid is formed or disappears in a system containing carbonate
buffer (Linderstr0m-Lang 1943, page 371).

In this case we find — under the assumption that the equilibrium pressure and
the "basic pressure" differ no more than 50 cm. from each other, that the vol-
ume of the oil seal is small {Yoii < 0.5m1), and that narrow-necked divers with
glass stoppers in the mouth seal are used:

' ^ V )

where Vw = the volume of the bottom drop, a'coi = the absorption coefficient
(not referred to 0", but to i°) of CO2 in the bottom drop at 760 mm. Hg and t"
(for water at 22.5°, has the value 0.89; for definition of q!'co2 (and /S'coj) see
Linderstr0m-Lang (1943, page 366).

The insertion of A F and AP for the original differentials dV and dP in-
stead of an integration is permissible also in this case, as shown by Linder-
str0m-Lang (1943, page 371), {i.e., the error involved is below 1%), provided
the pressure change A P does not exceed 50 cm., Brodie solution — which it
never does in practice.

Case 3. Like case 2, except that also one of the sparingly soluble gases (O2)
varies in quantity. This case includes respiration measurements in which the
CO2 evolved is not absorbed (Linderstr0m-Lang, 1943, pages 372 and 390).

In this case we find, under the same conditions as in case 2, augmented by
the condition v^ < 0.2 V, as well as under the assumption that the amount of
CO2 does not exceed 5% of the total gas:

^y^^ _,_ AFco^ ^ VAP

1 4- (Vwa'c02 + Voil /3C02)/F Pq

Where Vot and FcOz = the volumes of O2 and CO2 in the diver, /3cOi the
absorption coefficient (as above) of CO2 in paraffin oil at 760 mm. Hg and t°

MICROLITER DIVER TECHNIQUE

381

(at 24° z= 0.91; also valid with sufficient approximation at 22.5°); the other
designations as before.

The chief practical significance of this equation is that it permits of the
determination of respiratory quotients (calculation of A^cOa when V q^ is
known from a parallel determination according to case 1). Since the deter-
mination of R.Q. calls for the highest possible accuracy, the correction term
for the solubility of CO2 in paraffin oil (T^^jj Z^'cOz) i^ ^^^ disregarded in the
above equation (as in case 2), though here too Vg^l is small in relation to V.

The above formulas should be adequate for the calculation of most of the
experimental combinations occurring in practice. If in special cases some of the
experimental conditions to which the formulas correspond cannot be realized,
or if it is desired to use other gas combinations than those mentioned, then
the calculations of Linderstr0m-Lang (1943) will enable one to derive the
corresponding equations.

Sample Calculation. In an experiment on the measurement of the respira-
tion rate of the amoeba Chaos chaos Linne, Holter (1943, page 467) obtained
the following data:

Diver No.

Weight,
mg.

Total
vol., /ll.

0eJ

Mouth

diameter,

mm.

Length

of neck,

mm.

Length of

mouth seal,

mm.

16 (amoeba)
16a (control)

17 (amoeba)

18.07
14.70
16.09

18.1
14.7
16.1

2.40
2.40
2.40

0.69
0.71
0.71

10.0
10.3
12.0

2.6
3.0
3.05

Bottom drop, 0.4 /A. (Pringsheim soln. plus amoeba).
Lower neck seal, 0.4 fxl. (0.1 A'^ sodium hydroxide).
Paraffin oil seal, 0.4 fA.

The curves in Figure 133 were plotted by Holter to show the time course of
the oxygen consumption, and from the data he made the following calculations :

From the graphs:

Diver 16:

Diver 16: ^p (1 hr.) = 2.43 cm.
Diver 17: Ap (1 hr.) = 1.83 cm.
Diver 16a (control diver) = constant.

18.07 + 0.4-0.87 + 0.8-1.0 - 0.4-1.325 - 0.8-1.325 -

V =

18.07-1.325
2.40

O2 consumption = AF =

1.325
5.77-2.43 273

1000 273 + 22.5

= 5.77m1.
12.9- 10-3 Ml-/hr.

Diver 17:

V = 16.09 + 0.4-0.87 + 0.8 -1.0 - 0.4-1.325 - 0.8-1.325 -

16.09-1.325
2.40

0-2 consumption = AF

1.325

5.11-1.83

273

1000 273 + 22.5

= 5.11 m1.
8.6-10-3 juh/hr.

382

GASOMETRIC-MANOMETRIC METHODS

The slight initial bend of the respiration curve for diver 17 has nothing to
do with the respiration, it simply means that in case of this diver, which was
the last one to be placed in the thermostat, the measurements were begun be-
fore the initial equalization of i)rossure and temperature had ended.

Fig. 133. Respiration of amoebae: p = pressure.
From Holier (1943)

(6) 0.1 Microliter or Capillary Diver Technique

Zeuthen (1943) developed a new type of Cartesian diver for
respiration studies which has a gas volume about 100 times smaller
than the /xl. diver. These smaller divers, with a gas volume in the
range 0.04-0.11 /xl., were designated by Zeuthen as 0.1 (A. or capillary
divers. Their smaller volume leads to a refinement of the measure-
ment to the point where respiration intensities of 2 X 10"'* to
2 X 10"^ lA. oxygen per hour can be determined. In comparative
studies with several divers Zeuthen observed an error of measure-
ment of 2 X 10"^ lA. oxygen per hour, while variations in the respira-
tion of the individual cell during an experiment were much less.
Although the greatest respiration rate thus far measured in capil-
lary divers is 2 X 10"^ /xl. oxygen per hour, Zeuthen is of the
opinion that all rates less than 10"^ /xl. oxygen per hour should be
measured in capillary divers of suitable dimensions, while higher
rates should be followed in /xl. divers. Aside from the design of the
diver pipettes, and the flotation vessel, the apparatus is the same
as that used with the /xl. diver.

(1) THE CAPILLARY DIVER

Dimensions. The diver (Fig. 134) which is chosen as an
example has a gas vol. of 0.072 /xl. It is closed with seals of flotation
medium (Mi and Afo). The medium is that of Holter having a

CAPILLARY DIVER TECHNIQUE 383

specific gravity of 1.325 (page 347j ; however, it contains 0.5%
sodium taurocholate and has been made 0.1 N vi^ith respect to
sodium hydroxide by adding the calculated amount of 7.35 N sodium
hydroxide (29.4 g. sodium hydroxide in 100 g. soln.), which also
has a specific gravity of 1.325.

The length of the various columns in the diver is shown in Figure
134. In general, the lengths of the various components in the charged
diver should fall in the range (given in mm.) :

Seals of medium {Mi and M2) 0.7 to 1.0

Air Space (Ln) ca. 0.5

Solid paraffin seal (P) + paraffin oil seal (POi) 0.7 to 1.0

Water drop (W) containing respiring cell 0.2 to 0.3

Air space (L2) 1.5 to 2 times the diameter of the diver capillary

Oil seal (PO2), between menisci 0.02 to 0.03

Air space (L3) 2 to 3

The paraffin oil seals prevent loss of water from W. The solid paraf-
fin fixes the positions of the various colimms. The oil seal (POo) is
made short so that the carbon dioxide formed by the respiration can
diffuse from W to il/o becoming absorbed due to the alkalinity of
the latter.

Preparation of the Diver. The divers were first made in the
Carlsberg Laboratory from Thuringer glass, different samples of
which proved to be rather variable in property. Later Jena glass
was used and it was found that the glass drawn in an ordinary
flame (600-700°) was too brittle but that glass drawn at 1200-
1400° was suitable. The final test of the suitability of a glass is the
preparation and testing of control divers made from it. No informa-
tion is available at the moment concerning the properties of
American-made glass for divers.

The tubing from which the diver capillaries are drawn should be
thin-walled (outer diameter/inner diameter = about 1.25). Only
capillaries should be selected which have constant outer diameters,
as tested by a gauge such as the Zeiss cover slip gauge. For those
selected, the ratio of the weight of the glass to the weight of mercury
required to completely fill each tube is determined. With Jena glass
(sp.gr. ca. 2.40) Zeuthen reported that the following requirement
must be met for the weight of the mercury and glass: 9.2 >
mercury/ glass > 7.7, and with Thuringer glass: 10.0 > mercury/
glass > 8.3. Variations of the inner diameter must be checked by
moving a 5—10 mm. column of mercury along the capillary and

384

GASOMETRIC-MANOMETRIC METHODS

determining its length at different positions. Capillaries varying in
bore less than 2-3% and having lengths of more than 5 cm. should
be chosen. The inner diameter is finally measured by weighing the
mercury needed to fill the capillary completely. Divers are cut 5~6
mm. long, their length is measured, and then they are stored in
numbered test tubes.

I

^

Af2 = 0.80 mm.

13 = 2.46 mm.

PO2 = 0.02 mm.

^=0.30 mm.
P0i=0.30 mm
P=0.40 mm.

Li= 0.50 mm,
Mi = 0.90 mm.

Fig. 134.

1. The "standard diver":
length = 6 mm.,
diameter = 0.17 mm.,
gas space = 0.072 /ul.

The data in the figure apply to
lengths of the individual spaces and
seals.

2. Connecting piece between flotation
vessel and manometer.

•S. The flotation vessel.

From Zeuthen (1943)

Cleaning the Diver. Zeuthen has given the following procedure
for cleaning the diver:

1. Hold the diver in cork-tipped forceps under water and use a
thin glass rod to push out seals and air bubbles.

2. Place the diver in a cylinder of glass-distilled water and suck
the diver into a pipette having an inside diameter of about 1 mm.
for a distance of 2-3 cm. from the tip, and a constriction at the
2-3 cm. point so that the diver cannot pass. Continue to draw water
up into the pipette in order to flush the diver which is held at the
constriction.

3. Place the tip of the pipette at a slant against the bottom of

CAPILLARY DIVER TECHNIQUE 385

the water cylinder and blow out the water from the pipette while
retaining the diver in the pipette.

4. In a similar fashion, flush the diver with alcohol, toluol,
alcohol, and glass-distilled water in the order given.

5. Transfer the diver into the tube in which it is to be stored,
draw off excess water, and dry in an oven at 110-120°.

6. Never touch the cleaned diver with the fingers.

(2) THE FLOTATION VESSEL

The flotation vessel (Fig. 134) has been so designed that the diver
floats in medium enclosed between two air spaces, and the distance
between the upper and lower surfaces of the medium is such that
the ends of the diver are only about 0.5 mm. from the nearest air
space. With this design it has been found that the seals (Mi and Mo)
are effective in minimizing the exchange of air between the gas phase
of the diver and the medium, for the reasons discussed by Zeuthen
(1943, page 483).

(3) PIPETTES

Braking Pipette. The form of braking pipette required for
filling the capillary diver is shown in A, Figure 135. The capillary
(I) has an outer diameter of about 0.5 mm., and an inner diameter
a good deal less than 0.1-0.2 mm., and a length of about 3 cm. One
end of it is drawn out in a micro flame to an exceedingly fine bore,
which allows water to rise in the capillary at a rate of about 0.5 cm.
per sec. when the wide end is dipped into water to test it. The tube
(II), which is 1.5 mm. wide, holds I by means of a bit of DeKhotin-
sky cement. A thin-walled rubber tube is connected to II. A block of
soft transparent crude rubber (III) (a piece of red rubber labora-
tory tubing may be used) about 2X3X5 mm. is cut from a larger
piece after the rubber has been pierced with a needle and attached
to the wide end of I as shown. At the end away from I, the block is
cut at a slant to aid in finding the hole under the microscope. The
capillary diver is fitted into this hole using a binocular microscope
with good lighting to lessen the danger of breaking the diver during
the operation. The procedure to be used is as follows:

1. Pick up the diver by means of two watchmaker's forceps*
which have their points covered with cork.

2. Push the diver into the rubber block (III) until it almost
touches the glass of I.

386 GASOMETRIC-MANOMETRIC METHODS

3. Test for tight fitting by dipping the open end of the diver
into alcohol, blowing through the rubber tubing attached to II, and
then sucking a small drop of alcohol into the diver. If the drop is not
movable with equal ease in both directions, change the position of
the diver in III and repeat the test with alcohol until tight fitting
is obtained.

4. Finally, blow out the alcohol and allow the diver to dry for a
few min.

The Ball-Tipped Pipette. This pipette is very much the same
as the one used with microliter divers (page 360), except that it is
drawn out to an extremely fine thin-walled capillary. The ball is
formed by passing the pipette tip through a micro flame while com-
pressed air at 0.3-2 atmospheres is connected to the pipette. The
greater the pressure, the larger the hole that will be blown in the
ball, and the longer the flame, the larger the ball itself will be. The
dimensions, relative to the diver capillary are apparent from D,
Figure 135.

(4) FILLING THE CAPILLARY DIVER

Determining the Length of the Seals of Medium (Mi, Mo) to
be Used. First, the length of the column of medium {Ui) , which
placed in the diver will make the otherwise empty diver float, should
be calculated. It is most convenient to work with divers in which:

Im= (0.4 to 0.5) Z„

where Id is the length of the diver. The value of Im may be calculated
from the following formula which was derived from that given by
Linderstr0m-Lang (1943, page 363):

where / is the ratio of the weight of mercury to glass which was
determined in the selection of the diver capillary (page 383) and
(},gi and (j)M are the densities of the glass and medium, respectively
, (page 354) {cj,m = 1.325).

As will be described later, the columns W, POi, P, and PO-j
are first placed in the diver in the order given. Mi and M2 are
placed last. The following conversion factors are used for the cal-
culation of the combined length of Mi and M2:

CAPILLARY DIVER TECHNIQUE

387

1 mm. water replaces 0.75 mm. medium

1 mm. paraffin or paraffin oil replaces 0.65 mm. medium

Zeuthen (1943, page 501) employed the following example for a
diver having a diameter of 0.17 mm. and a charge as shown in A,
Figure 134:

lo = 5.87 mm.

/.If =: 2.15 mm. (calculated)

P -\- POr = 0.54 mm.

W = 0.26 mm.

1/2 = 0.37 mm.

PO2 = 0.05 mm.

P + POi + PO2 = 0.59 mm. (which replaces 0.38 mm. medium)

W — 0.26 mm. (which replaces 0.20 mm. medium)

Total: 0.58 mm. medium

Ml + Mz = 2.15 — 0.58 = 1.57 mm.
Ml chosen 0.82 mm.
il/2 chosen 0.75 mm.

I

M

^

1 cm.
A

1 cm

I cm.

5 r:im

D

Fig. 135. The techniciue of filling the diver.
Fro7n Zeuthen (19^)

"The equilibrium pressure of the diver is here 1 atmosphere —12.5 cm. H2O.
Most frequently, however, it will be found that the equilibrium pressure at the
first filling of the diver is rather far removed from 1 atmosphere. When the
diver floats it is possible to measure Mi and M2 accurately at the equilibrium
pressure. The figures found in this way are used in calculating Mi and M2 at
the next filling. As a rule we find that the diver can be filled in such a way that
the equilibrium pressure at the beginning of the experiment is 1 atmosphere
±40 cm. H2O (in recent experiments, though, 1 atmosphere ±20 cm. H2O)."

388 GASOMETRIC-MANOMETRIC METHODS

Placing IF, POi, and P. With the diver attached to the rubber
block of the pipette, the biological sample in its aqueous medium
W is drawn in, followed at once by the short seal of oil POi. The
filling is carried out under the microscope. The liquids should be in
small narrow cylinders which prevent the rubber block from coming
into contact with them. Speed is essential in bringing in the oil seal
after W has been taken in, since Zeuthen has found that if W is
exposed to evaporation for not more than 10 sec. at about 50%
humidity a loss of less than 10% of W will be incurred. It is advis-
able to work in a moist chamber in order to reduce evaporation
losses.

The paraffin seal (P) is next placed by pushing the end of the
diver through a layer of paraffin 0.2-0.3 mm. thick. The paraffin
(m.p. 58-60°) must be absolutely clear and white; the layer is
formed by kneading the material between the fingers. It is then
placed on a piece of rubber to minimize damage to the diver when
piercing the paraffin. The diver is held with the forceps and wiped
off to remove any paraffin or oil from the outside. A small blunt
glass rod which can go into the diver is used to push in the P,
POx, W combination about 1.2-1.5 mm. to leave space for Mx.
This must be done under the microscope and very slowly. Caution
is required to prevent the sample in W from coming into too inti-
mate contact with the air-water interface or particularly the oil-
water interface. Now the brake is clamped vertically and the length
of the diver is measured, if this has not been done previously. The
transparency of the rubber block permits the end of the diver to
be seen.

Placing PO2' A rubber stopper with dimensions approximately
the same as those in B, Figure 135, is fastened on its wide end to
a glass rod by which it may be clamped. A slit, 2-3 mm. deep, is
cut into the surface of the small end of the stopper and the end of
the diver is placed in this slit. With the diver mounted vertically in
this fashion, the ball-tipped pipette is clamped directly over it, and
the clamp holding the rubber stopper is raised so that the tip of the
pipette enters the diver. The ball is brought to the point where PO2
is to be placed and the oil is carefully blown out of the pipette to
form the seal. A slight jerk is used to disengage the ball from the
seal and then the pipette is withdrawn from the diver entirely. The
flexibility of the pipette stem makes strict alignment between the

CAPILLARY DIVER TECHNIQUE 389

pipette and diver less important than it is when microliter divers
are used.

The ball-tipped pipette may also be used to place oil in direct
contact with W, if desired. This is accomplished by moving the ball
very close to W so that the oil will be able to come into contact
with W before a separate seal can be formed. The thickness of such
an oil layer can be regulated by drawing back into the pipette any
excess oil.

Placing Mo. A glass tube is fitted on the rubber stopper (r) to
form a vessel around the diver as in C, Figure 135. The vessel is
filled with flotation medium to 2-3 mm. over the top of the diver.
In order to facilitate observation under the microscope, a square
piece of cover slip is cemented on the tube with Canada balsam and
the edges of the cover slip are held with DeKhotinsky cement to
prevent it from sliding off {E, Fig. 135).

M2 is then placed by withdrawing air from the end of the diver
through the tip of a pipette, arranged as shown in E, Figure 135.
As the tip is brought into the end of the diver, gentle suction is
applied so that the end of the tip is kept at the surface of the
medium. In this way, the length of Mo will be equal to the distance
the end of the pipette tip is brought into the diver.

Zeuthen (1943, page 506) has described the process of placing a
1.00 mm. M2 as follows:

"Before the pipette is introduced into the diver, a horizontal Une in the ocular
01 the micro.scope is brought to intersect with the image of the pipette 1.00 mm.
from its tip ; in placing il/n, the diver, encircling the pipette, is raised so much
that its edge is just level with the line chosen in the ocular. After making sure
that actually a current of medium has passed from the surrounding liquid
through the diver's neck into the pipette, the latter may be withdrawn from
the diver, emptied by blowing, and rinsed with water."

Placing Ml. The medium in the vessel surrounding the diver is
poured out; the diver is inverted and the M2 end is placed in the
slit. The vessel is again filled with medium, and the ensuing pro-
cedure is the same as that used for placing M2.

Transfer of Diver to Flotation Vessel.

1. Remove diver from the slit and allow to float in the medium
over the rubber stopper with the M2 end upward.

2. Draw the diver up into an ordinary pipette having a drawn-
out tip, along with about 0.5 ml. medium. Keep the ilfo end up.

390 GASOMETRIC-MANOMETRIC METHODS

3. Make the diver go up and down quickly a few times by suck-
ing and blowing to free it of air bubbles which might be adhering.

4. Pipette the diver and enough medium into the flotation vessel
so that the column of medium exceeds the length of the diver by
about 1 mm.

Replacement of Mo. Some evaporation of Mo during the filling
of the diver necessitates replacement of the medium in M2 in order
that its density be the same as that of the surrounding liquid. The
replacement is effected by applying alternate suction and pressure
three times to the flotation vessel. The air space (L3, Fig. 134),
which is next to M2, is relatively large and the expansions and
contractions of the air force the medium out and then into the end
of the diver. The effect on Mi is naturally very much less. The
process of filling the diver and placing it in the flotation vessel
takes 20-30 min.

(5) MEASUREMENT

For the observation of the diver during an experiment, Zeuthen
used the horizontal microscope with the vertical micrometer move-
ment illustrated in Figure 151. A particular mark on the diver, such
as one of the menisci, is made to coincide with a reference mark in
the ocular of the microscope to establish the equilibrium position.
For the finest measurements, the diver is kept in a floating position
throughout the experiment. This is not very difficult, since the
motion of the diver is quite slow when pressure changes on the
medium fall within 1-2 mm. (water) of the equilibrium pressure.
In experiments in which the greatest accuracy is not required, the
diver may be left to sink to the bottom of the medium after a
measurement. Should the diver stick to the lower medium-air inter-
face, making it difficult to raise it, gentle tapping of the flotation
vessel will free it. Before taking another reading, the diver should be
made to float for a fixed interval (about three minutes) to allow
time for complete equilibration of the pressure.

The equilibrium pressure is taken as the mean of the pressures
required to cause the diver just to begin to rise and to sink. The
two pressures are within 1-3 mm. water from one another, and the
readings require about one minute. While the accuracy of microliter
diver measurements is ±0.5 mm. water, that of the capillary diver
is ±1 mm.

CAPILLARY DIVER TECHNIQUE 391

After the experiment the diver is inimediatcly removed from the
flotation vessel by filling tli(^ latter with water and pouring out the
water with the diver into a small cylinder. Using a microscope, the
lengths of Lj, Lo, and L-? are measured, taking the greatest distance
from meniscus to meniscus. The boundary between Li and P is not
a meniscus; hence there will he five menisci in a diver such as
that in A, Figure 134.

(6) CALCULATION

Considering the menisci to be hemispherical, the gas volume T
of the diver at the barometric pressure prevailing at the end of the
experiment is:

V = {h + k + hUr-"- - -—

o

where I represents the length of L, and r is the radius of the diver
capillary.

At the floating position the volume is F X B/B — P2) , where B
is the barometric pressure (mm. water) and {B — po) the final
equilibrium pressure observed. No correction for the height of the
rise of the medium in the diver capillary is necessary.

When aP is the change in the equilibrium pressure (mm. w^ater),
the oxygen consumption (AT^) in a respiration experiment is:

AP B

AV = ——-■¥

10,300 B - p2
or corrected to standard conditions:

Ap B B -p, 273

av = • V

10,300 B - p2 10,300 273 + /

where {B — pi) represents the average pressure of measurement
during the given period.

Usually the corrections are small enough to render it sufficient to
calculate the change in vol. by the formula:

AV=V ^^

10,300

where aV and V are given in microliters and p in millimeters water
per hour.

392 GASOMETRIC— MANOMETRIC METHODS

(7) CHOICE OF DIVERS FOR DIFFERENT
RESPIRATION RATES

The choice of divers for measurements of respirations of approxi-
mately known intensities should follow the general considerations
given by Zeuthen (1943, page 509) :

1. The changes in equilibrium pressure should fall in the range
of 3-10 cm. water per hour (2-20 if necessary) .

2. The respiration rate must not be great enough to move (PO2)
toward W. This can be checked by observation through the micro-
scope.

3. Total changes in equilibrium pressure over 50 cm. water
(corresponding to a drop in oxygen in the diver from 21 to 16%)
should be avoided.

4. It is well to calculate, for the diver dimensions used, the
magnitude of the respiration rate which would result in the danger
of oxygen deficiency (perhaps even the carbon dioxide absorption
should be calculated according to the formulae of Zeuthen, 1943,
pages 492-494) . The following formula is used to relate the dimen-
sions to the respiration rate:

_ OAlAao, . <^o.

R o'lOj

where I (in mm.) is the greatest length of TT^ permissible for the
respiration intensity R (in /tl./hr.), A is the cross-sectional area of
the diver capillary, in mm.^ 0-02 is the "standard rate of passage"
for oxygen through water at 20° ; its value is 0.204. o-q is the
"standard rate of passage" for oxygen through paraffin oil to which
the value, 0.10, has been ascribed; and L^, in mm., is the average
length of the diffusion path through PO2, which is defined as Ld =
L -{- (2r/3), where L is the shortest distance between the oil menisci
and r is the radius of the diver capillary. Thus:

A I 2r\ 0.204

When there is a tendency to condensation of moisture on the
walls of L2, when W consists of fresh water, a film of oil about 0.05
mm. thick is placed on W . Then:

CAPILLARY DIVER TECHNIQUE 393

A

/ = - X 0.11 X 0.204 -
R

where {Lo) is the thickness of the oil film.

For a T7 of 0.2-0.3 mm. in the diver {A, Fig. 134) , lO'^ ^l./hr. is
the maximum respiration intensity which can be determined.

5. When the diameter of the diver is about 0.02-0.03 mm. greater
than the diameter of the cell, the change in equilibrium pressure
will usually be large enough even for weakly respiring cells. The
smallest glass divers which have been used have a diameter of 0.13
mm., and the smallest cells whose respiration can be measured
(about 10"^ /xl./hr.) will therefore be around 100 to 50 jx in diameter
for weak and strong respirations, respectively.

(c) Methods Other Than for Respiration

CHOLINESTERASE

Linderstr0m-Lang and Glick (1938) developed a Cartesian-diver
method for the measurement of cholinesterase based on the principle
of the method utilizing the Warburg apparatus (Ammon, 1933).
This principle depends on the fact that if the enzymatic scission of
a choline ester proceeds in a bicarbonate buffer, the acid liberated
will cause an equivalent evolution of carbon dioxide which can be
measured gasometrically. Some investigators use a bicarbonate-
Ringer medium while others employ only a bicarbonate solution for
the estimation of the enzyme. The advantage of the presence of the
other salts which are in the Ringer solution is that they activate
the enzyme. By substitution of other ester substrates, the present
method can be applied to the measurement of lipolytic enzymes,
atropinesterase, etc. For a titrimetric method see page 310.

Linder8tr0m-Lang and Glick Method for Cholinesterase

SPECIAL REAGENTS

Buffer Substrate Solution. Prepare 0.5% acetylcholine chloride in
bicarbonate-Ringer soln.

Bicarbonate Ringer Solution (pH 7.4). Add 2 ml. of 1.2% potas-
sium chloride, 2 ml. 1.76% calcium chloride (CaCl2.6HoO), and
20 ml., 1.26% sodium bicarbonate to 100 ml. 0.9% sodium

394 GASOMETRIC-MANOMETRIC METHODS

chloride. Saturate the soln. with a mixture of 5% carbon dioxide
and 95% nitrogen.

PROCEDURE

1. Pipette 1 ix\. buffer-substrate soln. into the bottom of a micro-
liter diver.

2. By means of a pipette extending nearly to the surface of the
soln. in the diver, pass a rapid stream of 5% carbon dioxide in
nitrogen through the diver for 30 sec.

3. Pipette 0.3 fxl. enzyme soln. into the bottom drop in the diver
and again pass the carbon dioxide-nitrogen mixture through the
dive]' .

4. Place the paraffin oil seal in the neck of the diver followed by
the mouth seal.

5. In a parallel fashion, prepare a control diver containing 0.3
jxl. inactivated enzyme or soln. without enzyme.

6. Proceed with the diver technique as described on pages 342-

382.

note: According to the work of Linderstr0m-Lang and Holter (1942),
the relativel.y high permeabihty of paraffin oil and flotation medium to
carbon dioxide would make it advisable to use oil and medium which are
saturated with the carbon dioxide-nitrogen gas mixture. The apparatus
illustrated in Figure 132 would be useful for charging the divers.

THIAMINE AND COCARBOXYLASE

The method of Ochoa and Peters (1938) for the estimation of
thiamine and cocarboxylase depends on the stimulating effect the
compounds have on the decarboxylation of pyruvic acid by alkaline-
washed yeast. Employing the Warburg technique these authors were
able to determine down to 0.01 /xg. cocarboxjdase and 0.05 [xg. thia-
mine. By adapting this method to the Cartesian /A. diver, Westen-
brink (1940) increased the sensitivity to the measurement of 0.05
ni/xg. cocarboxylase and 0.5 m^ag. thiamine. The sensitivity of the
method "v^aries with the variety of the yeast and the manner in
which it is washed. The thiamine determination may be carried out
most suitably when the concentration of cocarboxylase present is
small compared to that of the thiamine. For instance, in one experi-
ment, thiamine had no influence on the carbon dioxide evolution

THIAMINE AND COCARBOXYLASE 395

when more than 0.01 /tg. cocarboxylase was present; however. 0.001
fig. proved to be a favorable amount.

There is little reason why the Cartesian diver could not be applied
to the method of Atkin et al. (1939) and Schultz et al. (1942), which
depends on the stimulation of yeast fermentation by thiamine.

Westenbrink Method for Thiamine and Cocarboxylase

SPECIAL REAGENTS

Yeast Suspension. Stir 100 mg. yeast with 5 ml. 0.1 M secondary
sodium phosphate for 4 min. at 16-20°. Centrifuge 1 min.; dis-
card supernatant and repeat the procedure twice. Finally wash
residue once with 5 ml. water in the same manner. To the residue
now add 0.12 ml. of 0.1 M magnesium chloride and enough 0.1 M
phosphate buffer, pH 6.2, to bring the total vol. to 1.2 ml. Use soon,
since the treated yeast deteriorates rapidly even in the cold.

1.0 M Sodium Pyruvate in 0.1 M phosphate buffer, pH 6.2.

Thiamine (0.8 mg. per ml.) in 0.1 M phosphate buffer, pH 6.2.

Cocarboxylase (0.005 mg. per ml.) in 0.1 M phosphate buffer, pH
6.2.

PROCEDURE

1. Pipette into a Cartesian diver, having a vol. of 20-30 lA., 0.2 /il.
portions of the pyruvate, thiamine and cocarboxylase solns. When
thiamine is to be determined, use the cocarboxylase soln. with 0.2
/xl. unknown; and when cocarboxylase is to be measured use the
thiamine soln. with the unknown.

2. Set up a control experiment by replacing the unknown soln.
by the phosphate buffer.

3. Place divers containing the reagents in glass tubes closed by
rubber stoppers to prevent evaporation.

4. Prepare the yeast suspension and add 0.5 /xl. of it to the solns.
in the divers.

5. Seal the neck of each diver with 1.7 /xl. mineral oil; place diver
in the apparatus and take readings of the gas evolution after 30 min.

6. Calibrate the measurements by comparisons with those ob-
tained using known amounts of the constituent being analyzed.

39b GASOMETRIC-MANOMETRIC METHODS

7. For the determination of both components in a mixture, first
measm'e the cocarboxylase. Then determine the thiamine in a sepa-
rate experiment by adding, to the 0.2 /xl. imknown, 0.2 //,!. cocar-
boxylase soln. containing sufficient of the substance to make the
total in the diver 0.001 /xg.

DIPHOSPHOPYRIDINE NUCLEOTIDE

Anfinsen (1944) adapted the method of Jandorf, Klemperer, and
Hastings (1941) to the measurement of diphosphopyridine nucleo-
tide ( DPN) in microtome sections of tissue by the use of the Carte-
sian-diver technique. The method of Anfinsen permits the estima-
tion of 1-6 m/xg. DPN with an error of less than 5% ; this represents
a thousandfold increase in the sensitivity of the Warburg procedure
of Jandorf et al. The principle of the method lies in the fact that the
enzymatic conversion of hexose diphosphate to phosphoglyceric acid
and phosphoglycerol can be made to take place under conditions
in which the quantity of DPN present is the limiting factor. These
conditions require the presence of a muscle extract to supply en-
zymes and arsenate to limit specifically the glycolytic process. The
dearsenylation of the l-arseno-3-phosphoglyceric acid formed occurs
spantaneously. The molecule of acid produced in this step is made
manifest by the liberation of carbon dioxide from a bicarbonate
buffer; thus the gas evolution becomes a measure of the concentra-
tion of the DPN.

Anfinsen Method for Diphosphopyridine Nucleotide

Extraction of DPN from Tissue Sections. Since DPN is
rapidly destroyed by concomitant enzymes in the tissue, the follow-
ing procedure is employed for the extraction:

1. Weigh frozen-dried sections on a quartz torsion balance (page
191) and transfer to micro centrifuge tubes (Fig. 136 A), made of 3
or 4 mm. tubing.

2. Draw out tube to form a constriction {B).

3. Add a known amount of distilled water with a constriction
pipette (page 172) and rapidly seal the tube (C).

DIPHQSPHOPYRIDINE NUCLEOTIDE

397

^~^

.

^

%^_^ Wooden
35*"^ cylinder

1

o

i

A

pO

■ 1

1

-, 50 ml.

1 centrifuge
cup

y

9

1^' ^

i

^^

A

B

c

D

E

Fig. 136. Apparatus used in extraction of DPN from frozen dried sections:
A, micro centrifuge tube with frozen dried section; B, tube after addition of
water; C, tube sealed off during heat inactivation ; D, tube during extraction
of DPN. From Anfinsen (WW

4. Plunge sealed tube into boiling water, and, after 2 min., re-
move and chill well in ice water.

5. Centrifuge down drops of soln. left on the sides and top. The
special centrifuge cup (£") is convenient for this purpose. Open tube
with a file.

SPECIAL REAGENTS

O.lSIf M Sodium Bicarbonate saturated with a mixture of 5% carbon
dioxide and 95% nitrogen.

0.003 M Disodium Hydrogen Arsenate.

0.016 M (approx.) Sodium Hexose Diphosphate (o 1 mg. phospho-
rus/ml.). Prepare from the calcium salt by adding about 700 mg.
of the latter to 40 ml. of 1 % oxalic acid. After shaking neutralize
the mixture with sodium bicarbonate to chlorophenol red. De-
colorize with charcoal and filter. Test filtrate for oxalate, and, if
present, precipitate it with a little solid calcium hexose diphos-
phate and refilter through the original filter paper. Determine the
phosphorus content of the filtrate and dilute the soln. until it con-
tains 1 mg. organic phosphorus/ml. The presence of 3-4% in-
organic phosphorus may be ignored. Store in refrigerator where
the soln. w^ill be stable for several months.

398 GASOMETRIC-MANOMKTRIC METHODS

Muscle Extract. Remove the fascia as well as possible from muscles
of the hind legs and back of a cat. Homogenize thoroughly with an
equal weight of water and crushed ice, and centrifuge. A Waring
blendor and a Sharpies centrifuge are useful for these steps. With
stirring, add 4 vol. ice-cold acetone to the soln. in a slow stream.
Let the precipitate stand for 30 min. in the cold ; decant the super-
natant, and centrifuge the remainder. Wash the residue twice with
cold acetone in the centrifuge, transfer to a Biichner funnel anrl
wash with acetone and ether. Break up the cake into small pieces
and dry overnight in vacuo over sulfuric acid. Prepare a paste
of this acetone powder by grinding 600 mg. with successive 2 ml.
portions of water until 20 ml. has been added. Centrifuge; dialyze
the supernatant liquid in the cold against running distilled water
for 24 hr., and centrifuge. Remove interfering nucleotides by
adding 600 mg. of Norit charcoal to the dialyzed soln. Shake
mechanically in the cold for 1 hr., centrifuge, and treat the super-
natant fluid with another portion of charcoal for 1.5 hr. Centri-
fuge and filter. The DPN blank for this light brown soln. is small
at first and disappears completely after 12-24 hr. at 6°. The
acetone powder is very stable when kept in vacuo in the cold. The
activity of the aqueous extract remains constant 4-5 days when
kept cold.

PROCEDURE

1. Combine 0.4 ml. bicarbonate soln. with 0.6 ml. hexose diphos-
phate and 0.3 ml. arsenate. To 1 vol. of this mixture add 0.17 vol.
DPN soln., either the standard or unknown soln.

2. Pipette 1.60 //.l. resulting soln. into the diver.

3. Make up to a total vol. of 3.14 fx\. in the diver with water and
muscle extract (1.0 ml. muscle extract equivalent to 30 mg. acetone
powder). About 1 vol. muscle extract, diluted with 0.5 vol. distilled
water will probably be required.

4. Flush the diver with the gas mixture containing 5% carbon
dioxide and 95% nitrogen for 3 min. to bring the pH to 7.4.

5. Seal the neck of the diver with paraffin oil saturated with the
gas mixture and transfer to the flotation tube.

OPTICAL— LEVER RESPIRO.M ETR V

399

6. Take readings at 5 min. intervals after allowing 5 min. for the
initial eqnilibration.

2. Optical-Lever Respironielry

Heatle}', Berenblnm, and Chain (1939) developed an apparatus
in which a respiration chamber of 40-80 fA. is ground into a glass
plate and covered with a mica membrane to which mirrors are
attached. A change of gas pressure within the chamber causes the

I

I

miJLp.

□
□

0
0

Fig. 137. Component parts of respiration chamber: A, B, C, cups witli
cavities for two, three, and four separate droplets, respectively; D, mica mem-
brane with mirrors attached; E. "plate"; F. complete assembly ready for plac-
ing in brass case. From Hentlcy (1940)

mica to bulge and a compensating external pressure can be applied
to restore the membrane to its original position as indicated by an
optical lever. From the volume of the gas space and the change in
pressure required to keep it constant, the change in the gas volume
may be calculated. The instrument is sufficiently sensitive to measure
changes in the gas volume of 1 lA. per hour (about 200 times as
sensitive as the usual Barcroft or Warburg apparatus).

400

GASOMETRIC-MANOMETRIC METHODS

Heatley (1940) described an improved model in which up to six
respiration chambers may be mounted on a revolving frame to

A-

Fig. 138. Respiratory chamber.
From Berenblum, Chain, and Heatley (1940)

SECTION AB

FRONT ELEVATION

MEDIAN
VERTICAL SECTION

r Rubber washer of
" F^ 7 5 uneven thickness

REAR ELEVATION

8 B A countersunk

screw, holding

B toA

Rubber
washer

Rubber washer of T_
Circular glass window uneven thickness

MEDIAN HORIZONTAL 4BA clearance slot
SECTION

1 0 1

(Dimpn<;ions m mm )

Fig. 139. Details of metal case for respiration chamber.
From Heatley (1940)

ROTATING
BACK

facilitate manipulation and measurement. Before the war the appa-
ratus was made by Unicani Instruments Ltd.; but at the date of

OPTICAL-LEVER RESPIROMETKY

401

this writing production of the instrument has not yet been resumed.
The apparatus is rather complicated, mechanically, and since there
are other more available instruments which have simpler construc-
tion and greater sensitivity the use of the optical-lever manometer
will probably be limited. Therefore, only a cursory description of
the apparatus will be given.

Fig. 140. Details of revolving frame.
From Heatley (1940)

The respiration chamber (Fig. 137) consists of a "cup" which is
a 25 mm. square of glass, 3-4 mm. thick, in which cavities are
ground and then coated with a paraffin film. Drops placed in these

402

(iASOMETRIC-MANOMETRIC METHODS

cavities will not How together, but may be mixed by means of a
magnetic "ilea." The "cups" are covered with glass "plates" con-
taining three holes, as shown in Figure 137. The large hole in the
"plate" is covered with a 12 mm. square mica membrane not over
18 fx thick. Two 1 mm. squares of cover slip glass are cemented to
the mica with Seccotine, and mirrors, 2X3 mm., are cemented to
the cover slij) squares. The mirrors are made by silvering or alumi-
nizing cover slips and then cutting to size. The "plates," "cups,"
and mica membranes are sealed together with lubricant (No.
591822/39210— ^nYfs/i Drug Houses Ltd.). Another form of cham-
ber (Fig. 138) was used by Berenblum, Chain, and Heatley (1940)
for tiie determination of the respiratory quotient of tissue.

Fig. 141. Schematic diagram of optical system.
The image of the spht (S) is thrown upon the
ground-glass screen (A'') by the mirror system {R',
W, R, Q. P).li the mirrors W are parallel, a single
image will result ; if the mica bulges, the mirrors W
will tilt in opposite directions and the image will
divide in two. Dial Z controls the position of one
half of the divided mirror Q. From Heatley (1940)

The chamber is mounted in a metal case (Fig. 139) and the cases
are fitted into a revolving frame (Fig. 140). The chambers may be
opened or closed by rotating the "plate" over the "cup" through
60°. This is accomplished by turning disc C of the metal case (Fig.
139). A diagram of the optical system is shown in Figure 141, and
the principle of the pressure regulator is illustrated in Figure 142.
Finally, a view of the complete apparatus is given in Figure 143,
in which the following designations are used:

.1. thciiuoharunieter; B. supporting bracket for revolving frame; C, safety
tube to legulate maximum gas pressure; D, thermoregulator ; E, manometer;

OPTICAL-LEVER RESPIROMETRY

4();i

Fig. 142. Principle of manometer

and pressure regulator.

From Hentley, Berenhlum,

and Chain (1939)

Fig. 143. Diagrammatic view of complete apparatus.
From Heatlev (1940)

404 GASOMETRIC-POLAROGRAPHIC METHODS

F, one of the reservoirs of the pressure-regulating system; G, crank-actuating
reservoirs of pressure-regulating system; //, optical box; /, control panel; J,
guide block for accurately placing frame bracket; K, stand for frame bracket
when not in use; L, brass tube projecting from underside of optical box; M,
black felt hood protecting ground-glass screen from stray light; N, dial operat-
ing divided mirror; 0, pulley over which passes cord for rotating frame; P,
revolving frame for brass cases; Q, ratchet arm for rotating frame; R, cord
attached to latter; S, support for frame bracket during experiment.

The constants of the respiration chambers are determined as for
"Warburg vessels, mercury being used to measure the total volumes
of the chambers.

C. POLAROGRAPHIC

Polarographic methods have been employed for the determina-
tion of certain elements in small amounts of tissue, e.g., the pro-
cedures of Carruthers for sodium (1943a), magnesium (1943b),
and copper (1945), which were used for studies of carcinogenic
changes in mouse epidermis. These procedures require several
hundred milligrams of tissue for analysis and hence are suited to
histochemical work only when a relatively large quantity of histo-
logically well-defined material is available. Because of the limited
application these methods will not be given here.

On the other hand, polarography has been applied more readily to
respiration studies on the histochemical level. The mercury from the
dropping electrode may affect biological systems, but this possible
difficulty has been eliminated by Laitinen and Kolthoff (1941a,b),
who developed a method for the estimation of oxygen in solution
using a platinum wire electrode in place of the mercury type. Con-
tact between mercury and the biological material is also avoided in
the double vessel of Selzer and Baumberger (1942). The determina-
tion of the oxygen content of body fluids by means of the polaro-
graph was described by Beecher et al. (1942), but the method
of Davies and Brink (1942) would appear "to offer the best oppor-
tunity for the application of the polarographic technique to
respirometry on a scale suitable for histochemical work. The ap-
paratus of the latter investigators will be discussed in detail.

Microelectrode Measurement of Local
Oxygen Tension in Tissue

Davies and Brink (1942) described two types of stationary
platinum microelectrodes by means of which local oxygen tensions in

liOCAL OXYGEN TENSION

405

animal tissues can be determined with a spatial resolution of 25 /x.
In one electrode the end of a platinum wire is recessed inside a
cylindrical glass tip; this instrument may be used to measure abso-
lute oxygen tensions as often as once every 5 min. In the other elec-
trode the end of the wire is directly exposed to the outer medium;
with this electrode relative oxygen tensions may be recorded con-
tinuously. An idea of the applicability of the method may be gained
from the work of Davies and Brink, who employed the electrodes for
the measurement of the oxygen tension at the surface of superficial
arterioles and venules of the cat cerebral cortex, in the cortical
substance, at the surface of muscle cells, and at chosen distances
from the surface of unicellular organisms.

The principle of the method is based on the reduction at a plati-
num electrode of the dissolved oxygen according to the following
equation:

2H+ + O2 + 2e -> H2O2

and the measurement of the currents developed when suitable
potentials are applied, the currents being proportional to the oxygen
tension.

The variation in electrode current with applied potentials is
illustrated in Figure 144. The currents in the plateau region of the

Fig. 144. Current-voltage curve for
recessed electrode no. 11 in air-satu-
rated 0.15 M NaCl. Temperature,
37°C. Recess length, 1.0 mm. Bore,
0.176 mm. Each point is the value of
current 20 sec. after closing the cir-
cuit. Re-equilibration time, 20 min.
between measurements. Fro?n Davies
and Brink (19^2)

0 -02 -0.4 -06 -08 -1.0
POTENTIAL iVOLTS: us. 0 15 M CALOMEL

curve are limited by the maximum rate at which oxygen can diffuse
to the cathode, and this rate is proportional to the oxygen tension.
Also, the effect of change in the applied potential is a minimum in
the plateau region, and, accordingly, it is in this region that the
potentials are chosen for the measurement. The increase in the

406

GASOMETRIC-POLAROGRAPHIC METHODS

furrent above 0.8 volt is due to a second reaction, i.e., reduction of
ionic to molecular hydrogen.

Calihration and Measuring Instruments. The set-up for cali-
bration is illustrated in Figure 145. The platinum electrode {A)

Fig. 145. Diagram of apparatus used for calibrating

a stationary platinum electrode.

Frov^ Davics and Brink (1942)

dips into a 0.15 ilf sodium chloride solution and mixtures of oxygen
and nitrogen may be bubbled through the solution as indicated. The
calomel half cell is filled with the same solution and the potential
is supplied by a voltage divider. The potential is measured on the
voltmeter (T) and the current on the galvanometer (G).

When the electrode is used on tissue the surrounding tissue fluid
serves as the indifferent electrolyte and the calomel half cell is
placed as near the electrode as possible, touching either the exposed
tissue or a salt pad on another area of the organism.

With an air-saturated solution and electrodes made with wire of
0.2 mm. diameter or larger, currents of the order of 1 X 10"^ amp.
or greater are obtained. A galvanometer with a sensitivity of 5 X
10"^" amp. per mm. may be used. With electrodes of smaller di-
ameter, e.g., 25 /i., the currents are of the order of 1 X 10^^" amp.
for air-saturated solution and may be measured using a direct -
coupled amplifier. In the latter instance the galvanometer is replaced
by a well-shielded resistance whose value (R) is chosen to effect a
l)otential drop of about 1 millivolt applied to the input of the am-
I)lifier. The current (i) is given by the expression:

LOCAL OXYGEN TENSION 407

i = AE/yR

where y represents the voltage gain of the amplifier and aE the
change in output voltage resulting from the change iR in input volt-
age. The grid current of the input tube of the amplifier also passes
through resistance R but does not affect the measurement appreci-
ably. The very small currents make it imperative to avoid leaks to
the ground, and for this reason the switch and current-measuring
instrument are put on the platinum electrode side of the circut. A
coaxial cable with polystyrene bead spacers is used as the shielded
lead from the electrode and the end of the glass electrode shank is
coated with Petrowax (Gulf Oil Co.). In this fashion leaks can be
held down to the negligible magnitude of 1 X lO"^-'' amp.

Recessed Electrodes. The recessed electrodes are prepared by
sealing a platinum wire in a soft-glass tube of comparable inside
diameter in such a fashion that the tube extends beyond the end of
the wire to form a recess of the chosen length (recesses of 0.6-1.6
mm. have been used by Davies and Brink). To avoid gas bubbles in
the seal it is essential to degas the platinum before the sealing by
flaming it to white heat. The recess should be a uniform cylinder
whose axis coincides with that of the wire. The completed electrode
is annealed at 425° to prevent formation of cracks, which would
cause electrical leakages. In the preparation of small electrodes,
e.g., with a recess of 0.6 mm. length and 25 fi inside diameter, greater
control in the sealing requires the use of electrically heated platinum
loops. One loop, 0.3 mm. diameter, is used for sealing the wire into
a small glass tube, while another loop, 4 mm. diameter, is used to
seal the small tube unit into a larger glass shank. A low-power
microscope aids in the observation of the sealing and allows for
greater control of the process. The current-potential curves for
these small electrodes have more poorly defined plateaus than those
obtained with larger electrodes.

In testing, the electrode is placed in the circuit shown in Figure
145, and sufficient time is allowed for the oxygen in the recess to
attain equilibrium with that dissolved in the solution. After setting
the potential the switch is closed, and the current falls as the concen-
tration gradient spreads into the solution. After a fixed number of
sec. from the time the switch was closed the current reading is taken.
Then the switch is opened and sufficient time is allowed for the

408

GASOMETRIC-POLAROGRAPHIC METHODS

restoration of equilibrium before another measurement is made. Tlie
operation is repeated at other potentials to obtain a current-voltage
curve (Fig. 144).

The time the circuit must be left open between readings must be
determined for each electrode. This is accomplished by plotting cur-
rent readings against time between successive closings of the circuit.
For each measurement the same number of sec. must be allowed to
elapse between the circuit closing and the reading. From Figure 146

CE 2

■=>

" 1

Recess length 1.6 mm.

pP°—

Recess length 0.6 mm.

10 20 30

TIME (MIN.) BETWEEN READINGS

40

Fig. 146. Twenty-second current
as a function of time between readings
for two recess lengths. Bore, 25 fi. Air-
saturated 0.15 M NaCl. Temperature,
37°C. Potential, -0.60 v. vs. 0.15 M
calomel. From Davies and Brink
(1942)

it is apparent that the readings can be taken every 20 min. with the
electrode having the 1.6 mm. recess, and every 10 min. with the
0.6 mm. electrode.

The current, as a function of the time elapsing between closing
the switch and taking the reading, can be predicted on the basis
of diffusion theory. If the electrode recess is a true cylinder and if
readings are taken before the concentration gradient has extended
beyond the orifice, the diffusion will be one dimensional and the
following relationships will hold:

.. - "fJ

'xi2y/m

Cr.« —

exp (-?/2) dy

it = nYyCAiD/irty/^

where Cx.i = concentration of oxygen (moles/ml.) at distance x
cm. from the platinum surface at t sec. after the start of diffusion
(closing the electrode switch) , C = initial uniform concentration of
oxygen, y = a, variable of integration, D = diffusion coefficient of
oxygen (cm.Vsec), it = electrode current (amp.) at time, t, n =z

LOCAL OXYGEN TENSION

409

no. of electrons used per molecule of oxyge nelectrolyzed, Fy = 1
faraday (96,500 coulombs), A = area of a platinum surface (cm.^).
From the preceding relationship the current is inversely propor-
tional to the square root of the time (t). When the time is increased
to the degree that the gradient extends into the solution outside the
recess, the oxygen diffuses to the tip of the electrode from all direc-
tions instead of from only one, and this results in a concentration
at the orifice higher than that for the linear diffusion. Thus the
current is increased beyond that expected from the l\/[ relation.
This is illustrated in Figure 147, which shows the linear relationship

Fig. 147. Current-time curve for
electrode no. 11. Air-saturated 0.15 M
NaCl. Temperature, 25 °C. Potential,
—0.60 V. vs. 0.15 M calomel. From
Davies and Brink (1942)

14

12 -

E 8
S' 6

UJ

cc

§ 4
o

2 -

■r — 1 r !

—I 173 — 1

/

/

/

-

y/

-

- J^

-

—*-'/

/

/

-

/

/

'. 1 1 1 1

1 1 r

01 0.2 0.3

TIME, l/vT

0.4

up to 40 sec. Although the profile of the current-time curve is in-
fluenced by the shape of the recess in the electrode, the current at a
given time after closing the circuit is still proportional to the oxygen
tension as long as there is no change in T>. Thus, if the concentration
gradient is confined to the recess when the current is measured, only
the length of the recess will be important and it will not matter
whether the cross-sectional area of the recess is uniform.

The distance from the platinum surface to which the concentra-
tion gradient should extend at times after the onset of electrolysis
is shown in Figure 148. Up to 20 sec. the gradient extends to 1 mm.
from the platinum surface.

In Figure 149, the electrode current at intervals after the start of
diffusion is shown as a function of the initial uniform oxygen tension
of the solution in recess after equilibration with known gas mix-
tures. While the electrodes are calibrated at the temperature of the

410

GASOMETRIC-POLAROGRAPHIC METHODS

tissue to be studied, the calibration curves in Figure 149 arc actually
unchanged within the experimental error between 28° and 37°. The
negligible effect of the temperature change results from counter-
balancing the associated changes in D and the solubility of oxygen.
However, the temperature must be known in order to convert
partial pressures into concentrations of oxygen.

0.2 04 06 0.8 1.0 1.2 1.4 1.6 1,8
DISTANCE FROM PLATINUM SURFACE, mm.

Fig. 148. Concentration-distance
curve at various times after beginning
linear diffusion. Assumed value of D,
4.0 X 10~^ cm.^/sec. From Davies and
Brink (194£)

20 40 50 80 100 120 140 160
OXYGEN TENSION, mm Hg

Fig. 149. Current vs. oxygen ten-
sion from current-time curves. Elec-
trode No. 11. Same solution, tempera-
ture, 28 °C. Potential, —0.60 v. vs.
0.15 M calomel. From Davies and
Brink (1942)

Control experiments have revealed no evidence that substances
other than oxygen are electrolyzed under the conditions employed.
The recessed electrode offers the distinct advantage that the meas-
urements may be made independent of the diffusion coefficient in
the medium beyond the orifice. To maintain constancy of the diffu-
sion coefficient within larger recesses, they are filled with agar gel
containing 0.15 M sodium chloride. The smaller recesses exclude
particulate matter by virtue of the narrow bore; the diffusion of
solutes into the recess has no appreciable effect on the value of D
Another advantage of the recessed electrode is its freedom from
the effects of convection in the external solution.

LOCAL OXYGEN TENSION 411

When used in blood, the electrode should have its recess filled with
agar containing salt to keep the red cells out of it, as these would
elicit abnormally high currents by virtue of their function as oxygen
sources.

The recessed electrode has been found to give calibration constants
which are reproducible to ±3% over a period of weeks. Continued
exposure to tissue fluids for some hours will result in a drift in the
calibration to yield oxygen tension values which are too low. How-
ever, an equal exposure to 0.15 M sodium chloride will bring the
values back to the correct level. Davies and Brink suggest that this
difficulty may be obviated by calibration in the tissue fluids. Later
they found that by filling the recess with distilled water and then
covering^ the tip of the electrode with a collodion membrane, the
drift in calibration during exposure to tissue fluids can be made
very small.

Open Electrodes. The open type of microelectrode is prepared
by fusing a platinum wire, 25 /j, diameter, into a soft-glass tube so
that one end of the wire is flush with the sealed end of the tube.
While the open electrode cannot be used for the measurement of
absolute oxygen tension, it can serve to measure rapid changes in
tension. Thus, Davies and Brink were able to record, to about 0.1
sec, the sudden oxygen consumption occurring when a muscle fiber
contracts.

The variable properties of the open electrode make for consider-
able instability in its performance. However, polarization for several
niin. at 1.0-1.2 volts effects some stabilization; even so the repro-
ducibility is only about 15% under favorable circumstances. Cali-
bration before and after each experiment will hold the error to a
minimum. Poorly defined plateaus are obtained in the current-poten-
tial curves, but as a rule with up to 0.8 volt nothing but oxygen is
electrolyzed in oxygen-free solution. A linear relation may be found
between current and oxygen tension in calibration experiments;
however, when applied to tissue, particularly near blood vessels,
there may be little correlation between the actual tension and that
derived from a calibration curve which is obtained with a solution
such as Ringer's.

F. DILATOMETRIC TECHNIQUES

As Sreenivasaya and Bhagvat (1937) pointed out in their review
of the subject, dilatometry has seen comparatively little application
in the study of chemical and physical changes, although the recogni-
tion of its possibilities is by no means new. The adaptation of
dilatometry to fine quantitative measurements, with particular
reference to histo- and cytochemistry, was made by Linderstr0m-
Lang (1937a). The feature of this adaptation is a density gradient
in a nonaqueous medium in which a very small drop of aqueous
reaction mixture is suspended. Changes in volume of the drop that
accompany the chemical changes taking place within it are made
manifest by a vertical displacement to a new position where the
specific gravities of the drop and the surrounding medium are again
equal. The magnitude of the displacement then becomes a measure
of the extent of the reaction that occurred within the drop. Stand-
ardization of the density gradient is accomplished by introducing
aqueous drops of known densities.

The method is obviously limited to those systems that do not
contain or evolve constituents soluble in the bromobenzene-kerosene
medium. Thus lipase measurements could not be made in this
manner. The sensitivity of the method largely depends on the magni-
tude of the contraction constant, which may be defined as the
volume change occurring when one gram molecule of reactant under-
goes chemical change. Expressed mathematically,

K = vM/cV

where K is the contraction constant, M the molecular weight, v the
change in volume, V the original volume, and c the concentration.
The constant for the hydrolysis of urea is 24.1; hence, when 60 g.
(1 mole) urea is hydrolyzed, the reaction mixture decreases in vol-
ume by 24.1 ml. It is important that the temperature be held very
constant during the measurement so that the change determined
is only the isothermal one. The advantage of this method lies, not
only in the circumstance that very small reaction volumes may be

418

414

DILATOMETRIC METHODS

employed, l)ut also in the fortunate fact that the measurements
may bo made without disturbing the system in any way.

DILATOMETRIC APPARATUS AND ITS USE

Gradient Tube. Tlu> glass gradient tube, that is employed to
furnish a ])ractically linear specific gravity gradient, has the form
and dimensions shown in Figure 150. The tube is mounted in a
thermostat that maintains its temperature constant to about
±0.002"; the group at the Carlsberg Laboratory have done their
work at 30°. The tube is first filled with bromobenzene (sp. gr. 1.48)

-T-

E

u
O

Surface of

water m

thermostat

6 cm

-»-2.5cm.

-»4-7 cm.-+*

Fig. 150. The gradient tube.

From Linderst r0m-Lan g

and Lanz (1938)

Fig. 151. Measuring micioscope.

Frt)))t Linderst r()m-La7ig

and Lanz (1938)

up to the middle of the part connecting the two bulbs. After the
bromobenzene has attained the temperature of the bath, "water-
white" kerosene (sp. gr. 0.79) having the same temperature is added
carefully through a funnel fitted with filter paper to fill the remain-
der of the tube. The surface of the liquid in the tube should be below
that of the thermostat water. A long spatula is used to agitate gently,
for about 10 sec, the liquids in the region of their junction in the
middle of the tube so that some mixing will occur. After 24-48 hr.
the density gradient is sufficiently linear. Usually the gradient is
maintained for months and even years.

APPARATUS

415

Saturation of Medium with Water. It is necessary to saturate
the medium with watpr at a suitable vapor pressure in order to
minimize the tendency of the aqueous droji to lose water to the
medium and thus change its specific gravity. It has been found
sufficient to employ a 0.2 .1/ potassium bromide solution for the
i-ange of specific gravity from 0.99-1.01. About 1 ml. of the salt
solution is shaken thoroughly with about 10 ml. of the less dense
kerosene-bromobenzene mixture. The resulting suspension is poured
at once into the gradient tube, and as the drops fall they saturate
the medium. The high specific gravity of the drops carries them down
far enough in the tube to avoid any interference with subsequent
measurements.

Gradient Calibration. Calibration of the gradient is accom-
plished by the simple expedient of placing in the tube small drops
(0.10-0.15 fA.) of potassium chloride solutions of known specific
gravities (Table VIII). It is well to store these standard solutions
under a 1 cm. layer of kerosene in stoppered vessels, such as 50 ml.
volumetric flasks, which restrict the surface exposed to the kerosene.
In some cases, such as in the determination of "reduced weight",
( page 420) , standard drops composed of mixtures of doubly distilled
water and deuterium oxide have been used. The^O.10-0.15 fA. pipette
used to add the drops may be either the type shown in Figure 51

TABLE VIII
Composition and Density" of Standards for Dilatometry

Standard

No.

Potassium
chloride, %

d\°

Standard
No.

Potassium
chloride, %

d^o

0

0

0.995673

6

1.0272

1.002155

1

0.1719

0.996785

7

1 . 1848

1.003149

2

0.3408

0.997823

8

1.3442

1.004155

3

0.5122

0.998905

9

1.5121

1.005214

4

0.6633

0.999857 1

10

1 . 6904

1.006339

5

0.8450

1.001005

11

1.8721

1.007486

" The densities are taken from the Landolt-Bornstein tables.

(page 173) or the constriction pipette (Fig. 53, page 173). These
pipettes may be calibrated by measuring out strong acid of known
concentration and titrating it with a microburette, but for the
present purpose it is sufficient to calibrate them more roughly by
weighing the quantity of water they deliver directly on the pan of a
microbalance.

416 DILATOMETRIC METHODS

Method of Adding Drops. The procedure for adding the drops
is as follows:

1. Dip the tip of the pipette below the kerosene layer into the
solution.

2. Rinse the pipette by drawing up and blowing out the solution
several times. »

3. Draw the solution to the mark on the pipette.

4. Raise the pipette until the tip is in the kerosene.

5. Draw up about 0.1 /xl. kerosene into the pipette.

6. Remove the pipette and wipe the outside with the edge of a
piece of filter paper.

7. Blow out about half the kerosene while the tip is touching a
piece of filter paper.

8. Dip the pipette into the gradient tube so that the tip is about
2 mm. below the surface of the medium.

9. Blow out the pipette charge.

10. Remove the drop from the tip by lifting the latter out of the
medium. The drop falls and reaches its equilibrium position usually
within 15 min.

Method of Removing Drops. Drops may be removed in the
gradient tube by inserting a long thin glass rod with a fine point to
which the drops will adhere. They may then be carried up and
placed on the wall of the tube near the surface of the medium. It is
impossible to remove them entirely from the medium in this fashion
since the surface forces act to detach the drops from the glass tip
when this is attempted. The glass rod should be handled carefully
to avoid undue disturbance of the density gradient.

Dr. Oliver H. Lowry instituted the practice of adding a little sand
(60-80 mesh) from a salt shaker to remove the droplets. This is an
effective method which disturbs the gradient less than the glass rod
procedure.

Measurement of Drop Position. The position of drops is
determined by a horizontal microscope of long focal length mounted
on a micrometer stand (Fig. 151). The microscope (A) is pivoted
at jB; C is a calibrated rough adjustment rack with a range of 100-
150 mm., and £' is a clamp to hold the rack at any given setting.
The micrometer screw D enables fine adjustment, it has a vertical

PEPTIDASE 417

•

range of 10-12 mm. with one complete revolution giving a movement
of 1 mm. Since the circumference is divided into 100 parts, measure-
ments may be made with an accm'acy of 0.01-0.02 mm. The appara-
tus is mounted on a triangular base fitted with leveling screws.

The position of a drop with reference to a standard drop is deter-
mined by turning the micrometer screw to zero, setting the reference
mark in the microscope ocular (cross hair or scale division) at the
lower edge of the standard drop by means of rack C and clamp E,
moving the microscope horizontally so that it will be in line with the
drop to be observed, and bringing the reference mark to the lower
edge of the latter drop by adjustment of the micrometer screw.

Measurement of Reaction Rate. When the gradient tube con-
tains a series of standard drops embracing the range of specific
gravity within which the reaction mixture falls, measurements of
reaction rate may be carried out by determination of positions at
suitable intervals. Readings are begun after 15-20 min. from the time
the drop of reaction mixture is introduced into the tube. It has been
shown that this is an adequate time to allow the drop to assume its
proper position for observation. A plot of the densities of the drop as
a function of time represents the course of the reaction.

PEPTIDASE

The use of the gradient tube for dilatometric determination of
peptidase activity was described by Linderstr0m-Lang and Lanz
(1938). They employed DL-alanylglycine as substrate, of which only
the D form is hydrolyzed enzymatically by pig stomach and intestinal
extracts, which were used as the source of the enzyme. The change
of density with time was found to be a linear function. The contrac-
tion constant for the scission of the peptide bond was found to vary
with both pH and the concentration of the phosphate buffer em-
ployed. The magnitude of this variation, as determined by both
direct macro dilatometry and the micro method, is shown in
Table IX. These measurements were made at 30° on reaction mix-
tures consisting of equal volumes of substrate and enzyme solutions.
The substrate solutions had the composition given in Table X; the
pH values were calculated on the assumption that the pK value for
DL-alanylglycine at 30° is 8.09. The enzyme solutions were prepared

418

DILATOMETRIC METHODS

by grinding 30 g. pig intestine with sand and 100 ml. 60% glycerol;
after filtering, 1:50 dilutions were made with phosphate buffers
having the different pH values and concentrations indicated in the
table. Controls were run in which the enzyme solution had been
heated at 100° for 30 min. in a sealed glass tube.

TABLE IX

Contraction Constants for Alanjdglycine Hydrolj^sis

by Peptidase from Pig Intestine at Different

pH Values and Phosphate Concentrations"

Phosphate
concn. in

pH6.8

pH7.1

Micro
method

pH7.4

pH7.7

pHS.O

reaction
mixt.

Micro
method

Direct
dilat.

Micro
method

Direct
dilat.

Micro
method

Micro
method

Direct
dilat.

ilf/60

M/120

M/240

9.l6

9.12

9.18

9.85

9.47

9.28
9.2,

9.44
9.I2

9.99

11.42
11.22

9.4,

" fi\./mM peptide bond.

TABLE X

Composition of Substrate for Peptidase Measurements
at Various pH Values

Molarity of Molarity of

pH DL-alanylglycine NaOH

6.81 0.2 0.0100

7.13 0.2 0.0193

7.42 0.2 0.0346

7.72 0.2 0.0611

8.00 0.2 0.0900

The deviations in the constants as determined by both methods at
pH 7.7 and 8.0 are not understood. Accordingly, Linderstr0m-Lang
and Lanz have chosen to limit the drop method, for the time being,
to reaction mixtures with pH values in the range 6.8 to 7.4.

Enzyme experiments lasting 24 hours may be safely carried out.
The accuracy of the method is about fifty times that attained by the
microtitration procedure ( page 302 ) . An idea of the quantity of
material that may be subjected to investigation by the density gra-
dient method is given by calculations included in the paper of Lin-
derstr0m-Lang and Lanz. They estimated that if 10 hours be taken

PEPTIDASE 419

as a maximal time for measm-ements, and the density determined to
1 X 10"■^ with 0.1 fA. drops, the least quantity of alanylglycine split-
ting that could be detected in 10 hours would be 1 X 10""^ millimole.
This splitting could be accomplished by one thousandth of one sea
urchin egg, corresponding to about 5 X 10"^ mg. dry organic mate-
rial.

Jacobsen (1942) showed that the contractions accompanying the
cleavages of peptide linkages in benzoyl-DL-argininamide by trypsin
and benzoyl-L-tyrosylglycinamide by chymotrypsin are 13.4 ± 0.2
and 15.6 ± 0.8 ml. per mole, respectively.

Method of Linderstr^ni-Lang and Lanz for Peptidase

SPECIAL REAGENTS

Enzyme Preparation. 60% glycerol extract of tissue diluted with

phosphate buffer, pH 6.8 to 7.4, or a bit of cellular material can be

used directly.
Substrate Solution. 0.2 M OL-alanylglycine containing sodium

hydroxide of the molarity indicated in Table X to give the desired

pH.
Mixture for Saturation of Medium. Combine equal vol. of 0.2 M

potassium bromide and substrate soln.

PROCEDURE

1. Saturate the medium in the gradient tube with the potassium
bromide-substrate soln. mixture in the manner described in the
preceding section. After 24 hr. the tube may be used.

2. Introduce a number of 0.15 fA. standard drops into the tube.
The sp. gr. of these drops should bracket the range encountered in the
determination. Two drops of the same standard should not deviate
from one another in their equilibrium levels by more than 0.1 mm.

3. Mix equal vol. enzyme and substrate soln. under kerosene in
a small test tube, and place 2-3 drops 0.15 ix\. each in the tube. For
microtome sections of tissue, or smaller cellular units, first place a
1 ix\ kerosene drop on the end of a glass needle about 0.2 mm.
thick; insert the tissue into this drop with the aid of a very fine glass
needle, dip the drop containing the tissue into the gradient tul)e

420 DILATOMETRIC METHODS

medium, and slowly raise the tip of the needle out of the liquid to
disengage the drop.

4. Repeat preceding step with control drop containing heat-
inactivated enzyme.

5. Record positions of drops, relative to standard drops, at inter-
vals of about 15 min., until the rate of change in position of both
active and control drops is equal, indicating that the enzymatic
process is complete.

6. Plot results to determine the slope relating density change
with time. When the size of the drops and experimental conditions
are kept constant, the millimoles of alanylglycine split per unit time,
R, is given by the equation:

R = {Sv/Kd)

where S represents the slope, y the drop vol., K the contraction
constant under the experimental conditions, and d the drop density.

DENSITY AND "REDUCED WEIGHT"

Aside from its use to follow the course of reactions, the density
gradient technique can be employed as a very convenient method
for the measurement of the density of very small amounts of mate-
rial. Linderstr0m-Lang, Jacobsen, and Johansen (1938) applied the
technique to the measurement of the deuterium content in mixtures
of water and deuterium oxide. Jacobsen and Linderstr0m-Lang
(1940) modified the apparatus for more rapid determinations of
specific gravity by the use of a 200 ml. graduated cylinder as a
gradient tube, omitting thermostatic control and the use of the
traveling microscope. This simplification is well adapted to the
measurement of the specific gravity of biological fluids with an
accuracy of 0.1%. Lowry and Hunter (1945) employed a similar
apparatus for the determination of serum protein concentration.

It is of particular interest that the gradient tube has been adapted
to the determination of "reduced weight" ( Linderstr0m-Lang and
Holter, 1940), since the latter value affords a measure of the size of a
tissue sample independent of its water content. In some respects this
circumvents one of the chief difficulties encountered in histochemical
and cytochemical investigations, i.e., obtaining a quantitative defini-
tion of structural elements in tissues or cells to which quantities of a

DENSITY' AND REDUCED WEIGHT 421

constituent measured can be referred. The "reduced weight" is the
weight of the sample minus the weight of an equal volume of water,
or expressed mathematically:

gr = g — (vg dj

where Qr is the "reduced weight," g the weight of the sample, Vg the
volume of the sample, and dy, the density of water at the experimental
temperature. When the gradient tube is employed to determine the
"reduced weight," it is only necessary to place the sample in a small
drop of water, measure the specific gravity of the unit as a whole,
obtain the diameter of the water drop with the aid of the micrometer
microscope and from this value calculate the volume, and finally
apply the equation :

Qr = (d — dj V

where d is the specific gravity of the drop containing the sample,
and V is the volume of this drop. For standardization of the gradient,
drops are used consisting of mixtures of double-distilled water and
deuterium oxide ; and they have density differences of about 1 X 10~^.

The accuracy of the method depends on the size of the drop. For
0.1 lA. drops it is about 5 X 10"'^ mg. and for 1.0 lA. drops it is about
5 X 10-^ mg.

The tissue sample employed for "reduced weight" measurement
can be used subsequently for enzymatic determination, provided the
kerosene-bromobenzol medium has no influence on the activity.
The drop containing the tissue sample is removed from the gradient
tube by means of a retriever consisting of a thin glass rod the tip of
which is fused to a piece of cover glass, 2X2 mm., at an angle a
little greater than 90°. Once removed from the tube, the material can
be transferred to the medium appropriate for the enzyme measure-
ment.

VL DETERMINATION OF AMOUNT OF
A BIOLOGICAL SAMPLE

A major problem in histo- and cytochemistry is the quantitative
definition of the samples of biological material which are to be con-
sidered. Weight, volume, numbers of cells, and nitrogen or nucleic
acid content have been variously employed for this purpose, and
both advantages and disadvantages are to be found in each case. The
actual choice of a reference quantity will depend on the particular
problem to be considered.

Weight. Wet- and dry-weight measurements may be made with
commercial microbalances including torsion balances such as those
manufactured by Roller-Smith Co., which are sensitive down to
about 2 fjLg. Quartz fiber balances of greater sensitivity are described
on page 189.

It is advantageous in some instances to employ the so-called "re-
duced weight" which is the weight of the sample minus the weight
of an equal volume of water. The feature of this value is that it is
independent of the water content of the sample. "Reduced weight"
can be determined rather simply in the gradient tube apparatus by
placing the sample in a drop of water and measuring the density of
the drop. From the density, the volume of the drop, and the density
of water, the "reduced weight" can be calculated (page 421).

Volume. When the sample, such as an Amoeba profeus, is of a
nature and size that permits it to be drawn into a capillary tube of
known diameter, the volume can be calculated after the length of the
sample in the tube is measured. However, this procedure cannot be
used if the sample has an irregular shape which does not allow it to
completely fill the lumen of the capillary over the entire length of
the body or if it is so sticky that it adheres to the wall of the capil-
lary and thus interferes with the manipulation or becomes damaged.
To determine volumes in these cases, Holter (1945) developed a
colorimetric method which he applied to measurements of the amoe-
ba, Chaos chaos. The details of the procedure are given on page 432.

The volume of microtome sections of tissue is regulated by cut-

423

424 AMOUNT OF A BIOLOGICAL SAMPLE

\

ting circular sections of a known thickness and diameter as de-
scribed on page 427.

Numbers of Cells. When the cells are in suspension their
number may be determined by counting in a hemocytometer cham-
ber. It may aid in the counting to stain the cells first. However, the
measurement of the number of cells in histological preparations is
more involved. A technique for cell counting in mounted stained
sections of tissue was described by Linderstr0m-Lang, Holter, and
S0eborg Ohlsen (1934).

The data of Rask-Nielsen (1944) for the number of cells per unit
volume of pyloric mucosa may be taken to illustrate the counting
technique. A microscope magnification of 300 X was used, and the
ocular was equipped with an ocular micrometer containing a circle,
divided into quadrants, which enclosed an area corresponding to
0.0638 mm.^ of the tissue section. Six to ten random counts, uni-
formly distributed over the stained section, were made of all nuclei
and fragments thereof within the bounds of the counting area.

In a given case a mean of the chief cells from eight random counts
was 229. The mean error of the mean value of the counts was ±4.4,
obtained from the formula:

rj^ / q-a

where ( SA^) is the sum of the squares of the individual deviations
from the mean value, q the number of counts, a the counting area,
and A the area of the section.

In order to convert the number of nuclei plus fragments per count-
ing area (229 ± 4.4) to cells per counting area, a correction factor
for the fragments must be applied. This factor is 12/(12 -\- h)
where 12 is the thickness of the section and h the height of the
nuclei, both expressed in microns. In this case the factor equals
0.71; hence:

(229 ± 4.4) 0.71 = 163 =>= 3.1 cells/counting area

Since the sections were cut from a cylinder of fresh-frozen tissue having a
diameter of 2.64 mm.:

2 (2.64)2 = 547 mm.2, area of a fresh section

6.47 X 0.012 = 0.066 /xl, volume of a fresh section

5 47
and - '„-„ = 85.7 counting areas/fresh section

U.Ubdo

- CELL COUNTING 425

However, the fresh-frozen sections were fixed and stained before
the counting so that a correction is required to compensate for the
contraction produced by this treatment. Since the counting was car-
ried out through the entire thickness of the section, the contraction
in thickness need not be considered because the ratio of the actual
number of cells to the cells counted will not be altered by a change
in thickness. The change in area must be considered, however. The
ratio of the area of each stained section to that of the fresh section
(5.47 mm.-) was obtained by projecting both images at the same
magnification and measuring the respective areas with a planimeter.
The areas might also be obtained by tracing the outlines of the pro-
jected images on paper and weighing the paper. Then the actual
number of cells per section is

/No. stained cells \ ^ /no. counting areas \ ^ /area of stained sectionX
V per counting area/ \per fresh section/ \ area of fresh section /

And this quantity divided by the volume of a fresh section equals
the actual number of cells per unit volume of fresh tissue.
In the present case:

/5.08\

(163 ^ 3.1) (85.7) V5.47; ^ (igg ^ 3 7) ^q, eells^l.
0.066

In regions of inhomogeneity within the section the relative areas
of the mutually deviating regions were measured after projecting
the images on paper, and the separate regions were counted. Each of
the regions of a given cell type was made to contribute to the final
mean result as illustrated in the following examples from Rask-
Nielsen (1944):

(l) In the case of a section containing one type of cell not homogeneously
distributed:

In 61% of the section area the cells were arranged in glandular tubules. Mean
of five random counts was 306. Mean error ±6.3.

In 39% of the section area the cells were arranged in pits. Mean of four
random counts was 203. Mean error ±14.8. Mean number of nuclei plus frag-
ments per counting area was:

(306 ± 6.3) 0.61 + (203 ^ 14.8) 0.39 = (187 ± 3.8) -f- (79 ± 5.8) = 266 ± 9.6

Applying the correction (0.71) for nuclei fragments:

(266 ± 9.6) 0.71 = 189 ± 6.8 cells /counting area

and then following the formula used previously:

426 AMOUNT OF A BIOLOGICAL SAMPLE

/5.30\
\5A7) _

^'"'"'•^n^fP — - = (238 =. 8.6) 103 cells/Ml.
0.066

(2) In the case of two different types of cells in the same section:

Epithelial cells comprise 25% of the area of a section.

Neck chief cells comprise 75% of the area of the section.

Epithehal cell count = 262 ± 6.7.

Epithelial cells per counting area = (262 ± 6.7) 0.71 = 186 ± 4.8.

/5.00\

aS6 =^ 4.8(85^) {5 A7) ^ ^.25 = (55 =. 0.14) 10^ epithelial cells/^l
0.066

Neck chief cell count = 326 ± 8.5.

Neck chief cell per counting area = (326 ± 8.5) 0.71 = 232 ± 6.0.

/5.00\
(232 ± 6.0) (85.7) \5.47/ ^ q ^^ ^ ^206 ± 5.3) 10« neck chief cells//xl.

The determination of a constituent in each type of cell in this section may
follow the example given:

From the nearest section containing onl.v epithelial cells a peptidase activity
of 0.125 X 10'^ units per cell was found. Therefore, the activity arising from the
epithelial cells was:

(55 ± 0.14) 10=" X (0.125) 10"^ = 6.8 units/Ml.
and

(6.8) (0.066) = 0.45 unit/section.

The activity of the section composed of both the epithelial and neck chief
cells was found to be 1.70 units, of which 0.45 unit was ascribed to the epithe-
lial cells present. Therefore, the activity of the neck chief cells in the section
was considered 1.70 — 0.45 = 1.25 units, and the activity of these cells was
1.25/0.066 = 19 units/yul., or 19/206 X lO' = 0.092 X lO"" units/neck chief cell.

Nitrogen. The total nitrogen in a sample has often been used
as an indication of the quantity of protoplasm present. Measure-
ments of the nitrogen content may be carried out by one of the pro-
cedures given on pages 230-239, and 283.

Nucleic Acid. Berenblum, Chain, and Heatley (1939) employed
the estimation of nucleic acid phosphorus as an indication of the
quantity of cellular material present, in an attempt to refer meas-
urements to the amount of metabolizing substance rather than to

SECTIONING METHODS

427

the total tissue, which would include inactive material. Berenblum
et al. first removed lipid phosphorus by extracting with alcohol-
chloroform (3:1). Organic and inorganic acid-soluble phosphorus
was removed by extracting with 0.1 A^ hydrochloric acid; the tissue
was then ashed with perchloric acid, and the phosphorus in the ash
was measured.

A. PREPARATION OF FROZEN TISSUE SECTIONS
OF ACCURATE THICKNESS

In some instances it is undesirable to subject tissue to the em-
bedding process and the associated treatments with the various sol-
vents prior to the application of histochemical tests or analyses on
microtome sections. When this is the case, the freezing microtome
may be used for either fresh tissue or that fixed in a suitable man-
ner. It may be necessary to obtain sections of accurately uniform
volumes for quantitative work. The cross-sectional area of sections
can be controlled by punching out cylinders of tissue from frozen
material with metal borers of known internal diameters (Fig. 152).

Fig. 152. Tissue borer and microtome cryostat.
From Linderstr077i-Lnng and Holler (1940)

The cylinder of tissue can be placed on some wet filter paper set on
the freezing head of the microtome and then the whole firmly frozen
to the head. This is the procedure that has been followed in many
cases by the Carlsberg Laboratory investigators. They have found
too that a rotary microtome is less subject to factors which lead to
variations in thickness than the hand sliding type, and that the

428 AMOUNT OF A BIOLOGICAL SAMPLE

greatest uniformity is achieved by continuous sectioning at con-
stant speed. The first section is always discarded, and, in order to
minimize the temperature effect of the intermittent cooling with
carbon dioxide as commonly practiced, it is particularly important
to discard the first section after cooling and proceed at once to cut-
ting before appreciable warming can occur. These difficulties have
been surmounted to a great degree by Linderstr0m-Lang and Mo-
gensen (1938), who devised a means of maintaining the entire micro-
tome at a constant temperature low enough to keep the tissue frozen
on the block, at the same time making possible the free manipula-
tion of the instrument and the sections. In addition they developed
a method to prevent undue distortion or curling of sections on the
knife edge.

Linderstr0iii-Lang and Mogensen Method for Accurate Cutting
and Special Handling of Frozen Tissue Sections

A cryostat large enough to hold a rotary microtome is arranged to
maintain a constant temperature of around —20°. The type devel-
oped by Linderstr0m-Lang and Mogensen (1938) is indicated by
Fig. 152. The cabinet is made of two layers of wood, an inner one of
22 mm. furniture board and an outer one of 10 mm. cross cut veneer.
A sloping lid is hinged to the front of the cabinet, and through two
openings {A and B), lambskin-lined leather gloves are attached. The
gloves are of a size to permit easy manipulation within the cabinet.
An observation window (C) consists of a fiat glass cell filled with
water, which is prevented from freezing by a small low-heat elec-
tric resistance coil fitted against it. An extra hole (D), which is kept
closed with a stopper, enables removal of sections from the cabinet.
The interior is illuminated by a lamp (E) which fits over a glass
window in the top. A wooden partition separates the interior into
a front and rear chamber, the latter being about half the size of the
former. The rear compartment, fitted with a well-insulated lid, is
arranged to hold dry ice. Two guide rails fixed on the fioor of the
front compartment serve to hold the microtome in position, and in
order to prevent the frosting of the microtome, an electric heater
capable of holding the microtome at 5° in placed between the rails.
A thermoregulator fits through the left side of the roof of the micro-

SECTIONING METHODS

429

tome chamber, and in the right side of this compartment an electric
blower, such as a hair dryer, is so placed that it blows air from the
compartment against a movable metal valve that permits the air
either to go back directly into the microtome chamber or to pass
through the Dry-Ice box before returning. The valve is controlled
by the thermoregulator.

Fig. 153. Device for prevention

of curling of sections.

From Linderstr0m-Lang

and Mogensen (193S)

Fig. 154. Use of device for prevention of section curling.
From Linderstr0m-Lang (1939)

430 AMOUNT OF A BIOLOGICAL SAMPLE

In practice, the inicrotome without its tissue freezing block is
placed in the cabinet, the Dry-Ice chamber is filled with small
pieces, the blower is started, and after about 45 min. the tempera-
ture is at —20 ±0.5°. A cylinder of tissue is frozen to the block
using carbon dioxide, and the block is then brought inside the
cabinet through a removable window (C). After the block is
fastened to the microtome, the knife is set and cutting is begun.
When not in use, the microtome is stored in a large desiccator with
sulfuric acid.

The sections are prevented from curling on the knife edge by the
arrangement shown in Figures 153 and 154. A plate of glass (A) is
held at a distance of 50 /x from the knife by two strips of cellophane
tape (B). The glass plate can be moved on the hinge (C), and the
spring (D) holds the plate against the cellophane strips. The upper
edge of the plate coincides with the knife edge ; screws (F) assist in
the adjustment. The microtome is operated at constant speed and
without stopping for each series of sections to obtain uniform cut-
ting. After the glass plate is swung back, the first section in each
series is discarded since its thickness is different from that of the
others, and the sections to be used are transferred to slides by a thin
glass rod or fine brush.

B. MICROSCOPIC EXAMINATION AND CHEMICAL
ANALYSIS OF THE SAME TISSUE SECTION

In the usual procedure, alternate sections are employed for chem-
ical determination and histological examination in order to corre-
late the analysis with the morphological constitution of the tissue.
This procedure is suitable, provided the histology of the section
analyzed and that of the adjacent section studied microscopically
are essentially the same. When this is not true, as in the case of
retinal tissue which has histological changes every 40 to 50 fx, it is
necessary to carry out the analysis and the microscopy on the same
section. A method for accomplishing this was worked out by An-
finsen et al. (1942), who first stained and examined the section and
then used it for chemical measurement.

MICROSCOPY, CHEMICAL ANALYSIS, VOLUME MEASUREMENT 431

Method of Anfinsen et al. for Microscopy and Analysis on the
Same Tissue Section

In the procedure of Anfinsen et al, frozen sections are cut in the
cryostat of Linderstr0m-Lang and Mogensen, and the sections are
allowed to stand in the cryostat until dry. Faster drying can be
effected by the use of a dehydrating agent, with or without vacuum.
(With phosphorus pentoxide as the desiccant, 20 fi sections of retina
were completely dehydrated within 1-1.5 hr. at —20° at atmospheric
pressure, or within 15-20 min. in vacuo.) A nonaqueous solvent is
then used for mild staining to minimize displacement or solution of
tissue constituents. (A mixture of 1 vol. of 40 milligram per cent
methyl violet in absolute alcohol to 50 vol. xylol was used.) The
stained section is washed with xylol, transferred to a slide, and flat-
tened with a cover slip for visual examination or photomicrography.
After this the cover slip is removed, excess xylol is absorbed on filter
paper, and the section is allowed to dry in the air. Then the dry sec-
tion may be employed for chemical determination. The preceding
treatment was found to have no significant effect on the peptidase
or diphosphopyridine nucleotide in rat liver or the cholinesterase in
rat brain. In some instances it should be possible to examine the
sections directly without staining.

C. VOLUME OF IRREGULARLY SHAPED,
SMALL BIOLOGICAL SAMPLES

The measurement of volume as a means of defining the quantity
of a biological sample has already been discussed (page 423). Holter
(1945) developed a colorimetric method for measuring the volume
of irregular objects of the order of 0.01 to 1.0 /xl. such as the amoeba
Chaos chaos. The simple method of drawing an amoeba into a capil-
lary of known diameter, measuring its length, and computing its vol-
ume, cannot be used with irregular organisms which do not fill the
lumen of the capillary.

In Holter's method, the object is drawn into a capillary tube of
known diameter which is wide enough to avoid deformation of the
object, and some dye solution of known concentration is taken into
the capillary with it. The total length of the object plus dye is meas-

432 AMOUNT OF A BIOLOGICAL SAMPLE

ured and the dye is emptied into a known volume of water. The con-
centration of dye is determined colorimetrically in a microcuvette
and from it the volume of dye that was in the capillary is obtained.
This volume is subtracted from the volume of the dye plus object to
give the volume of the object. In the range 0.01 to 0.1 /A., objects
were measured with a probable error of about ±5% in single meas-
urements.

Holler Method for Measurement of Volume

SPECIAL APPARATUS

Capillaries. Choose capillaries which have a cylindrical constant
bore for at least 10 mm. from one end and which have an inside
diameter suitable for the object to be measured (0.3 mm. for Chaos
chaos). Draw out the uncalibrated end of the capillary to a fine
tip and mount as a braking pipette (Fig. 123). Fire-polish the wide
end of the capillary but do not constrict the mouth. Place a mark
about 5 mm. from the mouth; a thread of DeKhotinsky cement
may be used. Dry the capillary with alcohol and air between
measurements.

Moist Chamber. To prevent evaporation of the small volumes of
solution used, manipulations must be carried out in a moist
chamber (page 181).

SPECIAL REAGENTS

Dye Solution. Dialyze a soln. of acid violet (sodium tetraethyl-di-
p-sulfobenzyl-p^p'-diaminofuchsonimonium) in glass-distilled
water, and bring to a concentration of 1 % . Add sodium taurocho-
late to 0.01% and filter through a fiber-free material. Store in a
refrigerator, and about once every two weeks refilter to remove
dust and dye crystals which may have formed.

PROCEDURE

1. Transfer the object into a moist chamber by means of a brak-
ing pipette (page 359) .

2. Place the object in a flat drop of the dye soln. contained in a
dish.

3. Wash the object with the dye soln. by drawing it up into the
pipette a few times.

.VOLUME MEASUREMENT

433

4. Transfer the object to a fresh flat drop of the dye soln.

5. Draw the object up into the capillary along with an amount
of dye soln. not greater than twice the vol. of the object.

6. Remove the end of the capillary from the drop and draw the
object in the dye soln. about 2 mm. in from the mouth.

7. Dip the mouth of the capillary into water and draw in a
column about 1 mm. long. Remove from the moist chamber.

8. Place a very small drop of mercury (which will fill 0.5-1 mm.
of the capillary) on a watch glass and carefully suck it halfway into
the mouth of the capillary and then push it all the way in with the
finger. The capillary should now look like the illustration in Figure
155.

Mark
r5 mm.

-2

-1

^0

Fig. 155. Capillary for volume measurement, filled.
From Hotter (1945)

9. With the capillary in a vertical position, measure the distance
between the menisci of the dye soln. with a micrometer miscroscope
(Fig. 151) to 0.01 mm. With a living amoeba, it is necessary to use a
filter to remove the heat from the source of illumination so that the
organism will not be disturbed. Movement of the amoeba to a menis-
cus may distort, or obstruct the view of the meniscus.

10. Clean the capillary on the outside with wet filter paper, and
remove the mercury gently by combining it with a large drop of
mercury.

11. Remove the water from the capillary with a tip of a strip of
hard moistened filter paper.

434 AMOUNT OF A BIOLOGICAL SAMPLE

12. Empty the dye soln. plus amoeba into a small reaction tube
containing a measured vol. water (100 fA.). Rinse the capillary with
the water and stir the soln. by means of a magnetic "flea" (page
179). Use a permanent magnet, rather than the periodic electromag-
net, to stir carefully to avoid damage to the organism.

13. Remove the "flea" with the magnet, and the object with a
braking pipette.

14. Transfer the diluted dye soln. to a cuvette and obtain its
colorimeter reading using a yellow filter (S57 with the Zeiss step
photometer) .

15. Run an empirical set of standards with different lengths of
dye columns in the capillary and plot a curve of the colorimeter
readings against the lengths of the dye columns. The standard curve
may be used as long as neither the capillary nor dye soln. is changed.
To check the constancy of the dye soln., run one or two samples of
dye alone, which have known lengths in the capillary, in each ex-
periment.

16. Calculate the vol. of the object (V) by the formula:

V = A{Lt-L)

where ^4 is the cross-sectional area of the capillary, Lt the length of
the dye plus object in the capillary, and L the length of the dye
alone. The value of L, corresponding to the colorimeter reading of the
unknown, is obtained from the standard curve.

VIL DEDUCTIVE METHODS

In certain instances deductive methods can be applied to obtain
histochemical data from macrochemical analyses, i.e., if the total
amount of a constituent in a tissue and the amount in the entire
extracellular portion is known, one can calculate the amount in the
intracellular fraction. This method has been exploited by Lowry
(1943) and the principle may be illustrated by one of his examples:

"As a first approximation, a tissue such as skeletal muscle may be considered
to be composed of 5 separate fractions, blood, fat, collagen plus elastin, extra-
cellular fluid, and cells. If the amount of the first 4 fractions can be determined,
the amount of the remaining intracellular fraction can be calculated. Further-
more, if one knows the composition of the blood and extracellular fluid, it be-
comes possible to calculate the concentration of a particular substance. A, in
the cells by simply (1) measuring the total amount of A, (2) calculating the
amount of A in the several extracellular fractions, (3) subtracting the extra-
cellular A from the whole, and finally, (4) dividing the net intracellular A by
the calculated amount of intracellular fraction. This is similar to the calcula-
tion of the concentration of chloride in red cells when the hematocrit and the
concentration of chloride in whole blood and serum are known."

435

MICROBIOLOGICAL TECHISIQUES

"Perfect as is the wing of a bird, it never could raise
the bird up without resting on air. Facts are the
air of a scientist. Without them you never can fly.
Without them your 'theories' are vain efforts. But
learning, experimenting, observing, try not to stay
on the surface of the facts. Do not become the
archivists of facts. Try to penetrate to the secret of
their occurrence, persistently search for the laws
which govern them."

Pavlov
in Bequest of Pavlov to the Academic Youth of His
Country, Science 83: 3G9 (193G).

INTRODUCTION

The quantitative assay of a number of biologically important
substances, by virtue of their ability to affect the metabolism of
certain microorganisms, is still relatively new. The principle
involved is the measurement of the influence of the substance on
the rate of formation of an end product of the metabolism, such as
the carbon dioxide developed by yeast or the acid formed by
Lactobacillus casei. In other instances the effect on the rate of growth
is measured directly by determining the mass or area of a colony,
as in the case of the assay of choline with a mutant of Neurospora
crassa. At the date of this writing, the microbiological methods have
been developed mainly for members of the vitamin B family and
for amino acids. The great sensitivity of these methods compensates
for the fact that in many cases the substances to be assayed occur
biologically in very high dilutions. The microbiological technique
has been employed almost exclusively on the macro scale. However,
no insurmountable problems are associated with the simple reduction
in volume and the use of the micro techniques already available,
which would be necessary for the adaptation of macro methods to
histological or cytological studies. The riboflavin method of Lowry
and Bessey ( 1944 ) marks a beginning in this direction.

RIBOFLAVIN

The method of Snell and Strong ( 1939) for the microbiological
determination of riboflavin in amounts of the order of 100 m/xg. was
modified by Lowry and Bessey (1944) so that measurements could
be performed in the range of 0.5-2.0 m/xg. with a probable error of
only about 1% for determinations, in triplicate, of pure riboflavin
solutions, and of about 3% for rat cornea. The modifications applied
were based on the observations that riboflavin is partially destroyed
during autoclaving in small tubes at pH 7.0, that air is inhibitory
to, while carbon dioxide stimulates the growth and acid production
of, the Lactobacillus casei used for the assay, that buffers in the pH
range of 4 to 6 stimulate the growth and acid production of the
organism, and that cysteine restores the basal medium if its effective-
ness deteriorates on standing after the autoclaving.

439

440 MICROBIOLOGICAL TECHNIQUES

Lowry and Bessey Method for Riboflavin

SPECIAL REAGENTS AND MATERIALS

Lactobacillus casei Culture and Inoculum. The procedure for
transfer of cultures is shown in the following diagram:

stock culture h^

Stock culture a

^culture d, etc.
culture c

inoculum e —*■ inoculum / — *■ inoculum g, etc.

assay tubes h

Make the stab transfers h, c, d, etc. into yeast-water agar, con-
taining 1% glucose and 0.15 M potassium acetate, and, after 24
hr. incubation at 37°, store in a refrigerator. Reserve at least one
tube, h, as a stock culture. When assays are made on successive
days, do not grow inoculum from stock cultures c and d each day,
but transfer a drop of inoculum e to a similar tube, /, which is
incubated for use the next day. Do not use inoculum cultures
after they are more than 36 hr. old. Return to a stock culture about
every 5 days to minimize the possibility of contamination and
variation. Each month prepare new stock cultures corresponding
to h, c, and d, from a tube such as h of the preceding month. For
inoculum, make a stab from a stock culture into a sterile tube of
basal medium containing 0.5-1.0 ju,g. added riboflavin per 10 ml.
Incubate for 24 hr. at 37°, centrifuge out the cells, and resuspend
in an equal vol. of sterile 0.9% sodium chloride soln., observing
aseptic technique throughout. The most consistent results are
obtained from a culture made from a work stab not over
a week old.
Basal Medium. Consists of the following: photolyzed sodium
hydroxide-treated peptone, 0.5%; glucose, 1%; sodium acetate,
0.6%; cystine, 0.01%; inorganic salts; and riboflavin-free yeast
supplement equivalent to 0.1% yeast extract.

• RIBOFLAVIN 441

(a) Prepare the photolyzed sodium hydroxide-treated peptone
by exposing a mixture of 40 g. of Difco Bacto peptone {Difco
Laboratories, Inc.) in 250 ml. water and 20 g. sodium hydroxide
in 250 ml. water in a 25 cm. crystallizing dish to light from a 100
watt lamp, with reflector, at a distance of about 30 cm. for 6-10
hr. After standing at room temperature for an additional 14-18
hr. the mixture is neutralized with 27.9 ml. glacial acetic acid,
7 g. anhydrous sodium acetate is added, and the mixture is diluted
to 800 ml. Preserve under toluene.

(5) Prepare the yeast supplement by adding a soln. of 150 g.
basic lead acetate in 500 ml. water to 100 g. Bacto yeast extract
(Difco) in 500 ml. water; filter off and discard the precipitate, add
ammonium hydroxide to a pH of about 10, filter off and discard
the precipitate, add glacial acetic acid to just acidify the filtrate,
precipitate excess lead with hydrogen sulfide, and filter off and
discard the lead sulfide. Make up the filtrate to 1000 ml. and store
in a refrigerator under toluene. One ml. of the preparation is
equivalent to 100 mg. of original yeast extract.

(c) Prepare inorganic salt solns. as follows: Solution A —
Dissolve 25 g. potassium monohydrogen phosphate and 25 g.
potassium dihydrogen phosphate in 250 ml. water. Solution B —
Dissolve 10 g. magnesium sulfate heptahydrate, 0.5 g. sodium
chloride, 0.5 g. ferrous sulfate heptahydrate, and 0.5 g. manganese
sulfate tetrahydrate in 250 ml. water.

(d) Combine 50 ml. of the treated peptone soln., 50 ml. 0.1%
cystine hydrochloride, 5.0 ml. yeast supplement, 5.0 g. glucose,
2.5 ml. inorganic salt soln. A, and 2.5 ml. soln. B. Add potassium
acetate to give a concentration of 0.3 M when the mixture is
diluted to 250 ml. With a sterile pipette, add 0.5 ml. of 0.4%
freshly boiled neutral cysteine soln. to each 10 ml. of sterile basal
medium just before inoculation.

Standard Riboflavin Solutions. Prepare standards containing 0.5,
1.0, 1.5, and 2.0 fig. per 100 ml. in 0.002 N hydrochloric acid. Use
0.002 N hydrochloric acid for the blank soln. See step 11 imder
Procedure.

Caprylic Alcohol.

0.1 N Hydrochloric Acid.

0.002 N Hydrochloric Acid.

0.3 N Sodium Hydroxide.

442 MICROBIOLOGICAL TECHNIQUES

0.030 N Sodium Hydroxide.
0.04% Bromotlnjmol Blue.

PROCEDURE

1. Add 0.1 ml. 0.1 A^ hydrochloric acid, either with a 0.2 ml.
graduated pipette drawn out at the end to a slender tip, or with a
constriction pipette (p. 172), to the sample containing 2-8 m/xg.
riboflavin in a 0.75 ml. serological tube (6 X 50 mm.). To clean
these tubes boil in half-cone, nitric acid for a few min. and then in
distilled water several times.

2. Plug tubes with cotton and autoclave for 15 min. at 15 lb.
pressure. Take care to protect from light, particularly while hot.
Check weight of several tubes before and after autoclaving to make
sure no significant volume change occurred.

3. After cooling, add exactly 0.3 ml. 0.030 N sodium hydroxide
with a slender-tipped 1 ml. graduated pipette, or a constriction
pipette, and mix at once by twirling a fine glass rod with a hooked
end in the tube. The soln. now has an excess acid concentration of
about 0.002 M.

4. Transfer a 0.1 ml. aliquot to each of three serological tubes,
and set up, in triplicate, each of the riboflavin standard solns. and
the blank using 0.1 ml. of soln. in each tube.

5. Plug tubes with cotton, wrap entire tube rack in black cloth,
and autoclave for 15 min. at 15 lb. pressure.

6. When the tubes have cooled, remove black cloth, and with a
sterile 1 ml. graduated pipette drawn out at the tip inoculate each
tube with 0.1 ml. basal medium which had been previously inoculated
with 2 drops of the washed bacterial suspension per 10 ml. Replace
cotton plugs and mix by tapping the tubes with the finger.

7. Place rack of tubes in a vacuum desiccator containing damp
cotton swabs. Replace the air with carbon dioxide by alternately
reducing the pressure to about 150 mm. mercury and adding carbon
dioxide back to reach atmospheric pressure. Repeat four to five times
and leave pressure at about 700 mm. mercury. Place the desiccator
in an incubator for 3 days at 38°.

8. After removing tubes from desiccator, add a minute droplet
of caprylic alcohol to each tube to prevent foaming, and blow off
the carbon dioxide by carefully bubbling air through the liquid for
1 min. by means of a capillary tube not over 0.5 mm. in diameter.

. RIBOFLAVIN 443

The bubbles must be small enough so that all spattering is confined
in the tube.

9. Add 0.02 ml. 0.04% bromotliymol blue and titrate with 0.3 N
sodium hydroxide from a 0.2 ml. Rehberg burette. Stir by a stream
of air bubbles. Neither the air bubbler nor the burette tip should
exceed 0.7 mm. diameter. The solubility of the indicator in caprylic
alcohol makes it desirable to use very little of the latter.

10. Plot the calibration curve from the titration values found
using the standard riboflavin solns. Obtain the riboflavin content of
the unknown from its titration value by reference to the calibration
curve.

11. To minimize the effect of possible interfering substances,
and to obtain a more precise assay, prepare the standards as follows
if it is possible: Extract some of the tissue to be analyzed as already
described. Irradiate the extract, which is in 0.002 N hydrochloric
acid, for 30 min. in a Pyrex tube at a distance of 3-4 cm. from a
mercury arc lamp, such as the General Electric HB-4, to destroy the
riboflavin present. With a minimum of dilution, prepare standards
from this extract containing 0, 0.5, 1.0, 1.5, and 2.0 m/xg. riboflavin
per 0.1 ml.

MECHANICAL SEPARATION OF
CELLULAR COMPONENTS

"When one sets out upon the serious and difficult
business of wresting from the universe more knowl-
edge of its character and qualities, his efforts must
all meet with failure unless he can move with perfect
freedom toward the truth wherever the path may
lead. No authority must stay him there; no tradition
perplex him; no dogma, no prejudice, no vested
interest prevent the thorough exploration of every
promising avenue he sees. From its beginning, science
has struggled to be free from all such man-made
bondage."

Edmund W. Sinnott
in Science and the Education of Free Men, American
Scientist 32: 210 (1944).

INTRODUCTION

Centrifugation techniques have become a cytological tool by-
means of which various celkilar components have been separated,
and the isolated constituents have been subjected to chemical study
in certain instances. The value of this procedure is twofold. The
point of view stressed by Bensley ( 1942) that "to separate separable
things before proceeding to their analysis" requires no elaboration
or defense. In addition, sufficient material can be isolated in many
cases to permit chemical investigations on a relatively macro scale,
thus obviating the need for special, and often less available, tech-
niques. Cytochemical work on the separated formed bodies is really
just beginning, and a large proportion of the work still must be
directed toward perfecting the separations themselves.

As Danielli (1946a) has warned, the centrifugal segregation of
particulates from cells could possibly alter the enzyme activities
associated with the particulates m situ. This may or may not be
a factor in specific instances, but in the interpretation of data it
should be kept in mind.

The centrifuge microscope, cleverly designed by E. N. Harvey
and A. L. Loomis, has enabled direct observation of cells at high
magnifications while they are being spun in a centrifuge. In this way
it has been possible to determine the manner in which some cellular
components become segregated under centrifugal force and to
observe the actual scission of certain cells. The instrument and its
application, not only to separations, but also to the determination
of particular physical characteristics, such as surface forces, have
been described by Harvey (1932, 1933).

447

/. TYPES OF HIGH-SPEED CENTRIFUGES

While the ordinary centrifuge and the Sharpies instrument suffice
for many separations of cellular constituents, the ultra- or high-speed
centrifuge offers particular advantages in other instances. It would
carry the present discussion too far afield to give many of the
details of the various types of high-speed centrifuges that have been
employed for cytological work, but the instruments will be mentioned
and the most significant references will be included.

The elaborate Svedberg oil-driven ultracentrifuge, adapted for
measurements of physical characteristics of large molecules and
various particles, has been the subject of a book by Svedberg and
Pedersen (1940) that gives complete details. It has been sufficient
for many cytological separations to employ much simpler apparatus
such as the gas-driven and electrically powered instruments.

The construction of a centrifuge driven by a high-speed electric
motor than can yield a centrifugal force of about 34,800 times
gravity at 18,000 R.P.M. was described by Pickels (1942).

A multispeed attachment may be used with the type SB, size 1,
centrifuge of International Equipment Co. This attachment with
head No. 295 may be used for centrifugal forces up to 18,000 times
gravity. The full capacity of this head is 84 ml. and the celluloid
tubes used with it each has a capacity of 14 ml. (may be obtained
from Lusteroid Container Co., Inc.). The centrifuge without the
multispeed attachment may be used at 1500 times gravity with the
horizontal yoke No. 242, which accommodates bottles of 250 ml.
capacity, or at up to about 2400 times gravity with the conical head
No. 823, which carries 50 ml. tubes.

Gas-Driven "Spinning Top" Centrifuge. This instrument
was developed since 1925 when Henriot and Huguenard first spun
a small rotor (11.7 mm.) at a speed of 660,000 R.P.M. by means of
an air stream that also served as a low-friction cushion to support
the spinning rotor. Many improvements have been made in this
centrifuge (Beams, 1938, 1940, 1941; McBain, 1939). The instrument

448

HIGH-SPEED CENTRIFUGES

449

is essentially as shown in the diagram of Figure 156. Figure 157,
taken from a review by Beams (1942), demonstrates the effect of
the air pressure on the rotor speed for rotors of various designs.
Curves A, B, C and D are for a l^/s in. rotor with superstructures of
various sizes. The highest speed that Beams had obtained by 1942
was almost 1,500,000 R.P.M. with a centrifuge field of almost
8,000,000 times gravity; this was accomplished with a 9 mm. rotor
driven by hydrogen. The substitution of hydrogen for air at the same
pressure produces higher speeds. The usefulness of the gas-driven
centrifuge is greatest when accurate temperature control is not
required, and a large centrifugal field is desirable over a small radial
distance. It has proved valuable for studies of sedimentation
within a cell.

S 6'

Fig. 156. Diagram of air-
driven, air-supported centrifuge.
Left: central section through
stator cone. Right: section
through complete machine. Sta-
tor (S) ; rotor (.R). From Beams
(1H2)

3000

K 2000

O

1000

0 10 20 30 40 50 60
GAGE PRESSURE, lb /m.'

Fig. 157. Relation between rotor
speed and driving air pressure for
rotors shown at right of curves. From
Beams (19.i£)

Plastic-Rotor, Air-Driven Centrifuge. This relatively simple
instrument (Fig. 158), developed by Stern (1942), consists of an
air driven rotor (A) made of a disc of Lucite 0.5 m. thick and 6 in.
diameter that has a center axle (D) made from Vie in. drill rod
which is supported by Torrington needle bearings (E).

A top speed of 17,400 R.P.M., at 48 Ib./in.^ air pressure, giving a
force of 20,200 times gravity has been obtained. The transparency of
the analytical fluid cell has enabled direct observation during the
centrifugation, when a stroboscopic light source and a low-power

450

MECHANICAL SEPARATION OF CELLULAR COMPONENTS

microscope are employed. The instrument is to be made com-
mercially available by Fisher Scientific Co.

Vacuum-Type Air-Driven Centrifuge. In this instrument the
rotor spins in a vacuum on a flexible vertical shaft. Originally
designed by Beams and Pickels (1935), it has since been improved
in various ways as described by Beams ( 1941, 1942) . The advantage

Fig. 158. Air turbine ultracentrifuge with plastic rotor.
From Stem (1942)

Also shown in the figure are: B, analytical fluid cell inserted in cyhndrical
cell hole; C, brass disc connected with similar disc on other side of rotor by
brass bushing and screws; D, fastened to C and turned down and surface-
hardened at ends to fit E mounted in casing, //, and carefully aligned with
bearing on opposite side; F, Fi, brass contacts, inserted in rotor surface; G,
contact brush, made from spring bronze, insulated from casing H, adjustable in
position; H, centrifuge casing, made from sheet brass; /, semicircular opening
in casing H to permit free escape of expanded driving air; J, air jet, '/32 in.
lumen, trumpet-shaped at inlet end and conforming with rotor shape at outlet
end; K, angle for mounting on wooden base. Insert B, analytical fluid cell,
made by cementing, with Lucite cement, two outer discs of colorless Plexiglas
resin to central disc of red Plexiglas into which a sector-shaped opening of
12 mm. height and 3 mm. depth has been cut, connected with periphery by
narrow drill hole, through which the solution under study is introduced with a
hypodermic syringe. When in use the cell is inserted into cell hole in rotor
center and the broad base of the sector pointing toward the periphery.
During operation, the centrifuge is covered by a steel guard, made from 0.5 in.
thick boiler plate by welding, equipped with openings opposite the cell holes
and slots near the base to permit escape of air stream.

EGGS OF ARBACIA PUNCTULATA 451

of this construction over the nonvacuum type is that it obviates the
undesirable effects of air friction and the need for great precision in
the dynamical balance of the rotor. Furthermore, it has the capacity
for handling much greater volumes of material. On the other hand,
it is considerably more complicated mechanically.

Electrically Powered, Magnetically Supported Vacuum-Type
Centrifuge. In order to overcome the irregularities occasioned by
variations in gas pressure in the preceding types of centrifuge,
various methods have been employed to regulate the pressure, but
the most successful among these have been rather complicated.
Hence Skarstrom and Beams (1940) devised an electric centrifuge
in which the rotor was supported magnetically and allowed to spin
in a vacuum. For constancy of speed and ease of operation this
instrument appears to be superior to gas-driven vacuum types.
Refinements of the apparatus have been appearing continuously
from Beams' laboratory at the University of Virginia. In more
recent models the centrifuge is suspended from a vertical iron rod
supported by the field of a solenoid. An automatic regulatory mech-
anism is employed whereby the current in the solenoid is controlled
by the height of the rotor, decreasing as the rotor rises and increas-
ing as it falls. In this fashion the height of the rotor is kept constant
and free from any mechanical contact. Consequently, when the rotor
is spun in a vacuum by a rotating magnetic field, extremely high
speeds can be obtained. With this technique MacHattie (1941)
spun a V32 in. steel ball at 6,600,000 R.P.M., and when the driving
field was cut off at 6,000,000 R.P.M., it lost only about 1% of its
speed in an hour.

//. SEPARATION OF COMPONENTS OF
A, PUNCTULATA EGGS (AFTER HARVEY)

The eggs of the sea urchin, Arbacia punctulata, have been em-
ployed particularly for cytophysical and cytochemical investiga-
tions. Therefore, it may be well to include here a short description
of the procedure developed by Harvey (1932, 1936) for the separa-
tion of components of this cell, especially since similar treatment has
been applied to the ova of other marine invertebrates and to other
cells employed for laboratory study.

Centrifugation is carried out in tubes, a little larger than hema-
tocrit tubes, having a capacity of 0.7 ml. Place the eggs in 1 vol.
sea water over a layer of 2 vol. 0.95 M sucrose soln. (95% of 342 g.
sucrose added to 1 1. tap water) in the centrifuge tube. Roll the tube
gently to effect partial mixing. Centrifuge the eggs in this medium,
which is isosmotic and isopycnotic with the eggs, for 3-4 min. with
a centrifugal force of 10,000 times gravity. Three layers form with
the white halves above, under them a pinkish layer of elongated
whole eggs, and on the bottom the red halves. With more sucrose
solution, further centrifugation results in breaking of the halves
into quarters, as indicated in the diagram in Figure 159.

452

KGGS OF ARBACI A PUNCTULATA

453

Oil

Nucleus
Clear layer

— Mitochondria

White Half
2

White Half

(recentrifuged)

3

cy.:-:rX — Y°ik

Granular Quarter
5

— Pigment

Whole Egg

(centrifugedj

I

Red Half
6

Red Half

(recentrifuged)

7

Pigment Quarter
9

Fig. 159. Unfertilized egg of Arbacia punctulata, stratified by centrifugal
force (about 3 min. at 10,000 g.), and the halves and quarters into which it
breaks. Drawn from camera lucida sketches and photographs, as accurately as
possible to scale (magnified X275). The clear area in 7 at the centripetal pole
is due to further packing of the granules with longer centrifuging. From
Harvey (1936)

///. ISOLATION OF CELL NUCLEI

The isolation of cell nuclei for purposes of chemical investiga-
tion seems to have been attempted first by Miescher (1871), who
digested leucocytes with pepsin to remove the cytoplasm; the
nuclei could then be collected on a filter. Other early attempts at
the separation of nuclei were made by Ackermann ( 1904 ) , who
laked chicken erythrocytes in distilled water and precipitated the
nuclei in 3.6% sodium chloride solution, and Warburg (1910), who
employed a freezing-thawing technique to disrupt the reel cells.

Since a number of methods for the isolation of nuclei have been
evolved during the past 15 years, each with its particular advantages
and drawbacks, and each more suited to certain types of cells than
to others, the most important of these methods will be described in
detail so that the investigator may use his own judgment in their
application.

Since the Warburg (1910) method of freezing and thawing results
in partial agglutination and damage to erythrocytes, and incom-
pletely hemolyzes them, Laskowski (1942) developed a technique
based on hemolysis by lysolecithin. In some instances, as in the case
of lipid studies, lysolecithin treatment would be undesirable; hence
Bounce and Lan (1943) employed saponin for the hemolysis.

A. NUCLEI FROM AVIAN ERYTHROCYTES

Laskowski Procedure for Isolation of Nuclei

Centrifuge the erythrocytes from 30—40 ml. citrated blood, and
wash them several times with isotonic saline soln. by successive
centrifugations and decantations. After the last centrifugation,
pipette out the red cells from the bottom of the layer and suspend
them in 30-40 ml. saline soln. Add 5-8 ml. lysolecithin soln. and
determine when hemolysis is complete by microscopic examination;
it usually takes 20-40 min. at room temperature. Centrifuge the
liquid, and wash the material thrown down five to six times with

454

ISOLATION OF CELL NUCLEI 455

saline soln. by resuspending in 30-40 ml. and centrifuging each time.
The final suspension of nuclei has a faint yellow color, and it is
stable for 2 weeks when stored in a refrigerator. The nuclei may be
stained with aqueous methylene blue. Agglutination occurs when the
suspension is diluted with water but the nuclei may be suspended
safely in 0.1 M potassium dihydrogen phosphate soln.

The lysolecithin is prepared by grinding the poison glands of 100
bees with an emulsion of 5 g. of lecithin in 20 ml. phosphate buffer
(pH 7.0-7.1) and allowing the mixture to digest at 37° for 24 hr.
The material is then filtered through a Berkfeld filter. Snake venom
could be substituted for the bee poison.

NOTE. In a private communication to the writer, R. R. Bensley pointed out
that "it seems a pity to spend the time to pull the stings out of a hundred
honey bees when a few drops of ether added to a suspension of the cells
in salt solution will accomplish the same purpose. As a matter of fact, this
process of hemolysis does not isolate the nuclei since the stroma of the
corpuscle can be demonstrated closely contracted around the surface of
the nucleus."

Bounce and Lan Procedure for Isolation of Nuclei

Wash the red cells from fresh defibrinated blood twice with 0.9%
sodium chloride soln. by centrifuging and decanting. Suspend in a
vol. of the saline soln. equal to that of the blood used and add
5 ml. of 0.11 M phosphate buffer (pH 6.8-7.0), containing 0.3 g.
Merck's purified saponin for each 100 ml. red cell suspension. The
laking is complete in 5 min. Wash the nuclei liberated four to five
times by centrifuging, decanting and resuspending in saline soln.
Each time add 2-3 ml. of the 0.11 M phosphate buffer to the
centrifuged nuclei, without stirring, just before adding the saline
soln.

The observation of Crossmon (1937), that nuclei are ejected from muscle
cells when a bit of the tissue is teased in 5% citric acid, led Stoneburg (1939)
to an application of this principle for the isolation of nuclei from beef heart
muscle, rabbit thigh muscle, tumor cells, and leucocytes. Bounce (1943a) has
pointed out that while the use of 5% citric acid may yield nuclei satisfactory
for studies on lipids, nucleic acids, and acid-resistant nucleoproteins, the low
pH involved denatures enzymes and proteins in general. Hence, Dounce (1943a)
modified the Stoneburg procedure to obtain nuclei suitable for enzj^me and
protein work.

456 MECHANICAL SEPARATION OF CELLULAR COMPONENTS

B. NUCLEI FROM OTHER CELLS

Stoneburg Procedure for Isolation of Nuclei

Nuclei from Muscle Tissue. Grind the tissue, cleaned of visible
fat, as fine as possible in a meat chopper. Place 100 g. in a 1 1. beaker
and cover with 500 ml. of 5% citric acid. Stir occasionally and let
stand overnight. Skim off sm'face fat, dilute soln. with an equal vol.
water, and filter through eight layers of cheesecloth. Run filtrate
through a Sharpies centrifuge at 26,000 R.P.M. The centrifuge
cylinder is lined with cellophane to prevent the solvent action of
the acid on the metal. The material washed off the cellophane sheet
will contain about 50% nuclei. Wash the material four times by
successive centrifugations and resuspensions in water. The nuclei
content will be raised to about 70% by this process. Digest the
residue in the centrifuge bottle with 250 ml. 1% hydrochloric acid
and 250 ml. 0.8% sodium chloride soln. containing 2-4 g. pepsin
(strength 1:10,000) for 4 hr. at 37°. Stir occasionally, and at the
end siphon off the supernatant from the settled nuclei. Wash the
nuclei with distilled water and then centrifuge. A smear of the
residue should show nuclei undamaged histologically and free from
debris.

Nuclei from Rat Tumor Tissue. After removal of necrotic
material the tissue is ground in a meat chopper, treated with citric
acid soln., and filtered as in the case of muscle. Add 1 vol. water to
the filtrate and let stand 3 hr. Omit supercentrifugation but other-
wise continue the procedure given for muscle.

Nuclei from Leucocytes. Add pus to five times its vol. of 5%
citric acid. The nuclei settle rapidly and are then subjected to the
procedure given for tumor tissue.

Dounce Procedure for the Isolation of Nuclei

Nuclei from Rat Liver. Make up approximately equal vol.
cracked ice and water to 500 ml. and add 1.05 ml. 1 M citric acid
(final cone. M/475, pH 6.0-6.2). Place the mixture in a Waring
blendor and add 100 g. frozen rat liver as rapidly as possible without
stalling the stirrer. The liver may be frozen conveniently in the freez-

ISOLATION OF CELL NUCLEI 457

ing compartment of a refrigerator and it should be used as soon as
frozen. Run the blendor until all the ice has melted (10-15 min.).
Strain the mixture through two layers of fine cheesecloth (twenty
threads per cm.) and, when the cloth becomes clogged, replace it
with new material. Wring out the cloths used and strain the liquid
into the already strained batch. Repeat the straining with four
layers of cheesecloth and centrifuge for 20 min. at 1500-2000 R.P.]\I.
in 250 ml. centrifuge bottles. Decant down to the first line demark-
ing the supernatant and the more loosely packed sediment. Add
distilled water to the sediment to make a final vol. of 400 ml.
and stir well to break up lumps. A drop of capryhc alcohol may
be used to break the foam. Centrifuge the suspension for 15
min. and again discard the supernatant fluid. Stir the residue with
about 400 ml. distilled water and centrifuge for 10 min. Wash
sediment with 200 ml. distilled water and centrifuge 5 min. at a
slower speed (1000-1500 R.P.M.). Discard the supernatant and stir
the nuclei with 200 ml. distilled water. Centrifuge for only 3 min.
this time and at a still slower speed. Repeat the washing twice with
200 ml. portions of distilled water and centrifuge 3 min. each time
at the lower speed. Before the last centrifugation, pass the suspension
through four layers of cheesecloth. Stir the nuclei well with about 100
ml. distilled water and let stand for 45 min. in a 100 ml. cylinder.
Carefully decant the top 95 ml. containing most of the nuclei from
the whole cells on the bottom. Recover the nuclei by centrifuging and
resuspending in a small vol. distilled water. The light reddish-brown
color of the final preparation is due to adsorbed hemoglobin.

There are some additional points that should be mentioned. At a
pH 6.5, or higher, the cells rupture in the presence of distilled water
but the purified nuclei do not seem to be harmed. In the pH range of
4.0 to 5.9 cytoplasmic granules agglutinate to a solid mass on cen-
trifugation making it impossible to separate nuclei, while at 3.8 to
4.0 good nuclei preparations are obtained but the acidity results in
some alteration of enzymes and proteins. Adsorbed hemoglobin can
be removed from the product to some extent by one washing, and
completely by two, with Ringer soln. at pH 7.4. However, the Ringer
soln. extracts a certain amount of protein from the nuclei and causes
them to shrink. Subsequently, Dounce (1943b) showed that in the
preparation of nuclei at pH 3.8 to 4.0 the pH tends to rise above 4.0
on successive washings with the result that the nuclei agglutinate.

458 MECHANICAL SEPARATION OF CELLULAR COMPONENTS

This may be prevented by adding a few drops of 1 M citric acid to
tlie wash water each time. Actually less washing is required at pH
3.8 to 4.0 than at 6.0 to 6.2 since the nuclei pack better on centrifug-
ing.

Nuclei from Tumor Tissue. In a later paper Dounce ( 1943c)
described the application of the preceding method to tumor cells. In
the case of Walker carcinosarcoma 256, add 37.6 ml. 1 M citric acid
in the blendor to the 500 ml. of mixture containing 100 g. tissue. Run
the blendor for 15 min. ; the final pH is 3.0. After the first washing,
add a drop or more of 1 M citric acid to prevent agglutination. The
reason for the lower pH in this instance is that cytoplasm cannot be
removed very well from the nuclei at pH 3.8 to 4.0. When hepatoma
31 was used, it was impossible to obtain a good preparation in the
pH range 3.0 to 4.0 necessitating the addition or 100 ml. 1 M citric
acid to the 500 ml. vol. The final pH was 2.4. Here, too, citric acid is
added in the washings to prevent agglutination.

The Dounce pioceduie has been criticized by Hoerr (1943), who claims
that 0.85% sodium chloride is preferable to distilled water for extractions
and washings, and who objects to the use of the Waring blendor as being
too drastic a treatment leading to a certain degree of gelling. While Hoerr
may be right, and even though it is obvious that the less drastic the treatment
of biological systems to attain a given end, the better, the fact that Dounce
obtained good preparations which were suitable for a variety of studies on
enzymes and other nuclear constituents cannot be disregarded. Hoerr favors
the Lazarow (1941, 1943) procedure of breaking up cells by forcing a suspension
of them through bolting silk, and handling the material at a temperature
just above 0° taking care not to let it freeze.

Lazarow Procedure for Isolation of Liver Nuclei
as Used by Hoerr

Perfuse the liver in situ with cold physiological salt soln. by re-
peated variations of hydrostatic pressure from 2 to 4 ft., clamping the
inferior vena cava for 10-20 sec. each min., and massaging the liver
through the abdominal wall while the blood is flowing freely from
the vein. Usually 500-700 ml. saline soln. is required to remove prac-
tically all the blood cells. Remove the liver and chill to 0° as rapidly
as possible, but do not allow to freeze. Triturate the tissue briefly
in a mortar with 0.85% sodium chloride soln. and gently knead
through bolting silk. Suspend the emulsion in 0.5 to 0.7% sodium
chloride soln. at a pH of 6.0 to 6.2. Separate the nuclei from por-

ISOLATION OF CELL NUCLEI 459

tions of this medium by successive centrifugations at 1500 R.P.M.,
carrying out all the operations in a cold room at 0°. Follow the de-
gree of purification by making wet smears of both supernatant and
sediment after each centrifugation. Smears may be stained after fix-
ing in osmic acid vapor.

The principle of separating cellular components by centrifugation in non-
aqueous media (benzene-carbon tetrachloride mixtures) of controlled specific
gravity was first applied by Behrens (1932) to the isolation of nuclei from
calf heart, muscle, and thymus (Behrens, 193S). Fuelgen, Behrens, and
Mahdihassan (1937) used this technique to obtain nuclei from rye germ cells,
and later Behrens (1939) employed the same procedure for the separation of
nuclei from liver cells. In this country Mayer and Gulick (1942) used a
modified Behrens technique to isolate nuclei from bovine thymus cells, and
Williamson and Gulick (1944) applied the method to other cells having a
large proportion of nucleus such as those of human tonsil and bovine super-
mammary lymph gland. The limitation should be borne in mind that the
benzene-carbon tetrachloride mixtures will extract lipids from the nuclei. It
is claimed that proteins are not significantly affected. In general the technique
seems to involve a great deal of time and manipulation, and it is too drastic
a process for many purposes.

Behrens Procedure for Isolation of Nuclei from Thymus
and Lymph Cells (as Modified by Gulick et al.)

Cut the tissue into pea-size bits and freeze in liquid air as soon as
possible after removing from the animal. Dehydrate by treatment
with a number of changes of 10 vol. portions of dry acetone cooled
below — 20°. Remove remaining fat by continuous extraction with
ether that has been dried over anhydrous sodium sulfate; after the
extraction, remove the residual ether in a vacuum desiccator over
cone, sulfuric acid. Grind the dry material in a power mill until
it can pass a 40 mesh sieve. After suspending the powder in a ben-
zene-carbon tetrachloride mixture of specific gravity 1.25, com-
minute further in a ball mill with glass beads. It usually takes 4-8
weeks at 30-50 R.P.M. to effect separation of the nuclear and cyto-
plasmic particles. Test for the completeness of this separation from
time to time by staining a drop of the suspension with hematoxylin
and eosin in the following manner:

Dry a smear of the suspension on a glass slide, and place for 1 min.
in a filtered fresh mixture of equal vol. of 1% yellowish eosin and
Harris ripened alum hematoxylin. Transfer to citric acid-sodium

460 MECHANICAL SEPARATION OF CELLULAR COMPONENTS

phosphate buffer (pH 3.9) and, after 3 min., dry, clear with immer-
sion oil, and examine microscopically. Eosin stains the cytoplasmic
particles, and hematoxylin the nuclear material.

Before separating the suspended cellular components, remove con-
nective tissue by several sedimentations in pure benzene. Discard the
benzene carrying the connective tissue fragments each time. Then
centrifuge the cellular powder from progressively denser mixtures of
benzene and carbon tetrachloride, retaining the sediment in the tube
each time. When the proportion of carbon tetrachloride has been in-
creased to the point where no more powder settles, add benzene until
the denser nuclear fraction begins to come down under centrifuga-
tion. At this final separation, the specific gravity of the suspension
medium was between 1.345 and 1.350 at 28° when bovine thymus
cells were used. Nuclear concentrates prepared in this manner were
found to possess less than 5% contamination. The product is a fine
granular light tan dust that is quite hydroscopic, and it is obtained
in 4.3 to 6.7% yield on the basis of the dry weight of the original
tissue. Highest yields are given by thymus material.

IV. ISOLATION OF CHROMATIN THREADS
FROM CELL NUCLEI

Claude and Potter ( 1943) succeeded in isolating chromatin threads
from the nuclei of spleen cells from leukemic mice and of liver cells
from normal guinea pigs and rats. Apparently these threads are
related to the chromosomes. On standing, especially in very dilute
salt solution, the filaments disintegrate and seem to be replaced by
free refractile granules. In distilled water the threads swell rapidly
and finally disappear.

Claude and Potter Procedure

Care should be taken to conduct all the operations in a cold room
at 0-5°. Grind gently batches of 20-30 g. of the cold tissue with an
equal weight of sand in a mortar for 3 min., and progressively add
six times the tissue weight of either distilled water or 0.9% sodium
chloride soln. having a pH of 7.4. Centrifuge the suspension for 1
min. at 1500 times gravity to throw down the sand and tissue debris.
Centrifuge the supernatant fluid for 10 min. at 1500 times gravity to
sediment the thread-like material, and discard the supernatant. Sus-
pend the material from 25 g. of tissue in 35 ml. of the saline soln. by
gentle shaking, and centrifuge by bringing to the speed correspond-
ing to a force of 1500 times gravity and then cutting off the current
in the centrifuge motor. This short "up-and-down run" throws down
any remaining sand and tissue debris. Subject the supernatant fluid
to another 10 min. centrifugation at 1500 times gravity and follow
by resuspension of the sediment in 35 ml. saline soln., another "up-
and-down run," and a final 10 min. centrifugation of the super-
natant fluid to bring down the white mass of chromatin threads.

461

F. ISOLATION OF CYTOPLASMIC
PARTICULATES

A. MITOCHONDRIA

The mitochondria, dispersed throughout the cytoplasm of cells, are
microscopically visible and have been known for some time. How-
ever, their isolation was not accomplished until 1934 when Bensley
and Hoerr succeeded in separating them from liver cells of the guinea
pig by differential centrifugation. The substantial gap in time be-
tween the recognition of mitochondria as structural entities in the
cytoplasm, and their isolation, was largely due to their highly labile
character, at least in cells of some animals. Mitochondria rapidly
disappear from the cells after death, and they also disappear when
tissue is treated with organic solvents or fluids containing appreciable
acetic acid, and when the temperature is elevated to 48-50°. An
exception has been found, in the case of mitochondria of the liver
cell of Amblystoma, by Bensley and Gersh (1933b), who observed
that, after freezing-drying the tissue, extraction with organic
solvents, treatment with acetic acid, or elevation of temperature
did not cause the disappearance of these bodies. The particles called
"secretory granules" by Claude (1943a) include mitochondria.
Subsequently Claude (1946) referred to particles from liver cells of
0.5-2.0 IX diameter as "large granules" which consist "of secretory
granules and mitochondria." Some additional details for the isolation
procedure of Bensley and Hoerr (1934) have been given by Hoerr

(1943) and are incorporated in the following description. Claude

(1944) and Claude and Fullam (1945) employed certain neoplastic
cells of the rat as a source of mitochondria, and Claude (1946) gave
a detailed description of the separation of particulates from liver.*

Bensley and Hoerr Procedure for Guinea Pig Liver

The entire procedure should be carried out in a cold room at 0°.

Remove blood from the liver by perfusion with cold 0.85% sodium

* Since this writing, Hogeboom et al. published a paper on the isolation of
mitochondria from rat liver; see Bibliography Appendix, Ref. 55.

462

ISOLATION OF CYTOPLASMIC PARTICULATES 463

chloride soln. of pH 6.0 to 6.2. Grind the tissue gently in a mortar
and knead through bolting silk. Suspend the liver emulsion (about
30 g.) from the liver of a 400-500 g. guinea pig in no more than 200
ml. of the physiological saline soln. and centrifuge for 30 min. at
not over 600 R.P.jNI. Centrifuge the supernatant fluid once or twice
at 1500-2000 R.P.M. for not over 1 min. Centrifuge the supernatant
for 10 min. at 2000 R.P.M. and suspend the sediment in 200-300 ml.
of the saline soln. Repeat this washing process until the supernatant
is free of soluble protein; it usually requires four or five washings.

Follow the degree of isolation at various steps by fixing a smear
of the material on a slide wdth osmic acid and staining with aniline-
acid fuchsin and methyl green. The distinct yellowish color of the
unstained mitochondria can also be employed as something of a
check on the separations.

The lower tip of the cake in the centrifuge tube may be contami-
nated with cell fragments ; if so, cut off and discard this portion after
drying. Finally wash the mitochondria with distilled water contain-
ing 1 drop 1 N acetic acid in 200 ml.; centrifuge, and dry the
sediment in the centrifuge tubes in vacuo over phosphorus pentoxide.

During the preceding treatments the mitochondria undergo some
swelling and change from rods to round granules. The granules do
not coalesce and clump unless the centrifugation is at too high a
speed, which results in fragmentation and consequent agglutination.

Claude Procedure for Certain Neoplastic Cells of the Rat

The cells used as the source of mitochondria were obtained from
10-15 g. tumors that develop within 10-12 days after subcutaneous
inoculation of leukemic cells into rats. The advantage of this source
of material is the uniformity of the cell type, the relative lack of
connective tissue, the scant blood supply, and the absence of ap-
preciable necrosis. Furthermore, the mitochondria compose the major
portion of the large cytoplasmic granules.

The following process should be carried out in the cold (2-8°).
Chill freshly removed tumors, pass through a 1 mm. mesh masher,
and grind the pulp for 3-5 min. in a mortar. Add, very slowly at first,
a 0.85% sodium chloride soln., buffered to pH 7.2 with phosphate
having a final concentration of 0.005 M, to a total volume equiva-
lent to five times the weight of tissue pulp. (It is important to main-

464 MECHANICAL SEPARATION OF CELLULAR COMPONENTS

tain the slightly alkaline reaction to prevent clumping of cyto-
plasmic components.) Centrifuge the cellular suspension at 1500
times gravity for 4 min., and follow by a 10 min. run, in order to
throw down most of the debris, unbroken cells, and nuclei. Separate
the mitochondria from the supernatant fluid by centrifuging at 2400
times gravity for 25 min., or else at 18,000 times gravity for 4 min.
Discard the supernatant liquid and resuspend all but the bottom
portion of the sediment in buffered saline soln. Wash the mitochon-
dria by two to three sedimentations and resuspensions in buffered
saline soln. discarding the bottom layer after each centrifugation to
eliminate erythrocytes or nuclei that may have escaped earlier
separation. The final mitochondria suspension consists of granules
0.5-1.5 fx in diameter. The centrifugations were conducted in the type
SB, size 1, centrifuge of the International Equipment Co. (page 448) .

Claude Procedure for Isolation of "Large Granules" from Liver

Rat livers are used which have been depleted of blood by bleeding,
as well as guinea pig livers which have been perfused from the portal
vein or aorta with physiological saline solution. In the latter in-
stance the animals are prepared by placing them under ether anes-
thesia and injecting heparin intravenously, or directly into the heart,
in a dose of 1-2 mg. per 100 g. body weight. All the operations in the
preparation of the extract and the separation of the "large granules"
are carried out in a cold room at around 0°, and all the solutions are
employed at this temperature.

Preparation of Extract. Chill livers immediately after removal
and force through a tissue masher fitted with a 1 mm. mesh screen.
Grind 60-80 g. of the pulp in a mortar about 5 min. and add drop-
wise 0.85% sodium chloride (made slightly alkaline to prevent ag-
glutination by adding 0.2 ml. 1 N sodium hydroxide/1.) until 20-30
ml. have been introduced. Then add the solution more rapidly until
a final volume is obtained equivalent to five times the weight of the
liver pulp used. Centrifuge the suspension at 1500 times gi-avity for
3 min. and discard all the sediment. Centrifuge the supernatant
twice more for 3 min. at 1500 times gravity and discard the deposit
each time. The resulting supernatant liver extract is used for sub-
sequent separations.

Separation of "Large Granules." Centrifuge the liver extract
for 25 min. at 2000 times gravity. Resuspend the sediment, except

ISOLATION OF CYTOPLASMIC PARTICULATES 465

for the portion at the very bottom which consists of nuclei, red cells,
and debris, in a small amount of the "supernate" which has been left
in the tube after decanting the bulk of it. Centrifuge this suspension
for 30 min. at 2000 times gravity and withdraw the "supernate" by
suction with a capillary pipette. Take up the sediment in enough
alkaline saline solution to bring the total volume to one-twelfth that
of the liver extract ; hence the volume at this point is usually 20-25
ml. Leave the small disc of packed debris and nuclei in the tube and
do not suspend it with the rest of the sediment. Finally dilute 15-20
ml. portions of the suspension which contains the "large granules" to
35-45 ml. with alkaline saline and centrifuge for 30 min. at 2000
times gravity. The sediment of "large granules" may be resuspended
in alkaline saline and recentrifuged in the same manner for an
additional washing, and the washing process may be repeated several
times. The "large granule" fraction represents about 10-15% by dry
weight of the total solids in the liver extract. Suspensions of "large
granules" become increasingly acid on standing and may require
additijn of alkali to maintain neutrality.

Instead of the use of 2000 times gravity to effect separation of
"large granules" from the extract a force of 18,000 times gravity for
3-5 min. periods will give better fractionation. Celluloid tubes of 14
ml. capacity were used in the high-speed attachment of the Inter-
national centrifuge (page 448) for this purpose.

B. SUBMICROSCOPIC PARTICULATES

Claude (1940) discovered and separated submicroscopic particu-
lates from saline extracts of embryonic chick tissue, and subse-
quently separated these bodies from a wide variety of other tissues
of various species as well, Claude (1943b). Originally, Claude con-
sidered that these cellular units might be mitochondria or their
fragments, but has since agreed that this is unlikely because the par-
ticles are much too small. Claude (1943a) proposed that the term
"microsome" be applied to this cellular entity; it has also been re-
ferred to by others as the submicroscopic lipoprotein complex. A
second submicroscopic particulate, composed of glycogen, was
demonstrated by Lazarow (1942, 1943) in the guinea pig liver cell.

466 MECHANICAL SEPARATION OF CELLULAR COMPONENTS

Lazarow Procedure for Separation and Isolation of

Lipoprotein and Glycogen Particles from Guinea Pig Liver

Prepare a suspension of fragmented liver cells in the manner de-
scribed for the preparation of mitochondria using a volume of saline
three times that of the liver (page 462) . Continue all operations in a
cold room. Clarify the suspension by spinning for two 10 min. periods
at 3000 R.P.M. in an 8 in. angle centrifuge, followed by 15 min. at
6000 R.P.M. Discard the precipitates after each centrifuging. Trans-
fer the final supernatant fluid to clean 10 ml. Lusteroid test tubes and
centrifuge at 12,000 R.P.M. for 30 min. The sediment consists of a
densely packed cake of the particulate glycogen, and a loosely
packed red precipitate which can easily be separated from the gly-
cogen by inverting the tube. The red precipitate is a mixture of the
glycogen and lipoprotein particles. After removing the red material,
wash the surface of the packed cake twice with physiological saline
soln. and then resuspend in saline allowing 30 min. for dispersion.
Centrifuge at 12,000 R.P.M. for 30 min. and discard the supernatant.
Repeat the resuspension and centrifugation four times. Dry the
final cake in a vacuum desiccator and a clean white powder con-
sisting mainly of glycogen is obtained.

The red precipitate containing both lipoprotein and glycogen can
be freed of the latter by digestion with a purified diastase prepara-
tion. The separated fractions of both submicroscopic particulates
form transparent gelatinous pellets on centrifugation.

Claude Procedure for Isolation of "Microsomes"

All operations are carried out in a cold room at 0° and all solu-
tions employed are cooled to this temperature.

Suspend the ground tissue mass in eight to ten times its weight
of either 0.005 M phosphate buffer, pB. 7.1, or 0.0002 A^ sodium
hydroxide and centrifuge 20 min. at 2400 times gravity. Subject the
supernatant fluid to high-speed centrifugation at about 18,000 times
gravity for 1 hr. Take up the sediment in a little water and centri-
fuge at the top speed for 3-5 min. to throw down the coarser
particles and then resuspend them and again centrifuge for 3-5 min.
Combine the supernatants from both short runs and repeat the
process of a 1 hr. run followed by two short runs two or three times.
In this fashion a concentration of particles ranging in diameter from

ISOLATION 'OF CYTOPLASMIC PARTICULATES 467

60 to 200 ni/x, which will include the "microsomes" will be obtained.
In the case of liver tissue, the following procedure has been fol-
lowed (Claude, 1946) : To the "supernate" obtained when the "large
granules" are separated from the liver extract (page 465) , add 0.1 A^
sodium hydroxide to bring the pH to 7.2 to 7.4. Centrifuge for 4 min.
at 18,000 times gravity and discard the sediment which contains
"large granules" not previously separated. Bring the "supernate" to
pH 7.2 to 7.4 if necessary and spin down the "microsomes" by centri-
fugation for 1.5 hr. at 18,000 times gravity. Wash the "microsome"
fraction which appears as a jellly-like pellet by resuspending in
neutral saline soln. and centrifuging for 1.5 hr. at 18,000 times
gravity. After a second washing the "microsome" yield on a total
solids basis is 10-20% of the original liver extract.

VL ISOLATION OF CHLOROPLASTS FROM

LEAF CELLS

Chloroplastic material was separated from spinach leaves by
Chibnall (1924), who ground the leaves in water, removed liquid by
squeezing through a silk bag, and filtered the fluid through paper
pulp. The chloroplastic material mixed with other cellular constit-
uents was retained on the filter. Menke (1937) found that ammo-
nium sulfate would precipitate chloroplastic material from an acidi-
fied suspension prepared by grinding spinach leaves in water. The
most satisfactory preparations are obtained by differential centrifu-
gation. This procedure was adopted by IMenke (1938), who employed
M/15 phosphate as the suspension medium, Mommaerts (1938) , who
used water containing a little calcium carbonate, Granick (1938),
who found 0.5 M glucose solution particularly suitable, and Neish
(1939) , who also used 0.5 M glucose to obtain intact chloroplasts and
distilled water if merely chloroplastic material was desired. Comar
(1942), fo.'.nd that freezing decreases the solubility of chloroplastic
substance, and he demonstrated that, in a suspension that had been
previously frozen, only a few minutes' centrifugation at 3700 R.P.M.
was required to bring down the substance.

The importance, in some cases, of the time of day the leaves are
picked was emphasized by Hill and Scarisbrick (1940), who found
that if the leaves of Stellaria media were picked at 10:00 A.M. active
chloroplasts could be isolated, but if picked later in the day the ac-
tivity fell, approaching zero on sunny afternoons.

The procedures employed by Granick (1938) and Neish (1939)
will be described. Galston (1943) showed that Granick's method was
applicable to fibrous grass leaves such as those of the oat plant.
Neish successfully employed his method with leaves of Trifolium
pratense, red clover, Elodea canadensis, and Arctium minus, com-
mon burdock, and Onoclea sensibilis, sensitive fern but fibrous or
mucilagenous leaves such as those from couch grass or basswood did
not yield satisfactory preparations.

468

ISOLATION OF CHLOROPLASTS 469

Granick Procedure

Weigh rapidly on a torsion balance 3 g. fresh leaf tissue, exclud-
ing mid ribs and main veins. Place the tissue between wet paper
toweling until ready for use. In this manner the cells absorb water,
become turgid, and are more easily torn apart. After removing from
the paper, wash the tissue with distilled water, dry superficially,
and place some of the material in a 150 ml. porcelain mortar con-
taining 1 g. sand and 25 ml. 0.5 M glucose soln., cooled to about 5°.
Rub gently until the liquid becomes dark green. Add more tissue and
continue the cellular disruption. Pour the suspension into a 50 ml.
centrifuge tube. Add 20 ml. cold glucose soln. to the residue in the
mortar, and after further grinding add the liquid to that in the
centrifuge tube. The time and rate of centrifugation depends on the
leaf material employed. The process and the number of centrifuga-
tions required must be controlled by microscopic examination of the
centrifugates under an oil immersion objective.

Neish Procedure

Remove as much of the fibrous material as possible, wash the leaf
tissue with distilled water, squeeze out excess water by hand, and cut
about 20 g. of the compressed mass with scissors into an 8 in. porce-
lain mortar. ]\Iash to a pulp, add about 200 ml. distilled water in
three portions, grinding after each addition, and filter the mixture
through 200 mesh bolting silk. Centrifuge the filtrate in 250 ml. tubes
at 2000 R.P.M. (With material prepared from sensitive fern a lower
speed is required since the larger chloroplasts in this species sediment
more rapidly.) The starch granules settle faster than the chloro-
plasts and hence may be separated from them at this point. Decant
the supernatant fluid containing the chloroplasts into all. gradu-
ated cylinder until 950 ml. is obtained, then add 2 M calcium chlo-
ride soln. to the 1 1. mark and mix the whole. After 30 min. the floc-
culated chloroplasts settle to about the 200 ml. level. Discard the
supernatant fluid and centrifuge the flocculated material at the same
speed as before. Remove the supernatant fluid and triturate the
chloroplasts with a glass rod fitted wdth a rubber policeman. Repeat
the centrifugations and washings until the concentration of the cal-
cium chloride is reduced to the point at which the chloroplasts again
begin to disperse in the liquid. The material is collected after a final

470 MECHANICAL SEPARATION OF CELLULAR COMPONENTS

centrifugation. To obtain intact chloroplasts, substitute 0.5 M glu-
cose soln. for the distilled water in the procedure and centrifuge the
chloroplasts out at high speed without using the flocculating agent.
(Although nothing is said about it in Neish's paper, the general
experience with chloroplast isolation would suggest that the separa-
tions be carried out in the cold.)

VIL ISOLATION OF OTHER PARTICULATES

FROM CELLS

In addition to those already considered, a wide variety of particu-
lates from different types of cells has been isolated by differential
centrifugation. Without going into the details in each case, a number
of the particulates will be mentioned.

Sevag, Smollens, and Stern (1941) isolated a green particle from
Streptococcus pyogenes by first disrupting the organisms by sonic
vibrations or grinding in a ball mill, precipitating the particles by
66% saturated ammonium sulfate, and finally subjecting the resus-
pended particles to repeated low- and high-speed centrifugations.
The green particulate is sedimented in a gravitational field of
60,000-90,000 times gravity in 1 hr. The final yield amounted to
0.43%.

The iron protein, ferritin, was isolated from horse liver by Stern
and Wyckoff (1938a, 1938b). Several hours' centrifuging at about
70,000 times gravity is required to bring the ferritin down. This
particulate is unique in having an extremely high sedimentation
constant, and an iron content of about 20%. The average particle
size is 10 mfx.

The association of cytochrome oxidase activity with macro-
molecular particles separated from heart muscle was described in
a review by Stern (1943) ; Stern also included a description of the
centrifugal segregation of particles bearing the activity of Rous sar-
coma agent, fowl leukemia virus, and dysentery bacteriophage.

Sonic vibrations, produced by the instrument of Chambers and
Flosdorf (1936), were successfully employed by Henle et al. (1938)
and Zittle and O'Dell (1941) for the disruption of spermatozoa into
heads, midpieces, and tails, which could then be separated by differ-
ential centrifugation. Henle's group found that 7 min. of vibration
at 9000 cycles per second of an intensity that promotes vigorous
cavitation in the fluid was sufficient to break up the sperm of bull,
dog, and rabbit; 15 min. was required for guinea pig sperm and 20
min. for human. The temperature was kept below 20° by water cool-

471

472 MECHANICAL SEPARATION OF CELLULAR COMPONENTS

ing the inside of the nickel vibrating element. Difficulty was expe-
rienced with separation of the parts of guinea pig sperm, since the
heads and tails come down at almost the same rate on centrifugation
giving fractions containing 12-15% of the unwanted portion in the
most successful cases. The contamination found in fractions from
other sources falls in the range of 1-4%. It is easy to obtain the
heads from human sperm since the tails are entirely dispersed by
the vibrations; however, a small part of the midpiece tends to re-
main with the head. The sonic treatment resulted in sc loss of the
acrosome from only the guinea pig sperm.

Claude (1942) succeeded in separating granules of melanin from
the liver of Amphiuma tridactylum by alternate centrifugations of
3 min. at 18,000 times gravity and 1 min. at 1500 times gravity. The
granules were elongated, having dimensions of approximately 1 X
0.5 [x. A small proportion of melanin granules was found in guinea pig
liver, and they bore a striking resemblance to those separated from
the amphibian liver.

BIBLIOGRAPHY

Ackermann, D. (1904). Zur Chemie der Vogelblutkerne. Z. physiol. Cheni.,

43 : 299-304.
Albert. S., and Leblond, C. P. (1946). The distribution of the Feulgen and

2,4-dinitrophenylhydrazine reactions in normal, castrated, adrenalectomized

and hormonally treated rats. Endocrinology, 39 : 386-400.
Allen, R. J. L., and Bourne, G. H. (1943). Some experiments on the micro-
scopical demonstration of zymohexase in animal tissues. J. Expil. Biol.,

20:61-64.
Ammon, R. (1933). Die fermentative Spaltung des Acetylcholins. Arch. ges.

Physiol. Ppigers, 233: 486-491.
Anfinsen, C. B. (1944). A micromanometric method for the determination of

diphosphopyridine nucleotide. J. Biol. Chem., 152: 285-291.
Anfinsen, C. B., Lowry, O. H., and Hastings, A. B. (1942). The application of

the freezing-drying technique to retinal histochemistry. J. Cell. Comp.

Physiol, 20:231-237.
Angeli, B. (1928). Microdetection of phosphorus in plant cells. Riv. biol.,

10: 702-708.
Armitage, F. L. (1939). A modified peroxidase stain for blood and bone

marrow films. J. Path. Bad., 49: 579-580.
Arnon, D. I., Stout, P. R., and Sipos, F. (1940). Radioactive phosphorus as an

indicator of phosphorus absorption of tomato fruits at various stages of

development. Am. J. Botany, 27: 791-798.
Atkin, L., Schultz, A. S., and Frey, C. N. (1939). Ultramicrodetermination of

thiamine by the fermentation method. J. Biol. Chem., 129:471-476.

Baker, J. R. (1942). Some aspects of cytological technique, in Bourne, G.,

Cytology and Cell Physiology. Clarendon Press, Oxford, p. 1.
Barnett, S. A., and Bom-ne, G. (1941). Use of silver nitrate for the histo-

chemical demonstration of ascorbic acid. Nature, 147:542-543.
Barth, L. G., and Kirk, P. L. (1942). A simplified differential microrespiro-

meter. J. Gen. Physiol., 25: 663-668.
Bates, J. C. (1942). On the structure and staining of starch grains of the

potato tuber. Stain Technol., 17: 49-56.
Bayliss, L. E., and Walker, A. M. (1930). The electrical conductivity of

glomerular urine from the frog and from Necturis. J. Biol. Chem.,

87:523-540.
Beams, J. W. (1938). High-speed centrifuging. Rev. Mod. Phys., 10: 245-263.
Beams, J. W. (1940). The ultracentrifuge, in Baitsell, G. A., Science in

Progress, Series 2. Yale Univ. Press, New Haven, pp. 232-264.
Beams, J. W. (1941). High-speed centrifuging. Reports on Progress in

Physics, The Physical Society, Great Britain, 8: 31-49.

473

474 BIBLIOGRAPHY

Beams, J. W. (1942). The production and maintenance of high centrifugal

fields for use in biology and medicine. Ann. N. Y. Acad. Sci., 43: 177-193.
Beams, J. W., and Pickels, E. G. (1935). The production of high rotational

speeds. Rev. Sci. InMruments, 6: 299-308.
Beecher, H. K., Follansbee, R., Murphy, A. J., and Craig, F. N. (1942).

Determination of the oxj'gen content of small amounts of body fluids by

polarographic analysis. J. Biol. Cheni., 146: 197-206.
Behrens, M. (1932). Untersuchungen an isolierten Zell- und Gewebsbestand-

teilen. Mitt. I. Isolierung von Zellkernen des Kalbherzmuskels. Z. physiol.

Chem., 209: 59-74.
Behrens, M. (1938). Zell- und Gewebetrennung, in Abderhaldcn, E., Hand-

buch der biologischcn Arbeitsmethoden. Urban & Schwarzenberg, Berlin

and Vienna, Abt. V, Teil 10, pp. 1387-1389.
Behrens, M. (1939). Uber die Verteilung der Lipase und zwischen Zellkern

und Protoplasma der Leber. Z. physiol. Chem., 258: 27-32.
Behrens, B., and Baumann, A. (1933a). Zur Pharmakologie des Bleis. IX

Mitteil. Weitere Untersuchungen iiber die Verteilung des Bleis mit Hilfe

der Methode der Autohistoradiographie. Z. ges. exptl. Med., 92: 241-250.
Behrens, B., and Baumann, A. (1933b). Zur Pharmakologie des Bleis. X

Mitteil. Die Beziehung der Bleiablagerung zum Calciumstoffwechsel.

Z. ges. exptl. Med., 92: 251-264.
Belanger, L. F., and Leblond, C. P. (1946). A method for locating radioactive

elements in tissues by covering histological sections with a photographic

emulsion. Endocrinology, 39 : 8-13.
Benedetti-Pichler, A. A. (1937). Qualitative analysis of microgram samples.

General technic. Ind. Eng. Chem., Anal. Ed., 9: 483-487.
Benedetti-Pichler, A. A., and Cefola, M. (1942). Qualitative analysis of

microgram samples. General technique and confirmatory tests. Ind. Eng.

Chem., Anal. Ed., 14: 813-816.
Benedetti-Pichler, A. A., and Rachele, J. R. (1940). Limits of identification

of simple confirmatory tests. Ind. Eng. Chem., Anal. Ed., 12: 233-241.
Benedict, S. R. (1914). Studies in creatine and creatinine metabolism. I. The

preparation of creatine and creatinine from urine. J. Biol. Chem., 18:

183-190.
Benedict, S. R. (1929). A note on the purification of picric acid for creatinine

determination. J. Biol. Chem., 82: 1-3.
Bennett, H. S. (1939). Localization of adrenal cortical hormones in the

adrenal cortex of the cat. Proc. Soc. Exptl. Biol. Med., 42: 786-788.
Bennett, H. S. (1940). The life history and secretion of the cells of the

adrenal cortex of the cat. Am. J. Anat., 67: 151-228.
Bensley, C. M. (1939). Comparison of methods for demonstrating glycogen

microscopicallJ^ Stain Technol., 14: 47-52.
Bensley, R. R. (1942). Chemical structure of cytoplasm. Science, 96: 389-393.
Bensley, R. R., and Bensley, S. H. (1938). Handbook of Histological and Cy to-
logical Technique. Univ. Chicago Press, Chicago.
Bensley, R. R., and Gersh, I. (1933a). Studies on cell structure by the freezing-

drjdng method. I. Introduction, Ayiat. Record, 57: 205-215.

. BIBLIOGRAPHY 475

Bensley, R. R., and Gersh, I. (1933b). Studies on cell structure by the freezing-
drying method. II. The nature of the mitochondria in the hepatic cell of
ambh^stoma. Anat. Record, 57: 217-233.

Bensley, R. R., and Hoerr, N. L. (1934). Studies on cell structure by the
freezing-drying method. VI. The preparation and properties of mito-
chondria. Anat. Record, 60: 449-455.

Bentley, G. T., and Kirk, P. L. (1936). Quantitative drop analysis. VI. Total
nitrogen by diffusion. Mikrochemie, 21: 260-267.

Berenblum, I., Chain, E., and Heatley, N. G. (1939). The study of metabolic
activities of small amounts of surviving tissues. Biochem. J., 33: 68-74.

Berenblum, I., Chain, E., and Heatley, N. G. (1940). The metabolism of normal
and neoplastic skin epithelium. Ain. J. Cancer, 38: 367-371.

Berg, W. (1926). Zum histologischen Nachweis der Eiweisspeicherung in der
Leber. Arch, ges Phijsiol. Ppigers, 214: 243-249.

Berg, W. E. (1946). A microanalyzer for very small gas samples (0.4-1 mm.^).
Science, 104: 575-576.

Bessey, O. A. (1938). A method for the determination of small quantities of
ascorbic acid and dehydroascorbic acid in turbid and colored solutions in
the presence of other reducing substances. J. Biol. Chem., 126: 771-784.

Bessey, 0. A., Lowry, O. H., Brock, M. J., and Lopez, J. A. (1946). The deter-
mination of vitamin A and carotene in small quantities of blood serum.
J. Biol. Chem., 166: 177-188.

Blanchetiere, A., and Romieu, M. (1931). La reaction histochemique de
Romieu pour les substances proteiques, ses conditions d'obtention, son
mechanisme et sa signification. Conipt. rend. soc. biol., 107: 1127-1129.

Blanchetiere, M. A. (1923). Sur une methode de dosage du sodium. Bull. soc.
chim.Belg.,55: 807-818.

Bloom, W., and Bloom, M. A. (1940). Calcification and ossification. Calcifica-
tion of developing bones in embryonic and newborn rats. Anat. Record,
78: 497-523.

Bock, E., and Oesterlin, M. (1939). Ueber einige fluoreszenzmikroskopische
Beobachtungen. Zentr. Bakt. Parasitenk. Abt I, Orig., 143: 306-318.

Boell, E. J. (1945). Technique for the estimation of total nitrogen in tissues
and tissue extracts. Trans. Connecticut Acad. Arts Sci., 36: 429-441.

Boell, E. J., Koch, H., and Needham, J. (1939). Morphogenesis and metabolism:
Studies with the Cartesian diver ultramicromanometer. III. Respirators^
rate of the regions of the amphibian gastrula. IV. Respiratorj' quotient of
the regions of the amphibian gastrula. Proc. Roy. Soc. London, B127:
363-387.

Boell, E. J., and Needham, J. (1939). Morphogenesis and' metabolism: Studies
with the Cartesian diver ultramicromanometer. II. Effect of dinitro-o-
cresol on the anaerobic glycolysis of the regions of the amphibian gastrula.
Proc. Roy. Soc. London, B127: 356-362.

Boell, E. J., Needham, J., and Rogers, V. (1939). ]Morphogene?is and metab-
olism: Studies with the Cartesian diver ultramicromanometer. I. Anaerobic
glycolysis of the regions of the amphibian gastrula. Proc. Roy. Soc. London,
B127: 322-356,

476 BIBLIOGRAPHY

Boettiger, E. G. (1946). Diphenjdamine as colorimetric agent in the determina-
tion of glycogen. J. Cell. Comp. Physiol., 27: 1-8.

Bolomey, R. A., and Kemmerer, A. R. (1946). The determination of ascorbic
acid; a simplification of the Roe method. J. Biol. Chem., 165: 377-378.

Booth, R. G. (1944). Cereal phosphatases. I. The assay of free wheat phos-
phomonoesterase and characterization of the free phosphatases of wheat.
Biochem. J., 38: 355-362.

Bordley, J., Ill, Hendrix, J. P., and Richards, A. N. (1933). Quantitative studies
of the composition of glomerular urine. XI. The concentration of creatinine
in glomerular urine from frogs determined by an ultramicroadaptation of
the Folin method. J. Biol. Chem. 101: 255-267.

Bordley, J., Ill, and Richards, A. N. (1933). Quantitative studies of the
composition of glomerular urine. VIII. The concentration of uric acid in
glomerular urine of snakes and frogs, determined by an ultramicro-
adaptation by Folin's method. J. Biol. Chem., 101: 193-221.

Borsook, H. (1935). Micromethods for determination of ammonia, urea, total
nitrogen, uric acid, creatinine (and creatine), and allantoin. J. Biol. Chem.,
110: 481-493.

Borsook, H., and Dubnoff, J. W. (1939). Methods for the determination of
submicro quantities of total nitrogen, ammonia, amino nitrogen, amides,
peptides, adenylic acid, and nitrates. ./. Biol. Chem., 131: 163-176.

Bott, P. A. (1943). Quantitative studies of the composition of glomerular urine.
XV. The concentration of sodium in glomerular urine of Neclun. J. Biol.
Chem., 147: 653-661.

Bottelier, H. P., Holter, H., and Linderstr0m-Lang, K. (1943). Studies on
enzymatic histochemistry. XXXVI. Determination of peptidase activity,
nitrogen content and reduced woigiit in roots of the barley, Hordeum
vulgare. Compt. rend. trav. lab. Carlsherg, Ser. chim., 24: 289-313.

Bouine, G. (1935). Mitochondria, Golgi apparatus and vitamins. Australian J.
Exptl. Biol. Med. Sci., 13: 238-249.

Bourne, G. (1936). The vitamin C technique as a contribution to cytology.
Anat. Record, 66: 369-385.

Bourne, G. (1942). Cytology and Cell Physiology. Clarendon Press, Oxford.

Bourne, G. (1943). The distribution of alkaline phosphatase in various tissues.
Quart. J. Exptl. Physiol, 32: 1-19.

Brachet, J. (1940). La detection histochemique des acides pentosenucleiques.
Compt. rend. soc. bioL, 133: 88-90.

Brandt-Rehberg, P. (1925). A method of microtitration. Biochem. J., 19:
270-277.

Broda, B. (1939). Uber die Verwendbarkeit von Chinahzarin, Titangelb und
Azoblau zum mikro- und histochemischen Magnesiumnachweis in Pflan-
zengeweben. Mikrokosmos, 32. 184.

Briiel, D., Holter, H., Linderstr0m-Lang, K., and Rozits, K. (1946). A micro-
method for the determination of total nitrogen (accuracy 0.005 fig. N).
Compt. rend. trav. lab. Carlsberg, Ser. chim., 25: 289-324; Biochim. et
Biophys. Acta, 1: 101-125 (1947).

BIBLIOGRAPHY 477

Busch, M. (1905). Gravimetrische Bestiiumung der Saltpetersauie. Ber., 38:

861-866.
Butler, A. M., and Tuthill, E. (1931). An application of the uranyl zinc acetate

method for determination of sodium in biological material. J. Biol. Chem.,

93: 171-180.

Cameron, G. R. (1930). Tlie staining of calcium. J. Path. Bad., 33: 929-955.
Carere-Comes, O. (1938). Neue Methoden zum histochemischen Nachweis des

Kahums und zur elektiven Fiirbung der kahreichen Gewebe. Z. vnss.

Mikroskop., 55: 1-7.
Carruthers, C. (1943a). Ultramicro method for sodium employing the polaro-

graph. hid. Eng. Chem., Anal. Ed., 15: 70-71.
Carruthers, C. (1943b). Microdetermination of magnesium with the polarograph.

Ind. Eng. Chem., Anal. Ed., 15: 412-414.
Carruthers, C. (1945). Microdetermination of copper with the polarograph.

Ind. Eng. Chem., Anal Ed., 17: 398-399.
Caspersson, T. (1940). Methods for the determination of the absorption

spectra of cell structures. J. Roy. Microscop. Soc, 60: 8-25.
Castel, P. (1934r-35a). Contribution a I'etude de la localisation histochimique

de certaines substances medicamenteuses. I. L'arsenic. Arch. soc. sci. med.

biol. Montpellier et Languedoc, 16: 430-435.
Castel, P. (1934-35b). Contribution a I'etude de la localisation histochemique

de certaines substances medicamenteuses. II. Le Bismuth. Arch. soc. sci.

med. hiol. Montpellier et Languedoc, 16: 453-456.
Castel, P. (1936). Recherches sur la detection histochimique du bismuth. Bull.

histol. appl. physiol. et path, et tech. m,icroscop., 13: 290-297.
Chambers, L. A., and Flosdcrf, E. W. (1936). Sonic extraction of labile bac-
terial constituents. Proc. Soc. Exptl. Biol. Med., 34: 631-636.
Chambers, R., and Kopac, M. J. (1937). Micrurgical technique for the study

of cellular phenomena, in McClung, Handbook of Microscopic Technique.

2nd ed., Hoeber, New York, pp. 62-109.
Chevremont, M., and Comhaire, S. (1939). Detection cytochimique de lacto-

flavine dans le foie de cobaye et etude de ses variations provoquees par le

cyclopentyldinitrophenol. Arch exptl. Zellforsch. Gewebeziicht., 22: 658-664.
Chibnall, A. C. (1924). Spinacin, a new protein from spinach leaves, J. Biol.

Chem., 61 : 303-308.
Christeller, E. (1926). (a) Histochemischer Nachweis des Wismuts in den

Organen. (b) Histochemischer Differenzierung der Gewebe mittels Eisen-

salzbildung. Med. Klin. Munich, 22: 619-621.
Claff, C. L., and Swenson, 0. (1944). Micro glass electrode technique for

determination of hydrogen ion activity of blood and other biological fluids.

J. Biol. Chem., 152: 519-522.
Clark, W. G., Levitan, N. I., Gleason, D. F., and Greenberg, G. (1942). Titri-

metric microdetermination of chloride, sodium and potassium in a single

tissue or blood sample. J. Biol. Chem., 145: 85-100.
Claude, A. (1940). Particulate components of normal and tumor cells. Science,

91 : 77-78.

478 BIBLIOGRAPHY

Claude, A. (1941). Particulate components of cytoplasm. Cold Spring Harbor

Symposia Quant. Biol., 9: 263-270.
Claude, A. (1942). Mechanical separation of the morphological constituents of

the cell. Trans. N. Y. Acad. Sci., 4: 79-83.
Claude, A. (1943a). The constitution of protoplasm. Science, 97: 451-456.
Claude, A. (1943b). Distribution of nucleic acids in the cell and the morpho-
logical con.stitution of cytoplasm. Biol. Symposia, 10: 111-129.

Claude, A. (1944). The constitution of mitochondria and micro.somes, and the
distribution of nucleic acid in the cytoplasm of a leukemic cell. J. Exptl.
Med.,^Q: 19-29.

Claude, A., and Fullam, E. F. (1945). An electron micro,scope study of isolated
mitochondria. Method and preliminar.y results. J . Exptl. Med., 81 : 51-62.

Claude, A., and Potter, J. S. (1943). Isolation of chromatin threads from the
resting nucleus of leukemic cells. J. Exptl. Med., 77: 345-354.

Cole, P. A., and Brackett, F. S. (1940). Technical requirements in the deter-
mination of absorption spectra by the ultraviolet microscope. Rev. Sci.
Instruments, 11: 419-427.

Coleman, L. C. (1938). Preparation of leuco basic fuchsin for use in the Fuel-
gen reaction. Stain Technol., 13: 123-124.

Colwell, R. N. (1942). The use of radioactive phosphorus in translocation
studies. Ayn. J. Botany, 29: 798-807.

Comar, C. L. (1942). Chloroplast substance of spinach leaves, Botan. Gaz., 104:
122-127.

Conway, E. J. (1934). A horizontal micro-burette. Biochem. J., 28: 283-287.

Conway, E. J. (1935a). An absorption apparatus for the micro-determination
of certain volatile substances. III. The micro-determination of chloride,
with application to blood, urine and tissues. Biochem. J., 29: 2221-2235.

Conwa}'-, E. J. (1935b). Apparatus for the micro-determination of certain votatile
substances. IV. The blood ammonia, with observations on normal human
blood. Biochem. J., 29: 2755-2772.

Conway, E. J., and Byrne, A. (1933). An absorption apparatus for the micro-
determination of certain volatile substances. I. The micro-determination of
ammonia. Biochem,. J ., 27: 419-429.

Cowdry, E. V. (1928). The microchemistry of nuclear inclusions in virus diseases.
Science, 68: 40-41.

Cowdry, E. V. (1943). Microscopic Technique in Biology and Medicine.
Williams & Wilkins, Baltimore.

Cramer, G. (1940). Ein Verfahren, Nitrate im Gewebe sichtbar zu machen.
Zentr. allgern. Path. u. path. Anat., 74: 241-244.

Cretin, A. (1924). Sur un nouveau reactif du calcium applicable aux recherches
histologiques. Bull, histol. appl. physiol. et path, et tech. microscop., 1:
125-132.

Cretin, A. (1929). Note sur la detection histochimique du plomb. Compt. rend,
assoc. anat., 24: 171.

Cretin, A., and Pouyanne, L. (1933). Action de quelques metaux sur la con-
solidation osseuse. (Contribution a I'etude de I'osteosynthese.) Bordeaux
chirurgical, 4: 321—364.

BIBLIOGRAPHY 479

Crossmon, G. (1937). The isolation of muscle nuclei. Science, 85: 250.
Cunningham, B., and Kirk, P. L. (1940). A new form of differential micro-

respirometer. J. Gen. Physiol, 24: 135-149.
Cunningham, B., and Kirk, P. L. (1942). The oxj^gen consumption of single

cells of Paramecium caudalum as measured by a capillary respirometer.

/. Cell. Comp. Plnjsiol, 20: 119-134.
Cunningham, B., Kirk, P. L., and Brooks, S. C. (1941a). Quantitative drop

analysis. XIV. Potentiometric determination of chloride. J. Biol. Chem.,

139: 11-19.
Cunningham, B., Kirk, P. L., and Brooks, S. C. (1941b). Quantitative drop

analysis. XV. Determination of potassium. J. Biol. Chem., 139: 21-28.
Custers, J. F. H. (1933). Eine evakuierte Verstarkeranordnung zur Messung

kleiner Photostrome. Z. tech. Physik., 14 : 154-157.

Danielli, J. F. (1946a). Establishment of cytochemical techniques. Nature,

157: 755-757.
Danielli, J. F. (1946b). A critical stud}^ of techniques for determining the

cytological position of alkaline phosphatase. J. Exptl. Biol., 22: 110-117.
Daniels, A., and Heidt, L. J. (1932). Photochemical technique. I. A simple

capillarj' mercury vapor lamp. J. Aiyi. Chem. Soc, 54: 2381-2384.
Davies, P., and Brink, F. (1942). Microelectrodes for measuring local oxygen

tension in animal tissues. Rev. Sci. Instruments, 13: 524-533.
Dean, R. B. (1941). The determination of chloride in single isolated muscle

fibers. /. Biol. Chem., 137: 113-121.
Dean, R. B., and Fetcher, E. S., Jr. (1942). Micrometer burette. Science, 96:

237-238.
Deane, H. W., Nesbett, F. B., and Hastings, A. B. (1946). Improved fixation

for histological demonstration of glycogen and comparison with chemical

determination in liver. Proc. Soc. Exptl. Biol. Med., 63: 401-406.
Deckert, W. (1935). Colorimetrische Zinkbestimmung mit Dithizon. Z. anal.

Chem., 100: 385-390.
De Ment, J. (1942). Fluorescent Chemicals and Their Applications. Chemical

Pub. Co., Brooklyn.
Dempsey, E. W., and Deane, H. W. (1946). The cytological locaHzation, sub-
strate specificity, and pH optima of phosphatases in the duodenum of the

mouse. J. Cell. Comp. Physiol., 27: 159-179.
Dempsey, E. W., and Singer, M. (1946). Observations on the chemical cj'-fology

of the thyroid gland at different functional stages. Endocnnology, 38: 270-

295.
Dempsey, E. W., and Wislocki, G. B. (1945). Histochemical reactions associated

with basophilia and acidophilia in the placenta and pituitary gland. Am.

J. Anat., 76: 277-301.
•Dempsey, E. W., and Wislocki, G. B. (1946). Histochemical contributions to

physiology. Physiol. Revs., 26: 1-27.
Dhere, C. (1939). La spectrochimie de fluorescence dans I'etude des produits

biologiques. Fortschr. Chem,. org. Naturstoffe, 2: 301-341.
Dixon, M. (1943). Manometnc Methods as Applied to the Measurement of

480 • BIBLIOGRAPHY

Cell Respiration and Other Processes. 2nd ed., Cambridge Univ. Press,

London.
Dobriner, K., and Rhoads, C. P. (1940). The porphyrins in health and disease.

Phijsiol. Revs., 20: 416-468.
Dols, M. J. L., Jan.sen, B. C. P., Sizoo. G. J., and van der Maas, G. J. (1938).

Distribution of phosphorus in the leg bones of chickens. Nature, 142:

953-954.
Doniach, I., Mottram, J. C, and Weigert, F. (1943). The fluorescence of 3,4-

benzpyrene in vivo. I. The distribution of fluorescence at various sites,

especially the skin of mice. Brit. J. Exptl. Path., 24: 1-9.
Dounce, A. L. (1943a). Enzyme studies on isolated cell nuclei of rat liver.

J. Biol. Chem., 147: 685-698.
Dounce, A. L. (1943b). Further studies on isolated cell nuclei of normal rat

liver. J. Biol. Chcm., 151 : 221-233.
Dounce, A. L. (1943c). The desoxyribonucleic acid content of isolated nuclei

of tumor cells. J. Biol. Chem., 151: 235-240.
Dounce, A. L., and Lan, T. H. (1943). Isolation and properties of chicken

erythrocyte nuclei. Science, 97: 584-585.
Dubhn, W. B. (1943). Bodian method applied to demonstration of melanin.

Aju. J. Clin. Path., Tech. Sect., 7: 127-128.
Du Bois, D. (1931). A machine for pulling glass micropipettes and needles.

Science, 73: 344-345.
Dubos, R. J. (1937). The decomposition of yeast nucleic acid by a heat resistant

enzyme. Science, 85: 549-550.
Dunn, R. C. (1946). A hemoglobin stain for histologic use based on the cyanol-

hemoglobin reaction. Arch. Path., 41 : 676-677.
Dunn, R. C, and Thompson, E. C. (1945). A new hemoglobin stain for histo-
logic use: a slightty modified Van Gieson stain. Arch. Path., 39: 49-50.
Dunn, R. C, and Thompson, E. C. (1946). A simplified stain for hemoglobin

in tissues or smears using patent blue. Stain Technol., 21 : 65-67.

Edwards, G. A., Scholander, P. F., and Roughton, F. J. W. (1943). Micro
gasometric estimation of the blood gases. III. Nitrogen. /. Biol. Chem...
148: 565-571.

Elftman, H., and Elftman, A. G. (1945). Histological methods for the demon-
stration of gold in tissues. Stain Technol., 20: 59-62.

Ellinger, P. (1938). Lyochromes in the kidney. With a note on the quantitative
estimation of lyochromes. Biochem. J., 32: 376-382.

Ellinger, P. (1940). Fluorescence microscopy in biology. Biol. Revs. Cambridge
Phil. Soc, 15: 323-350.

Ellinger, P., and Koschara, W. (1933). tJber eine neue Gruppe tierischer Farb-
stoffe (Lyochrome). I, II, III Mitteil. Ber. 66: 31^317, 808-813, 1411-1414.

Engstrom, A. (1944). The localization of mineral salts in striated muscle fibres.
Acta Physiol. Scand., 8: 137-151.

Engstrom, A. (1946). Quantitative micro- and histochemical elementary
analysis by roentgen absorption spectrography. Acta Radiol., Suppl. 63:
106 pp.

BIBLIOGRAPHY 481

Elder, H. von, Hellstrom, H., and Adler, E. (1935). Fluoreszenmikrr.skopische
Studien iiber das Flavin in Augen. Z. verleich. Physiol., 21: 739-750.

Fautrez, J., and Lambert, P. P. (1937). Une nouvelle methode de coloration
histologique de I'liemoglobine. Bull. hist. appl. physiol. et path, et tech.
microscop., 14: 29-31.

Feulgen, R., and Behrens, M. (1938). Zur kenntnis des Plasmalogens. III. Eine
v/eitere Methods zur Identifizierung des Plasmals. Z. physiol. Chem., 256:
15-20.

Feulgen, R., Behrens, M., and Mahdihassan, S. (1937). Darstellung und Identi-
fizierung der in den pflanzlichen Zellkernen vorkommenden Nucleinsaure.
Z. physiol. Chem., 246: 203-211.

Feulgen, R., and Behrens, M. (1938). Zur Kenntnis des Plasmalogens. III. Eine
neuartige Gruppe von Phosphatiden (Acetolpho.sphatide). Z. physiol.
Chem., 260: 217-245.

Feulgen, R., and Voit, K. (1924). Uber einen weitverbreiteten festen Aldehyd;
Seine Entstehung aus einer Vorstufe, sein mikrochemischer und mikro-
skopisch-chemischer Nachweis und die Wege zu seiner praparativen Dar-
stellung. Arch ges. Physiol. P fingers, 206: 389-410.

Fischl, v., and Singer. E. (1935). Die Wirkungsweise chemotherapeutisfh
verwendeter Farbstoffe. Z. Hyg. Injektionskrankh., 116: 348-355.

Folin, O. (1930). An improved method for the determination of uric acid m
blood. J. Biol. Chem., 86: 179-187.

Forsgren, E. (1928). Mikroskopische Untersuchungen iiber die Gallenbildung
in den Leberzellen. Z. Zelljorsch. mikroscop. Anat., 6: 647-688.

Frankenberger, Z. (1921). Recherches histologiques sur la localisation du plomb.
Compt. rend, assoc. anat., 16: 241-244.

Fujita, H. (1939). trber die Mikrobestimmung der Blutphosphatase. J. Bio-
chem,. Japan, 30: 69-87.

Gage, S. H. (1938). Apparatus and methods for microincineration. Stain Tech-
nol, 13: 25-36.

Galston, A. W. (1943). The isolation, agglutination and nitrogen analysis of
intact oat chloroplasts. Am. J. Botany, 30: 331-334.

Gerard, P., and Cordier, R. (1932). Etudes histophysiologiques sur le rein des
Anoures. Arch. biol. Liege, 43: 367-413.

Gerard, R. W., and Hartline, H. K. (1934). Respiration due to natural nerve
impulses. A method for measuring respiration. J. Cell. Comp. Physiol., 4:
141-159.

Gerlach, W., and Gerlach, W. (1933). Die chemische Emissionsspektralanalyse.
II. Anwendung in Medizin, chemie, und mineralogie, Voss, Leipzig. Trans-
lated by Twyman, J. H. (1934), Clinical and Pathological Applications of
Spectrum Analysis, Hilger, London.

Gersh, I. (1932). The Altmann technique for fixation by drying while freezing.
Anat. Record, 53: 309-337.

Gersh, I. (1938). Improved histochemical methods for chloride, phosphate-
carbonate and potassium applied to skeletal muscle. Anat. Record, 70:

2" 329 .<^»C^2^

482 BIBLIOGRAPHY

Gersh, I., and Baker, R. F. (1943). Total protein and organic iodine in the

oolloid of individual follicles of the thyroid gland of the rat. J. Cell. Comp.

Physiol, 21: 213-227.
Gersh, I., and Bodian, D. (1943a). Histochomical analysis of changes in Rhesus

motoneurons after root section. Biol. Symposia, 10: 163-184.
Gersh, I., and Bodian, D. (1943b). Some chemical mechanisms in chroma-

tolysis. J. Cell. Comp. Physiol, 21: 253-274.
Gersh, I., and Stieglitz, E. J. (1933). A critical study of histochemical methods

for the determination of iodides in tissues. Anat. Record, 56: 185-193.
Gibbs, G. E., and Kirk, P. L. (1934). Quantitative drop analysis. IV. The

determination of urea and ammonia. Mikrochemie, 16: 25-36.
Ginsel, L. A. (1939). The ultraviolet absorption of sheep thyroglobulin.

Biochem. J., 33: 428-434.
Giroud, A. (1929). Recherches sur la nature chimque du chondriome. Proto-

plasma, 7: 72-98.
Giroud, A., and Bulliard, H. (1933). Reaction des substances a fonction

sulfhydryle. Methode de mise en evidence dans les tissus. Protoplasma,

19: 381-384.
Giroud, A., and Leblond, C. P. (1936). L'acide ascorhique dans les tissus et

sa detection. Hermann, Paris.
Giroud, A., and Leblond, C. P. (1937). Histological study of renal elimination

of ascorbic acid. Anat. Record, 68: 113-126.
Glick, D. (1934). Studies on enzymatic histochemistry. VIII. A micro method

for the determination of lipolytic enzyme activity. Conipt. rend. trav.

lab. Carlsberg, Sir. chim., 20: No. 5, 6 pp.; Z. physiol. Chem., 223: 252-

256.
Glick, D. (1935a). Methods and applications of enzyme studies in histological

chemistry by the Linderstr0m-Lang Holter technic. J. Chem. Education.

12: 253-259.
Glick, D. (1935b). The chemical determination of minute quantities of vitamin

C. /. Biol Chem., 109: 433-436.
Glick, D. (1937). The quantitative distribution of ascorbic acid in the develop-
ing barley embryo. Compt. rend. trav. lab. Carlsberg, Sir. chim., 21: 203-

209; Z. physiol chem., 245: 211-216.
Glick, D. (1938). Studies on enzj'matic histochemistry. XXV. A micro method

for the determination of choline esterase and the activity-pH relationship

of this enzyme. Compt. rend. trav. lab. Carlsberg, Sir. chim., 21: 263-268;

J. Gen. Physiol, 21 : 289-295.
Glick, D. (1946). Histochemical localization of adenosinetriphosphatase. Sci-
ence, 103: 599.
Glick, D., and Biskind, G. R. (1935). The histochemistry of the adrenal gland.

II. The quantitative distribution of lipolytic enzymes. /. Biol. Chem., 110:

575-582.
Glick, D., and Fischer, E. E. (1945a). The histochemical localization of

adenosinetriphosphatase in plant and animal tissues. Science, 102 : 429-430.
Glick, D., and Fischer, E. E. (1945b). Studies in histochemistry. XVI. Methods

for the histochemical localization of adenosinetriphosphatase, thiamine-

BIBLIOGRAPHY 483

pyrophosphatase and glycerophosphatase in grains and sprouts. Arch.
Biochem., S: 91-96.

Click, D., and Fischer, E. E. (1946a). Studies in histochemistry. XVII. Locali-
zation of phosphatases in the wheat grain and in the epicotyl and roots
of the germinated grain. Arch. Biochem., 11: 65-79.

Glick, D., and Fischer, E. E. (1946b). Studies in histochemistry. XVIII. Locali-
zation of arginine in the wheat grain and in the epicotyl and roots of the
germinated grain. Arch. Biochem., 11: 81-87.

Gomori, G. (1939). Microtechnical demonstration of phosphatase in tissue
sections. Proc. Soc. Exptl. Biol. Med., 42 : 23-26.

Gomori, G. (1941a). The distribution of phosphatase in normal organs and
tissues. J. Cell. Coin-p. Physiol., 17: 71-83.

Gomori, G. (1941b). Distribution of acid phosphatase in the tissues under
normal and under pathologic conditions. Arch. Path., 32: 189-199.

Gomori, G. (1942). Histochemical reactions for lipid aldehydes and ketones.
Proc. Soc. Exptl. Biol. Med., 51 : 133-134.

Gomori, G. (1943). Calcification and phosphatase. Am. J. Path., 19: 197-209.

Gomori, G. (1945a). Recent advances in histochemistry. J. Mt. Sinai Hosp.
N. Y., 11: 317-26.

Gomori, G. (1945b). Microtechnical demonstration of sites of lipase activity.
Proc. Soc. Exptl. Biol. Med. 58: 362-364.

Gomori, G. (1946a). Distribution of lipase in the tissues under normal and
under pathologic conditions. Arch. Path., 41: 121-129.

Gomori, G. (1946b). A new histochemical test for glycogen and mucin. Am.
J. Clin. Path., Tech. Sect., 10: 177-179.

Gomori, G. (1946c). The study of enzymes in tissue sections. Am. J. Clin. Path.,
16: 347-352.

Goodspeed, T. H. and LTaer, F. M. (1934). Application of the Altmann freez-
ing-drying techniques to plant cytology. Proc. Natl. Acad. Sci. U.S., 20;
495-501.

Goodspeed, T. H., and Uber, F. M. (1935). Application of the Altmann freez-
ing-drying technique to plant cytology. II. Character of the fixation. Univ.
Calij.', Berkeley, Pubs. Botany, 18: 23-32.

Gorbach, G. (1936). Die Mikrowaage. Mikrochemie, 20: 254-336.

Gorbman, A., and Evans, H. L. (1941). Correlation of histological differentia-
tion with beginning of function of developing thyroid gland of frog. Proc.
Soc. Exptl. Biol. Med., 47: 103-106.

Goulhart, M. (1939). Sur una forme cristalline qui precede les cristaux de
Teichmann dans la recherche de I'hemoglobine. Compt. rend. soc. bioL,
132: lOS-111.

GouUiart, M. (1941). La detection histochimique de I'hemoglobine dans les
tissues, sous la forme de microcristaux de protoiodoheme. Compt. rend. soc.
biol, 135: 1260-1262.

Graffi, A. (1939). Zellulare Speicherung cancerogener Kohlenwasserstoffe. Z.
Krebsforsch., 49: 477-495.

Graffi, A. (1940). Intracellulare Benzpyrenspeichrung in lebenden Normal-
und Tumorzellen. Z. Krebsforsch., 50: 196-219.

484 BIBLIOGRAPHY

Grafflin, A. L. (1942). Histological observations upon the porphyrin-excreting
Harderian gland of the albino rat. Am. J. Anat., 71: 43-64.

Granick, S. (1938). Quantitative isolation of chloroplasts from higher plants.
Am. J. Botany, 25: 558-561.

Groat, R. A. (1939). Two new mounting media superior to Canada balsam
and gum damar. Anat. Record, 74: 1-6.

Gunther, W. H. (1941). trber den histologischen Nachweis des Benzpyrens.
Z. Krebsforsch., 52: 57-60.

Gunther, P., and Wilcke, G. (1926). Beitrage zur Rontgenspektralanalyse, II.
tJber die Verwendung der Methode der Silberkornziihlung zur Photo-
metrierung von Rontgenspektrallinien. Z. physikal. Chem., 119: 219.

Hackmann, C. (1942). The determination of sulfonamides in histological sec-
tions. Med. u. Chem. Abhandl. med. chem. Forschungsstdtten I. G. Far-

beyiind., 4: 134-137.
Hadfield, I. H. (1942). Two simple micro-burettes and an accurate wash-out

pipette. J. Soc. Chem. Ind. London, 61: 45-50.
Haitinger, M. (1938). Fluoreszenz-Mikroskopie. Akadem. Verlagsgesellschaft,

Leipzig.
Hale, C. W. (1946). Histochemical demonstration of acid polysaccharides in

animal tissues. Nature, 157: 802.
Hamilton, J. G. (1941). Applications of radioactive tracers to biology and

medicine. J. Applied Phys., 12: 440^60.
Hamilton, J. G., Soley, M. H., and Eichorn, K. B. (1940). Deposition of

radioactive iodine in human thyroid tissue. Univ. Calif., Berkeley, Pubs.

Pharmacol., 1 : 339-367.
Hammett, F. S., and Chapman, S. S. (1938-39). The quantitative unrehability

of the nitroprusside test for — SH and — S— S— . J. Lab. Clin. Med., 24:

293-298.
Hamos, L. von, and Engstrom, A. (1944). Microanalysis by secondary roentgen

spectrography. Acta Radiol., 25: 325-331.
Hand, W. C, Edwards, B. B., and Caley, E. R. (1943). Studies in the pharma-
cology of mercury. III. Histochemical demonstration and differentiation of

metallic Hg, mercurous Hg, and mercuric Hg. J. Lab. Clin. Med., 28: 1835-

1841.
Harrison, B. F., Thomas, M. D., and Hill, G. R. (1944). Radioautographs

showing the distribution of sulfur in wheat. Plant Physiol., 19: 245-257.
Harvey, E. B. (1932). The development of half and quarter eggs of Arbacia

punctidata and of strongly centrifuged whole eggs. Biol. Bull., 62: 155-167
Harvey, E. B. (1936). Parthenogenetic merogony or cleavage without nuclei in

Arbacia punctulata. Biol. Bull., 71: 101-121.
Harvey, E. N. (1931). The tension at the surface of marine eggs, especially

those of the sea-urchin, Arbacia. Biol. Bull., 61 : 273-279.
Harvey, E. N. (1932). The microscope-centrifuge and some of its applications.

J. Franklin Inst., 214: 1-23.
Harvey, E. N. (1933). A new form of centrifuge-microscope for simultaneous

observation of control and experimental material. Science, 77: 430--431.

BIBLIOGRAPHY 485

Hawes, R. C, and Skavinski, E. R. (1942). Diffusion micromethod for nitrogen.
Ind. Eng. Chem., Anal. Ed., 14: 917-921.

Heatley, N. G. (1935a). A new type of microburette. Biochem. J., 29: 626-630.

Heatley, N. G. (1935b). The distribution of glycogen in the region.s of the
amphibian gastrula; with a method for the micro-determination of glyco-
gen. Biochem. J., 29: 25G8-2572.

Heatley, N. G. (1939). An easily made microburette. Mikrochemie, 26: 147-149.

Heatley, N. G. (1940). An improved model of a new type of microrespirometer.
J. Sci. Instruments, 17: 197-202.

Heatley, N. G., Berenblum, I., and Chain, E. (1939). A new type of micro-
respirometer. Biochem. J., 33: 53-67.

Heatley, N. G., and Lindahl, P. E. (1937). Studies on the nature of the am-
phibian organization centre. V. The distribution and nature of glycogen
in the amphibian embryo. Proc. Roy. Soc. London, B122: 395-402.

Heck, K., Brown, W. H., and Kirk, P. L. (1937). Quantitative drop analysis.
IX. Determination of glucose. Mikrochemie, 22: 306-314.

Helander, S. (1945a). Detection of chemotherapeutics in thin sections of tissue
by the aid of fluorescence microscopy. Nature, 155 : 109.

Helander, S. (1945b). On the concentrations of some sulfanilamide derivatives
in different organs and tissue structures. Acta Physiol. Scand., 10: Suppl.
XXIX, 1-103.

Hempelmann, L. H., Jr. (1940). Staining reactions of the mucoproteins. Anat.
Record, 78: 197-206.

Henle, W., Henle, G., and Chambers, L. A. (1938). Studies on the antigenic
structure of some mammalian spermatoza. J. Exptl. Med., 68: 335-352.

Henriot, E., and Huguenard, E. (1925). Sur la realisation de tres grandes
vitesses de rotation. Comyt. rend., 180: 1389-1392.

Hepler, 0. E., Simonds, J. P., and Gurley, H. (1940). Renal phosphatase in ex-
perimental nephropathies. Proc. Soc. Exptl. Biol. Med., 44: 221-223.

Hevesy, G. (1940). Application of radioactive indicators in biology. Ann. Rev.
Biochem., 9: 641-662.

Hevesy, G. (1948). Radioactive Indicators, Their Application in Biochemistry,
Animal Physiology, and Pathology. Interscience, New York, 1948.

Hill, R., and Scarisbrick, R. (1940). The reduction of ferric oxalate by isolated
chloroplasts. Proc. Roy. Soc. London, B129: 238-255.

Hirt, A., and Wimmer, K. (1939a). Luminescenzmikroskopische Beobachtung
iiber das Verhalten von Vitaminen im lebenden Organismus. Das Vitamin
B. in der Leber. Klin. Wochschr., 18: 733-740.

Hirt, A., and Wimmer, K. (1939b). Luminescenzmikroskopische Beobachtungen
iiber das Verhalten von Nicotinsaure und Nicotinsiiureamid im lebenden
Organismus. Klin. Wochschr., 18: 765-767.

Hirt, A., and Wimmer, K. (1940). Luminescenzmikroskopische Untersuchungen
am lebenden Tier. Klin. Wochschr., 19: 123-128.

Hoerr, N. L. (1936). Cytological studies by the Altmann-Gersh freezing-drying
method. I. Recent advances in the technique. Anat. Records, 65: 293.

Hoerr, N. L. (1943). Methods of isolation of morphological constituents of the
liver cell. Biol. Symposia, 10: 185-231.

486 BIBLIOGRAPHY

HofiT, N. L., and Scott, G. H. (1944). Frozen-dehydration method for histologic

fixation, in Glasser, D., Medical Physics. Year Book Publishers, Chicago,

pp. 466-468.
Hollande, A. C. (1931). L'insolubilisation des urates figures dans les coupes

histologiques. Bull, histol. appl. physiol. et path, et tech. microscop., 8:

176-178.
Holmgren, H., and Wilander, O. (1937). Beitrag zur Kenntnis der Chemie und

Funktion der Ehrlichschen Mastzellen. Z. mikroskop. anat. Forsch., 42:

242-278.
Holter, H. (1943). Technique of the Cartesian diver. Compt. rend. trav. lab.

Carlsberg, Sir. chim., 24: 399-478.
Holter, H. (1945). A colorimetric method for measuring the volume of large

amoebae. Compt. rend. trav. lab. Carlsberg, Sir. chim., 25: 156-167.
Holter, H., and Doyle, W. L. (1938). Studies on enzymatic histochemistry.

XXVin. Enzymatic studies on protozoa. J. Cell. Comp. Physiol., 12:

295-308.
Holter, H., and Linderstr0m-Lang, K. (1932). Beitrage zur enzymatischen

Histochemie. III. tJber die Proteinasen von Drosera rotundijolia. Z. phys-
iol. Chem., 214: 223-240.
Holter, H., and Linderstr0m-Lang, K. (1936). Enzymverteilung im Proto-

plasma. Monatsh., 69: 292-313.
Hotchkiss, R. D. (1946). Personal covimunication.
Huggins, C, and Talalaj^ P. (1945). Sodium phenolphthalein phosphate as a

substrate for phosphatase tests. J. Biol. Chon., 159 : 399^10.
Humphrey, A. A. (1935). Dinitrosoresorcinol — A new specific stain for iron in

tissues. Arch. Path., 20: 256-258.
Huntson, R. D. (1935). An inexpensive direct-current amplifier. Rev. Sci. Instru-
ments, 6: 322-323.

Imhauser, K. (1927). tJber das Vorkommen des Plasmalogens. II. tJber das
Vorkommen des Plasmologens bei Tieren. Biochem. Z., 186: 360-375.

Isaacs, M. L. (1922). A colorimetric determination of blood chlorides. J. Biol.
Chem., 53: 17-19.

Jackson, C. (1944). An improved method of staining lipides: acetic-carbol-
Sudan. Onderstcpoort J. Vet. Sci. Animal Ind., 19: 169-177.

Jacobsen, C. F. (1942). The contraction accompanying the cleavage of peptide
linkages in synthetic substrates for trypsin and chymotr3'psin. Compt. rend.
trav. lab. Carlsberg, Sir. chim., 24: 281-287.

Jacobsen, C. F., and Linderstrom-Lang, K. (1940). Method for rapid determina-
tion of specific gravity. Acta Physiol. Scand., 2 : 149-152.

Jandorf, B. J., Klemperer, F. W., and Hastings, A. B. (1941). A manometric
method for the determination of diphosphopyridine nucleotide. J. Biol.
Chem., 138: 311-320.

Jenkins, R. (1937). Microscopy with fluorescent light. Stain. Technol., 12:
167-173.

Johansen, D. A. (1940). Plant Microtechnique. McGraw-Hill, New York.

BIBLIOGRAPHY 487

Johnson, H. C, and Kirk, P. L. (1940). Quantitative drop analysis. XII. Deter-
mination of the ultrafilterable calcium of blood. Mikrochemie ver. Mikro-
chim. Acta, 28: 254-257.

Johot, F., and Curie, I. (1934). Artificial production of a new kind of radio-
element. Nature, 133: 201-202.

Kabat, E. A., and Furth, J. (1941). A histochemical study of the distribution of
alkaline phosphatase in various normal and neoplastic tissues. Am. J. Path.,
17: 303-318.

Kalmus, H. (1928). Die Messungen der Atmung. Garung und CO2- Assimilation
kleiner Organismen in der Kapillare. Z. vergleich. Physiol., 7: 304-313.

Kaufmann, H. P. (1926). Weitere Versuche zur Bromometrie der Fette. Z.
Untersuch. Lebensm., 51: 3-14.

Kay, W. W., and Whitehead, R. (1935). The staining of fat with Sudan IV
J. Path. Bact., 41: 303-304.

Keilin, D., and Hartree, E. F. (1938). Cytochrome oxidase. Proc. Roy. Sac.
London, B125: 171-186.

King, E. J. (1932). The colorimetric determination of phosphorus. Biochem. J.,
26: 292-297.

King, E. J., and Delory, G. E. (1939). The rates of enzymic hydrolysis of phos-
phoric esters. Biochem.. J., 33: 1185-1190.

Kinsey, V. E., and Robison, P. (1946). Micromethod for the determination of
urea. J. Biol. Chem., 162: 325-331.

Kirk, P. L. (1933). Quantitative drop analysis. I. General apparatus and tech-
nique. Mikrochem,ie, 14: 1-14.

Kirk, P. L. (1935). Preparation of sintered glass filters. Ind. Eng. Chem., Anal.
Ed., 7: 135-136.

Kirk, P. L., and Bentley, G. T. (1936). Quantitative drop analysis. V. Determi-
nation of iron in simple and biological media. Mikrochemie, 21 : 250-259.

Kirk, P. L., and Craig, R. (1932). Note on a method for the manipulation of
blood-dilution and micro-pipettes. J. Lab. Clin. Med., 18: 81-83.

Kirk, P. L., and Tompkins, P. C. (1941). Micro- and drop-scale titration of
oxalate. Ind. Eng. Chem., Anal. Ed., 13: 277-280.

Klemperer, P. (1947). Some recent biologic investigations and their significance
for pathology. J. Mount Sinai Hosp., 14: 442-448.

Kochakian, C. D., and Fox, R. P. (1944). Microdetermination of calcium by
titration of the oxalate with ammonium hexanitratocerate. Ind. Eng. Chem,.,
Anal. Ed., 16: 762-763.

Kolthoff, I. M., and Lingane, J. J. (1941). Polarography (Revised Reprint,
1946). Interscience, New York.

Komaya, G. (1925). tJber eine histochemische Nachweismethode der Resorp-
tion, Verteilung und Asscheidung des Wismutes in den Organen. Arch.
Dermatol. Syphilol, 149: 277-291.

Kortiim, G. (1934). Uber einen neuen rotierenden Sektor fiir Lichschwach-
ungen grosser Genauigkeit. Z. Instrumentenk ., 54: 373-377.

Kossa, J. von (1901). Ueber die im Organismus kiinstlich erzeugbaren Verkal-
kungen. Beitr. path. Anat. u. allgem. Path., 29: 163-202.

Kretchmer, N., Holman, R. T., and Burr, G. 0. (1946). A submicro method for
the determination of iodine number of lipids. Arch. Biochem., 10: 101-105.

488 BIBLIOGRAPHY

Krogh, A. (1935). Syringe pipets. Ind. Eng. Chem., Anal. Ed., 7: 130-131.

Krogh, A., and Keys, A. B. (1931). A syringe-pipette for precise analytical
usage. J. Chem. Soc, 2436-2440.

Krugelis, E. J. (1946). Distribution and properties of intracellular alkaline
phosphatases. Biol. Bull, 90: 220-233.

Ku, D. Y. (1933). Microincineration studies on human coronary arteries. Am.
J. Path., 9: 23-36.

Kunitz, M. (1940). CrystalHne ribonuclease. /. Gen. Physiol., 24: 15-32.

Kurbatov, J. D., and Pool, M. L. (1943). Radioactive isotopes for the study of
trace elements in living organisms. Chem. Revs., 32: 231-248.

Kuttner, T., and Cohen, H. R. (1927). Micro colorimetric studies. I. A molybdic
acid, stannous chloride reagent. The micro estimation of phosphate and
calcium in pus, plasma, and spinal fluid. J. Biol. Chem., 75: 517-531.

Kuttner, T., and Lichtenstein, L. (1930). Micro colorimetric studies. II. Estima-
tion of phosphorus. Molybdic acid-stannous chloride reagent. J. Biol.
C/iem., 86: 671-676.

Lacassagne, A., and Lattes, J. S. (1924). Methods auto-histo-radiographique

pour la detection dans les organes du polonium injecte. Compt. rend., 178:

488-490.
Laidlaw, G. F. (1932). The dopa reaction in normal histology. Anat. Record, 53:

399-413.
Laidlaw, G. F., and Blackberg, S. N. (1932). Melanoma studies. II. A simplified

technique for the dopa reaction. Am. J. Path., 8: 491-498.
Laitinen, H. A., and Kolthoff, I. M. (1941a). Voltammetry with stationary

microelectrodes of platinum wire. J. Phys. Chem., 45: 1061-1079.
Laitinen, H. A., and Kolthoff, I. M. (1941b). Voltammetric determinations and

amperometric titrations with a rotating microelectrode of platinum wire.

J. Phys. Chem., 45: 1079-1093.
Landow, H., Kabat, E. A., and Newman, W. (1942). Distribution of alkaline

phosphatase in normal and in neoplastic tissues of the nervous system.

Arch. Neurol. Psychiat., 48: 518-530.
Laskowski, M. (1942). Preparation of living nuclei from hen erythrocytes.

Proc. Soc. Exptl. Biol. Med., 49: 354-356.
Lavin, G. I. (1943). SimpHfied ultraviolet microscopy. Rev. Sci. Instruments,

14: 375-376.
Lawrence, E. 0., and Cooksey, D. (1936). On the apparatus for the multiple

acceleration of light ions to high speeds. Phys. Rev., 50: 1131-1140.
Lawrence, E. 0., and Livingston, M. S. (1932). The production of high speed

light ions without the use of high voltages. Phys. Rev., 40: 19-35.
Lazarow, A. (1941). The oxygen uptake of mitochondria and other cell frag-
ments. J. Biol. Chem., 140: Proc. XXV-XXVI.
Lazarow, A. (1942). Particulate glycogen: A submicroscopic component of the

guinea pig liver cell; its significance in glycogen storage and the regulation

of blood sugar. Anat. Record, 84: 31-50.
Lazarow, A. (1943). The chemical structure of cytoplasm as investigated in

Bensley's laboratory during the past ten years. Biol. Symposia, 10: 9-26.

, BIBLIOGRAPHY 489

Lee, Bolles (1937). The Microtomists Vade-Mecum. 10th Ed., Blakiston, Phil-
adelphia.

Leger, M. E. (1888). Sur una reaction caracteristique du bismuth. Bull. soc.
chim. biol, 50: 91-93.

Levvy, G. A. (1946). The cerimetric determination of glucuronic acid, using the
Conway burette. Biocheni. J., 40: 396-400.

Levy, M. (1936). Studies on enzymatic histochemistry. XVII. A micro Kjeldahl
estimation. Compt. rend. trav. Carlsberg, Sir. chim., 21: 101-110; Z.
physiol. chem., 240: 33-42.

Levy, M., and Palmer, A. H. (1940). lodometric estimation of small quantities
of nitrogen without distillation. J. Biol. Chem., 136: 57-60.

Levy, M., and Palmer, A. H. (1943). Chemistry of the chick embryo. IV. Ami-
nopeptidase. J. Biol. Chem., 150: 271-279.

Lillie, R. D. (1941). Stain fading in various histologic mounting media. Stain
Technol, 16: 127.

Linderstr0m-Lang, K. (1936). Studies in enzymatic histochemistry. XIX.
Microestimation of alkalies in tissue. Compt. rend. trav. lab. Carlsberg, Ser.
chim., 21: 111-122.

Linderstr0m-Lang, K. (1937a). Dilatometric ultra-microestimation of peptidase
activity. Nature, 139: 713.

Linderstr0m-Lang, K. (1937b). Principle of the Cartesian diver applied to
gasometric technique. Nature, 140: 108.

Linderstr0m-Lang, K. (1939). Distribution of enzymes in tissue and cells.
Harvey Lectures, Ser. 34: 214-245.

Linderstr0m-Lang, K. (1942). Approximate solution of certain diffusion prob-
lems in liquid sj^stems. Compt. rend. trav. lab. Carlsberg, Ser. chim., 24:
249-280.

Linderstr0m-Lang, K. (1943). On the theory of the Cartesian diver micro respi-
rometer. Compt. rend. trav. lab. Carlsberg, Ser. chim., 24: 333-398.

Linderstr0m-Lang, K., and Duspiva, F. (1936). Studies on enzymatic histo-
chemistry. XVI. The digestion of keratin by the larvae of the clothes
moth. (Tineola biselliella Humm). Compt. rend. trav. lab. Carlsberg, Ser.
chim., 21: 53-84; Z. physiol. Chem., 237: 131-158 (1935).

Linderstr0m-Lang, K., and Engel, C. (1937). Uber die Verteilung der Amylase
in den aussern Schicten des Gerstenkornes. Enzymologia, 3: 138-146;
Compt. rend. trav. lab. Carlsberg, Ser. chim., 21: 243-258 (1938).

Linderstr0m-Lang, K., and Glick, D. (1938). Micromethod for determination of
choline esterase activity. Compt. rend. trav. lab. Carlsberg, Ser. chim., 22:
300-306.

Linderstr0m-Lang, K., and Holter, H. (1931). Contributions to the histological
chemistry of enzymes. I. The estimation of small cleavages caused by en-
zymes. Compt. rend. trav. lab. Carlsberg, Ser. chim. 19, No. 4: 1-19; Z.
physiol. Chem., 201 : 9-30.

Linderstr0m-Lang, K., and Holter, H. (1932). Contribution to the histochem-
istry of enzymes. II. The distribution of peptidase in the roots and sprouts
of malt. Compt. rend. trav. lab. Carlsberg, Ser. chim., 19, No. 6: 1-39; Z.
physiol. Chem., 204 : 15-33.

490 BIBLIOGRAPHY

Linderstr0m-Lang, K., and Holier, H. (1933a). Studies on enzymatic histo-
chemistry. V. A micro method for the estimation of sugars. Compt. rend.
trav. lab. Carlsberg, Ser. chim., 19, No. 14: 1-12.

Linderstr0m-Lang, K., and Holter, H. (1933b). Studies in enzymatic histo-
chemistry. VI. A micro-method for the estimation of ammonia. Compt.
rend. trav. lab. Carlsberg, Ser. chim., 20: 1-9; Z. physiol. chem., 220:
5-12.

Linderstr0m-Lang, K., and Holter, H. (1940). Die enzymatische Histochemie,
in Bamann, E., and Myrback, K., Die Methoden der Ferment j or schung .
Thieme, Leipzig, pp. 1132-1162.

Linderstr0m-Lang, K., and Holter, H. (1942). Diffusion of bases through pro-
tective seals of oil or flotation medium in the Cartesian diver micro respi-
rometer. Compt. rend. trav. lab. Carlsberg, Ser. chim., 24: 105-138.

Linderstr0m-Lang, K., Holter, H., and S0eborg-Ohlsen, A. (1934). Studies on
enzymatic histochemistry. XTII. The distribution of enzymes in the
stomach of pigs as a function of its histological structure. Z. physiol. Chem.,
227: 1-50; Compt. rend. trav. lab. Carlsberg, Ser. Chim., 20: 66-127 (1935).

Linderstr0m-Lang, K., Jacobsen, O., and Johansen, G. (1938). On the measure-
ment of the deuterium content of mixtures of H2O and DaO. Compt. rend,
trav. lab. Carlsberg, Ser. chim., 23: 17-25.

Linderstr0m-Lang, K., and Lanz, H., Jr. (1938). Studies on enzymatic histo-
chemistry. XXIX. Dilatometric micro-estimation of peptidase activity.
Compt. rend. trav. lab. Carlsberg, Ser. chim., 21: 315-338; Mikrochim.
Acta, 3: 210-230.

Linderstr0m-Lang, K., and Mogensen, K. R. (1938). Studies on enzymatic
histochemistry. XXXI. Histological control of histochemical investigations.
Compt. rend. trav. lab. Carlsberg, Ser. chim., 23: 27-35.

Linderstr0m-Lang, K., Palmer, A. H., and Holter, H. (1935). Studies on enzy-
matic histochemistry. XIV. A micro-determination of chloride in tissues.
Compt. rend. trav. lab. Carlsberg, Ser. chim., 21: 1-5; Z. physiol. Chem.,
231 : 226-230.

Linderstr0m-Lang, K., and S0eberg-Ohlsen, A. (1936). Studies on enzymatic
histochemistry. XX. Distribution of urease in dog's stomach. Emymologia,
1: 92-95.

Linderstr0m-Lang, K., Weil, L., and Holter, H. (1935). Studies on enzymatic
histochemistry. XV. A micro-estimation of arginase. Compt. rend. trav. lab.
Carlsberg, Ser. chim., 21: 7-13; Z. physiol. Chem. 233: 174-180.

Lindner, R., and Kirk, P. L. (1937a). Quantitative drop analysis. VII. Further
investigations on the determination of calcium. Mikrochemie, 22: 291-300.

Lindner, R., and Kirk, P. L. (1937b). Quantitative drop analysis. VIII. Deter-
mination of phosphorus. Mikrochemie, 22: 300-305.

Lindner, R., and Kirk, P. L. (1938). Quantitative drop analysis. X. Determina-
tion of sodium. Mikrochemie, 23: 269-279.

Lindsay, E., and Craig, R. (1942). The distribution of radiophosphorus in wax
moth, mealworm, cockroach and firebat. Ann. Entomol. Soc. Am., 35:
50-56.

- BIBLIOGRAPHY 491

Links, R. (1934). Hahnlose Mikroburette mit Sperrfliisigkeit fiir Serienbestim-
mungen. Microchemie, 15: 87-94.

Lison, L. (1935). fitudes sur la metachroraasie colorants metachromatiques et
substances chromotropes. Arch. biol. Liege, 46: 599-668.

Lison, L. (1936). Histochemie animale. Gauthier-Villars, Paris.

Lison, L. (1938). Ziir Frage der Ausscheidung und Speicherung des Hamo-
globins in der Amphibienniere Beitr. path. Anat. u. allgem. Path., 101:
94-108.

Loscalzo, A. G., and Benedetti-Pichler, A. A. (1945). Titration of microgram
samples. Ind. Eng. Chem., Anal. Ed., 17: 187-191.

Lowry, O. H. (1941). A quartz fiber balance. J. Biol. Chem., 140: 183-189.

Lowry, O. H. (1943). Electrolytes in the cytoplasm, Biol. Symposia, 10:
233-245.

Lowry, O. H. (1944). A simple quartz torsion balance. /. Biol. Chem., 152:
293-294.

Lowry, O. H., and Bessey, 0. A. (1944). A method for the determination of
five-tenths to two millimicrograms of riboflavin. /. Biol. Chem., 155: 71-77.

Lowry, 0. H., and Bessey, 0. A. (1946). The adaptation of the Beckman spec-
trophotometer to measurements on minute quantities of biological mate-
rials. J. Biol. Chem., 163: 633-639.

Lowry, O. H., and Hunter, T. H. (1945). The determination of serum protein
concentration with a gradient tube. J. Biol. Chem., 159: 465-474.

Lowry, O. H., and Lopez, J. A. (1946). The determination of inorganic phos-
phate in the presence of labile phosphate esters. J. Biol. Chem., 162:
421-428.

Lowry, 0. H., Lopez, J. A., and Bessey, 0. A. (1945). The determination of
ascorbic acid in small amounts of blood serum. J. Biol. Chem., 160: 609-615.

Lundsteen, E., and Vermehren, E. (1936). Micro methods for the estimation of
phosphatases in blood plasma and inorganic phosphorus in blood. Compt.
rend. trav. lab. Carlsberg, Ser. chim., 21: 147-166; Enzymologia, 1: 273-279.

McBain, J. W. (1939). Opaque or analytical ultracentrifuges. Chem. Revs., 24:

289-302.
McClung, C. E. (1937). Handbook oj Microscopical Technique. 2nd ed.,

Hoeber, New York.
MacHattie, L. E. (1941). The production of high rotational speed. Rev. Set.

Instruments, 12: 429-435.
McJunkin, F. A. (1922). Peroxj'dase staining with benzidin in paraffin sections

of human tissue. Anat. Record, 24: 67-77.
MacKee, G. M., Herrmann, F., Baer, R. L., and Sulzberger, M. B. (1943a).

Demonstrating the presence of sulfonamides in the tissues. Science, 98:

66-68.
MacKee, G. M., Herrmann, F., Baer, R. L., and Sulzberger, M. B. (1943b).

A histochemical method for demonstrating the presence of sulfonamides in

the tissues. J. Lab. Clin. Med., 28: 1642-1649.
McLean, F. C., and Bloom, W. (1940). Calcification and ossification. Calcifica-
tion in normal growing bone. Anat. Record, 78: 333-359.

492 BIBLIOGRAPHY

McMillen, J. H., and Scott, G. H. (1937). A magnetic electron microscope of

simple design. Rev. Sci. Instruments, 8: 288-290.
Macallum, A. B. (1908). Die Mothodcn und Ergebnisse der Mikrochemie in

der biologischen Forschung. Ergeb. Physiol., 7: 552-652.
Mallory, F. B., and Parker, F., Jr. (1939). Fixing and staining methods for lead

and copper in tissues. Am. J. Path., 15: 517-522.
Maj^er, D. T., and Gulick, A. (1942). The nature of the proteins of cellular

nuclei. /. Biol. Chem., 146: 433-440.
Mendel, L. B., and Bradley, H. C. (1905). Experimental studies on the physiol-
ogy of the molluscs. Am. J. Physiol., 14: 313-327.
Menke, W. (1937). Untersuchungen der einzelnen Zellorgan in Spinatblattern

auf Grund preparativchemisches Methodik. Z. Botan., 32: 273-295.
Menke, W. (1988). Untersuchungen liber das Protoplasma griiner Pflanzen-

zellen. I. Isolierung von chloroplasten aus Spinatblattern. Z. physiol.

Chem., 257: 43-48.
Menten, M. L., Junge, J., and Green, M. H. (1944). A coupling histochemical

azo dye test for alkaline phosphates in the kidney. J. Biol. Chem., 153:

471^77.
Metcalf, R. L. (1943). The storage and interaction of water soluble vitamins in

the Malpighian sj^stem of Periplaneta americana (L). Arch. Biochem., 2:

55-62.
Metcalf, R. L., and Patton, R. L. (1942). A study of riboflavin metabolism in

the American roach by fluoresence microscopy. /. Cell. Comp. Physiol.,

19: 1^.
Metcalf, R. L., and Patton, R. L. (1944). Fluorescence microscopy applied to

entomology and allied fields. Stain Technol., 19: 11-27.
Michaelis, L., and Granick, S. (1945). Metachromasy of basic dyestuffs. J. Am.

Chem. Soc, 67: 1212-1219.
Miescher, F. (1871). Ueber die chemische Zusammensetzung der Eiterzellen, in

Hoppe-Seyler, F., Medizinisch-chemische Untersuchungen, Berlin, 4: 441;

and in Jones, W. (1920), Niicleic Acids, 2nd ed., Longmans, Green, London

and New York, p. 2.
Milovidov, P. (1928). Sur les methodes de double coloration du chondriome

et des grains d'amidon. Arch. anat. microsp., 24: 9-18.
Milovidov, P. (1938). Bibliographie der Nucleal- und Plasmalreaktion. Proto-
plasma, 31: 246-266.
Mirsky, A. E. (1943). Chromosomes and nucleoproteins, in Nord, F. F., and

Werkman, C. H., eds., Advances in Enzymology, Vol. III. Interscience,

New York, pp. 1-34.
Mitchell, A. J., and Wislocki, G. B. (1944). Selective staining of glycogen by

ammoniacal AgNOa: a new method. Anat. Record, 90: 261-266.
Mommaerts, W. F. H. M. (1938). Some chemical properties of the plastid-

granum. Proc. Koninkl. Nederland, Akad. Wetenschap., 41: 896-903.
Montgomery, C. G., and Montgomery, D. D. (1938). Grid controlled ioniza-
tion gauge. Rev. Sci. Instruments, 9: 58.
Moog, F. (1943a). The use of manganese in the histochemical demonstration of

acid phosphatase. J. Cell. Comp. Physiol., 22: 95-97.

BIBLIOGRAPHY 493

Moog, F. (1943b). Cytochrome oxidase in early chick embryos. J. Cell. Comp.

Physiol, 22: 223-231.
Moog, F. (1944). Localizations of alkaline and acid phosphatases in the early

embiyogenesis of the chick. Biol. Bull., 86: 51-80.
Moog, F., and Steinbach, H. B. (1946). Notes on the possibility of a histo-

chemical method for localizing adenosinetriphosphatase. Science, 103: 144.
Moreau, P. (1931). Essais de localisation histochimique du calcium dans les

organes pen riches en cet ion par la microincineration combinee a des re-
actions chimiques. Bull, histol. appl. physiol. et path, et tech. microscop.,

8: 245-248.
Morris, D. L. (1946). Colorimetric determination of glycogen. Disadvantages

of the iodine method. J. Biol. Chem., 166: 199-203.
Mullen, J. P. (1944). A convenient and rapid method for staining glycogen in

paraffin sections with Best's carmine stain. Ajn. J. Clin. Path., 14: 9-10.
Miiller, E. (1935). Die Phosphatasebestimmung in kleinen Serummengen.

Z. physiol. Chem., 237: 35-39.
Murray, J. A. (1937). Technique for paraffin sections of formal fixed insect

material. J. Roy. Microscop. Soc, 57: 15.

Needham, J., and Boell, E. J. (1939). An ultramicro-Kjeldahl technique.

Biochem. J., 33: 149-152.
Neish, A. C. (1939). Studies on chloroplasts. I. Separation of chloroplasts. a

study of factors affecting their flocculation and the calculation of the

chloroplast content of leaf tissue from chemical analysis. Biochem. J.,

33: 293-299.
Nimmo-Smith, R. H. (1946). A note on the cerimetric determination of blood

glucose. Biochem,. J., 40: 414.
Norberg, B. (1937). Beitrage zur Enzymatischen Histochemie. XXII. Mikro-

bestimmung von kalium. Compt. rend. trav. lab. Carlsberg, Ser. chim.,

21: 233-241; Mikrochim. Acta, 1: 212-219.
Norberg, B. (1942). Histo- and cytochemical determination of phosphorus.

Ada Physiol. Scand., 5, Suppl. 14: 99 pp.

O'Brian, H. C, and Hance, R. T. (1940). A plastic cover glass, isobutyl

methacrylate. Science, 91: 412.
Ochoa, S., and Peters, R. A. (1938). Vitamin Bi and cocarboxylase in animal

tissues. Biochem. J., 32 : 1501-1515.
Ohmori, Y. (1937). Phosphomonoesterase. Enzymologia, 4: 217-231.
Okamoto, K., Akagi, T., and Mikami, G. (1939). Biologische Untersuchungen

des Goldes. I. Uber die histochemische Goldnachweismethode. Acta Schol.

Med. Univ. Imp. Kioto, 22: 373-381.
Okamoto, K., Mikami, G., and Nishida, M. (1939). Biologishe Untersuchungen

des Goldes. I. tJber die histochemische Goldnachweismethode. Acta Schol.

Med. Univ. Imp. Kioto, 22: 382-387.
Okamoto, K., and Utamura, M. (1938). Biologische Untersuchungen des

Kupfers. I. Histochemische Kupfernachweismethode. Acta Schol. Med.

Univ. Imp. Kioto, 20: 573,

494 BIBLIOGEAPHY

Okamoto, K., Utamura, M., and Akagi, T. (1939). Biologische Untersuchungen

des Silbers. I. Histochemische Silbernachweismethode. Acta Schol. Med.

Univ. Imp. Kioto, 22: 361-372.
Okkels, H. (1927). Disposition de la chaux dans les reins dan Tintoxication

experimentale par le calcium., Bull, histol. appl. physiol. path, et tech.

microscop., 4: 134-141.
Opie, E. L., and Lavin, G. I. (1946). Localization of ribonucleic acid in the

cytoplasm of liver cells. J. Exptl. Med., 84: 107-112.
Oster, K. A., and Mulinos, M. G. (1944). Tissue aldehydes and their reaction

with amines. /. Pharmacol. Exptl. Therap., 80: 132-138.
Oster, K. A., and Oster, J. G. (1946). The specificity of sex hormones on the

tissue aldehyde shift in the rat kidney and of fuchsin sulfurous acid

reagent on aldehydes. J. Pharmacol. Exptl. Therap., 87: 306-312.
Oster, K. A., and Schlossman, N. C. (1942). Histochemical demonstration of

amine oxidase in the kidney. J. Cell. Camp. Physiol., 20: 373-378.

Packer, D. M., and Scott, G. H. (1942). A crycstat of new design for low

temperature tissue dehydration. J. Tech. Meth. Bull. Intern. Assoc. Med.

Museums, 22: 85-96.
Pap, T. (1929). Eine neue Methode zur Impragnation des Retikulums. Zent.

allgem. Path. u. path. Anat., 47: 116-117.
Pecher, C. (1941). Biological investigations with radioactive calcium and

strontium. Proc. Soc. Exptl. Biol. Med., 46: 86-91.
Peters, J. P., and Van Slyke, D. D. (1932). Quantitative Clinical Chemistry.

Williams & Wilkins, Baltimore.
Pickels, E. G. (1942). An improved type of electrically driv^en high speed

laboratory'' centrifuge. Rev. Sci. Instruments, 13: 93-100.
Pickford, G. E. (1937). A sealed-in micro glass electrode. Proc. Soc. Exptl.

Biol. Med., Z%: 154-157.
Pijoan, M., and Gerjovich, H. J. (1946). The use of 2,4-dinitrophenylhydrazine

for the determination of ascorbic acid. Science, 103: 202-203.
Pincussen, L. (1927). Methodische Mitteilungen. VII, 3. Mikrotitrationsapparat.

Biochem. Z., 186: 32-35.
Policard, A. (1931-32). Some new methods in histochemistry. Harvey Lectures,

Ser. 27: 204-226.
Policard, A. (1933a). L'histospectrographie. Protoplasma, 19: 602-629.
Policard, A. (1933b). Mineral constituents of blood vessels as determined by

the technique of microincineration, in Cowdry, E. V., Arteriosclerosis.

Macmillan, New York, pp. 121-130.
Policard, A., and Martin, E. (1933). Recherches d'histochimie quantitative sur

la silico-tuberculose et la tuberculose pulmonaires. Bull, histol. appl.

physiol. et path, et tech. m,icroscop., 10: 22-36.
Policard, A., and Morel, A. (1932). Application de la spectrographie d'emission

aux problemes histochimiques. L'histospectrographie par etineelage direct

des coupes. Bidl. histol. appl. physiol. et path, et tech. microscop., 9: 57-68.
Policard, A., and Okkels, H. (1930). Localizing inorganic substances in micro-
scopic sections. The microincineration method. Anat. Record, 44: 349-361.
Policard, A., and Okkels, H. (1931). Die Mikroveraschung (Mikrospodographie)

BIBLIOGRAPHY 495

als histochemische Hilfsmethode, in Abderhalden, E., Handbuch der

biologischen Arbeitsmethoden, Abt. V, Urban & Schwarzenberg, Berlin,

pp. 1815-1828.
Policard, A., and Pillet, D. (1926). Recherches sur certaines reserves calcaires

revelees par microincineration au niveau de la region cephalique chez les

larves de Batraciens. Bidl. histol. appl. et 'path, et tech. mikroscop., 3:

230-235.
Policard, A., and Ravant, P. (1927). Procede perrmettant la microincinerati

sans vetraction d'organes riches en tissu fibreux. Bull, histol. appl. physiol.

et path, et tech. microscop., 4: 170.
Pollack, H. (1928). Micrurgical studies in cell physiology. VI. Calcium ions in

living protoplasm. J. Gen. Physiol., 11: 539-545.
Pouting, J. D. (1943). Extraction of ascorbic acid from plant materials.

Relative suitabiUty of various acids. Ind. Eng. Cheni., Anal. Ed., 15:

389-391.

Pool, M. L., and Kurbatov, J. D. (1943). Principal Artificial Radioactive Iso-
topes as Trace Elements. (A chart of data compiled from nuclear trans-
mutation literature.) Form 4605, Nuclear Physics Laboratory, Ohio State
University, Columbus, Ohio.

Popper, H. (1941). Histologic distribution of vitamin A in human organs under
normal and under pathologic conditions. Arch. Path., 31 : 766-802.

Popper, H. (1944). Distribution of vitamin A in tissue as visualized by
fluorescence microscopy. Physiol. Revs., 24: 205-224.

Popper, H., and Elsasser, M. (1941). The photographic visualization of vitamin
A in tissue by means of fluorescence microscopy. Can. J. Med. Technol.,
3: 45-50.

Post, E. E., and Laudermilk, J. D. (1942). A new microchemical reaction for
cellulose. Stain. Technol., 17: 21-26.

Pringsheim, P., and Vogel, M. (1946). Luminescence of Liquids and Solids.
Rev. Ed., Interscience, New York.

Querner, F. R. von (1935). Der mikroskopische Nachweis von Vitamin A in
animalen Gewebe. III. Zur Ivenntnis der paraplastischen Leberzellein-
schliisse. Klin. Wochschr., 14; 1213-1217.

Radley, J. A., and Grant, J. (1939). Fluorescence Analysis in Ultraviolet Light.

3rd ed.. Van Nostrand, New York.
Rafalko, J. S. (1946). A modified Feulgen technique for small and diffuse

chromatin elements. Stain Technol., 21: 91-93.
Ralph, P. H. (1941). The histochemical demonstration of hemoglobin in blood

cells and tissue smears. Stain Technol., 16: 105-106.
Ramsay, W. N. M. (1944). The micro-volumetric determination of iron, with

particular reference to blood. Biochem. J., 38: 467-469.
Raoul, Y., and Meunier, P. (1939). Sur I'essai des preparations a base de

vitamine A. J. pharm. chim., 29: 112-118.
Rappaport, F. (1935). Mikrochemie des Blutes. Haim, Vienna, p. 86.
Rask-Nielsen, R. (1944). Studies on enzymatic histochemistry. XXXVII. On

496 BIBLIOGRAPHY

the peptidase content of the pyloric glandular cells. Compt. rend. trav. lab.

Carlsberg, Sir. chim., 25: 1-32.
Richards, A. N., Bordley, J. Ill, and Walker, A. M. (1933). Quantitative studies

of the composition of glomerular urine. VII. Manipulative technique of

capillary tube colorimetry. J. Biol. Chem., 101: 179-191.
Roberts, W. J. (1935). Recherche de I'or dans Ics tissus animaux par la

methode du developpement physique. "Keimmethode" de Lieseganz, Bull.

histol. appl. physiol. et path, et tech. microscop., 12: 344-361.
Rocher, H. (1942). Micromanometre base sur le principe du ludion. Compt.

rend. soc. bioL, 136: 366-367.
Rocher, H. (1943). Micromanometre base sur le principe du ludion. Bull. soc.

chim. 10: 486-492.
Roe, J., and Kuether, C. A. (1943). The determination of ascorbic acid in

whole blood and urine through the 2,4-dinitrophenylhydrazine derivative

of dehydroascorbic acid. J. Biol. Chem., 147: 399^07.
Romieu, M. (1925). Sur la detection histochimique des substances proteiques.

Bull, histol. appl. physiol. et path, et tech. microscop., 2: 185-191.
Romieu, M. (1927). Une reaction histochimique nouvelle des lecithines, la

reaction iodophile. Compt. rend., 184: 1206-1208.
Roughton, F. J. W., and Scholander, P. F. (1943). Micro gasometric estimation

of the blood gases. I. Ox>'gen. J. Biol. Chem., 148: 541-550.
Russell, J. A. (1944). The colorimetric estimation of small amounts of ammonia

by the phenol-hypochlorite reaction. J. Biol. Chem., 156: 457-461.

Sakaguchi, S. (1925). Uber eine neue Farbenreaktion von Protein und Arginin.

J. Biochem. Japan, 5: 25-31.
Salomon, K., Gabrio, B. W., and Smith, G. F. (1946). A precision method for

the quantitative determination of calcium in blood plasma. Arch. Biochem.,

11: 433-443.
Sawj'er, C. H. (1943). Cholinesterase and the behavior problem in Amblystoma.

I. The relationship between the development of the enzyme and early

motility. /. Exptl. Zool, 92: 1-29.
Schmidt-Nielsen, K. (1942). Microtitration of fat in quantities of 10"^ gram.

Compt. rend. trav. lab. Carlsberg, Ser. chim., 24: 233-247.
Schmidt-Nielsen, K. (1944a). Micro determination of the iodine number of fat

in quantities of 10^ gram. Compt. rend. trav. lab. Carlsberg, Ser. chim.;

25: 87-96.
Schmidt-Nielsen, K. (1944b). Extraction and fractionation of the lipids in 1 mg.

of tissue. Compt. rend. trav. lab. Carlsberg, Ser. chim., 25: 97-104.
Schneider, G. (1903). Ein Beitrag zur Physiologie der Niere niederer Wirbel-

tiere. Skand. Arch. Physiol, 14: 383-389.
Scholander, P. F. (1942). Microburette. Science, 95: 177-178.
Scholander, P. F. (1942a). Volumetric microrespirometers. Rev. Sci. Instru-
ments, 13: 32-33.
Scholander, P. F. (1942b). A micro-gas-analyzer. Rev. Sci. Instruments, 13:

264-266.
Scholander, P. F. (1942c). Microgasometric determination of nitrogen in blood

and saliva. Rev. Sci. Instruments, 13: 362-364.

BIBLIOGRAPHY 497

Scholander, P. F., and Edwards, G. A. (1942). Volumetric microrespirometer

for aquatic organisms. Rev. Sci. Instruments, 13: 292-295.
Scholander, P. F., Edwards, G. A., and Irving, L. (1943). Improved micrometer

burette. J. Biol. Chem., 14S: 495-500.
Scholander, P. F., and Roughton, F. J. W. (1942). A simple micro-gasometric

method of estimating carbon monoxide in blood. J. Ind. Hyg. Toxicol.,

24: 218-221.
Scholander, P. F., and Roughton, F. J. W. (1943a). Micro gasometric estima-
tion of the blood gases. II. Carbon monoxide. J. Biol. Chem., 148: 551-563.
Scholander, P. F., and Roughton, F. J. W. (1943b). Micro gasometric estimation

of the blood gases. IV. Carbon dioxide. J. Biol. Chem., 148: 573-580.
Schultz, A. (1924). Eine Methode des mikrochemischen Cholesterinnachweises

am Gewebsschnitt. Zentr. allgem. Path. u. path. Anat., 35: 314-317.
Schultz, A. (1925). tJber Cholesterinesterverfettung. Verhandl. deut. path. Ges.,

20: 120-123.
Schultz, A., and Lohr, G. (1925). Zur Frage der Spezifitat der mikrochemischen

Cholesterin-Reaktion mit Eisessig-Schwefelsaure. Zentr. allgem. Path. u.

path. Anat., 36: 529-533.
Schultz, A. S., Atkin, L., and Frey, C. N. (1942). Determination of vitamin B,

by yeast fermentation method. Ind. Eng. Chem., Anal. Ed., 14: 35-39.
Schultz-Brauns, O. (1929). Histo-topochemische Untersuchungen an krankhaft

veranderten Organen unter Anwendung der Schnittveraschung. Arch. path.

Anat. Physiol. Virchow's, 273: 1-50.
Schultz-Brauns, O. (1931). Die Methode der Schnittveraschung unfixierter

tierischer Gewebe. Z. wiss. Mikroskop., 48: 161-191.
Scott, G. H. (1932). Topographic similarities between materials revealed by

ultraviolet light photomicrography of living cells and by microincineration.

Science, 76:148-150.
Scott, G. H. (1933a). A critical study and review of the method of microin-
cineration. Protoplasma, 20: 133-151.
Scott, G. H. (1933b). Quantitative estimation of ash after microincineration.

Proc. Sac. Exptl. Biol. Med., 30: 1304-1305.
Scott, G. H. (1937). The microincineration method of demonstrating mineral

elements in tissues, in McClung, C. E., Microscopical Technique. 2nd ed.,

Hoeber, New York, pp. 643-665.
Scott, G. H. (1943). Mineral distribution in the cytoplasm. Biol. Symposia;

10: 277-289.
Scott, G. H., and Packer, D. M. (1939a). The electron microscope as an

analytical tool for the localization of minerals in biological tissues. Anat.

Record, 74: 17-29.
Scott, G. H., and Packer, D. M. (1939b). An electron microscope study of

magnesium and calcium in striated muscle. Anat. Record, 74: 31-45.
Scott, G. H., and Williams, P. S. (1934). A new development in histospectrog-

raphy. Proc. Soc. Exptl. Biol. Med., 32: 505-507.
Scott, G. H., and Williams, P. S. (1935). The spectrographic analysis of

biological materials. Anat. Record, 64: 107-127.
Scott, G. H., and Williams, P. S. (1936). A simplified cryostat for the dehydra-
tion of frozen tissues. Anat. Record, 66: 475-^81

498 BIBLIOGRAPHY

Selzer, L., and Baiimberger, J. P. (1942). The influence of metallic mercury on

the respiration of cells. J. Cell. Comp. Physiol., 19: 281-287.
Semenoff, W. E. (1935). Mikrochemische Bestimmung der Aktivitat der

Succinodehydrase in den Organen der Rana temporaria. Z. Zelljorsch.

viikroskop. Anat., 22: 305-309.
Sen, P. B. (1930). A method of locating urease within tissue by a micro-
chemical method. Indian J. Med. Research, 18: 79-82.
Sendroy, J., Jr. (1939a). Note on tests for purity of solid silver iodate prepared

for chloride determination. J. Biol. Chem., 127: 483-485.
Sendroy, J., Jr. (1939b). Photoelectric microdetermination of chloride in bio-
logical fluids, and of iodate and iodine in protein-free solutions. J. Biol.

Chem., 130: 605-623.
Sendroy, J., Jr. (1942a). Note on the photoelectric microdetermination of

chloride in biological fluids. J. Biol. Chem., 142: 171-173.
Sendroy, J., Jr. (1942b). Photoelectric determination of oxalic acid and calcium,

and its application to micro- and ultramicroanalysis of serum. J. Biol.

Chem., 144: 243-258.
Sendroy, J., Jr. (1944). Determination of serum calcium by precipitation with

oxalate. A comparative study of factors affecting the results of several

procedures. J. Biol. Chem., 152: 539-556.
Sendroy, J., Jr., and Alving, A. S. (1942). Photoelectric microdetermination of

iodate and iodine. /. Biol. Chem., 142: 159-170.
Serra, J. A. (1944a). Eine neue histochemische Reaktion: die Reaktion des

Arginins. Naturwissenschajten, 32: 46; Z. wiss. Mikroscop., 60.
Serra, J. A. (1944b). Improvements in the histochemical arginine reaction and

the interpretation of this reaction. Porlugaliae Ada Biol., 1 : 1-7.
Serra, J. A. (1946). Histochemical tests for proteins and amino acids; the

cl'iaracterization of basic proteins. Stain Technol., 21 : 5-18.
Serra, J. A., and Queiroz Lopes, A. (1944). Direkter Nachweis von basischen

Proteinen im Zellkern. Naturwissenschaften, 32: 47; Chromosoma, 2.
Serra, J. A., and Queiroz Lopes, A. (1945). Une methode pour la demonstration

histochimique du phosphore des acides nucleiques. Portugaliae Acta Biol.,

1: 111-122.
Sevag, M. G., Smolens, J., and Stern, K. G. (1941). Isolation and properties of

pigmented heavy particles from Streptococcus pyogenes. J. Biol. Chem.,

139: 925-941.
Sharlet, H., Sachs, W., Sims, C. F., and Salzer, B. H. (1942). Histochemical

observations on melanin production in the skin. Arch. Dermatol. Syphilol.,

45: 103-111.
Simpson, W. L. (1941). An experimental analysis of the Altman technic of

freezing-drying. Anat. Record, 80: 173-189.
Simpson, W. L. (1943). Radioactive isotopes, in Cowdry, E. V., Microscopic

Technique In Biology and Medicine, Williams & Wilkins, Baltimore,

pp. 165-170.
Simpson, W. L., and Cramer, W. (1943). Fluorescence studies of carcinogens

in skin. I. Histological localization of 20-methylcholanthrene in mouse

skin after a single application. Cancer Research, 3: 362-369.

BIBLIOGRAPHY 499

Simpson, W. L., and Cramer, W. (1945). Fluorescence studies of carcinogens

in skin. II. ISIouse skin after single and multiple applications of 20-methyl-

cholanthrene. Cancer Research, 5: 449-463.
Sinnott, E. W. (1944). Science and the education of free men. Am. Scientist,

32: 205-215.
Sisco, R. C, Cunningham, B., and Kirk, P. L. (1941). Quantitative drop

analysis. XIII. The formal titration of amino nitrogen. J . Biol. Chem.,

139: 1-10.
Siwe, S. A. (1935a). Stufenphotometrische Bestimmung von Phosphor in

kleinen Blutmengen. Biochem. Z., 278: 437^41.
Siwe, S. A. (1935b). Einige Methoden zur Bestimmung von Ca in kleinen

Blutmengen. Biochem. Z., 278: 442-446.
Sjostrand, F. (1945/1946). trber die Eigenfluorescenz tierischer Gewebe mit

besonderer Beriicksichtigung der Saiigetierniere. Acta Anat., 1, Suppl. 1;

163 pp.
Sjostrand, F. (1946a). Cytological localization of riboflavin and thiamine by

fluorescence microspectrography. Nature, 157: 698.
Sjostrand, F. (1946b). The fluorescence microspectrographic localization of

riboflavin (vitamin B2) and thiamin (vitamin Bi) in tissue cells. Acta

Physiol. Scand., 12 : 42-52.
Skarstrom, C, and Beams, J. W. (1940). The electrically driven magnetically

supported vacuum type ultracentrifuge. Rev. Set. Instruments, 11: 398-403.
Snell, E. E., and Strong, F. M. (1939). A microbiological assay for riboflavin.

Ind. kng. Chem., Anal. Ed., 11: 346-350.
Sobel, A. E., and Kaye, I. A. (1940). Estimation of submicroquantities of

calcium. Ind. Eng. Chem., Anal. Ed., 12: llS-120.
S0eberg-Ohlsen, A. (1941). Contribution to the histochemistry of the stomach.

Compt. rend. trav. lab. Carlsberg, Sir. chim., 223: 329-482.
Sreenivasaya, M., and Bhagvat, K. (1937). Dilatometiy and its applications in

the study of enzymes. Ergeb. Enzymforsch., 6: 234-243.
Stadie, W. C, O'Brien, H., and Lang, E. P. (1931). Determination of the pH

of serum at 38° with the glass electrode and an improved electron tube

potentiometer. J. Biol. Chem., 91 : 243-269.
Stehle, R. L. (1921). The gasometric determination of urea in urine. J. Biol.

Chem., 4:7: 13-17.
Steiger, A. (1941). Mikrochemischer Nachweis des Karotins. Microkosmos, 34:

121-122.
Stein, J. (1935). Reaction histochimique stable de detection de la bilirubine.

Compt. rend. soc. bioL, 120: 1136-1138.
Stern, K. G. (1932). Uber die Katalase farbloser Blutzellen. Z. physiol. Chem.,

204: 259-282.
Stern, K. G. (1942). A simple ultracentrifuge with plastic rotor. Science, 95:

561-562.
Stern, K. G. (1943). Studies on macromolecular particles endowed with specific

biological activity. Biol. Symposia, 10: 291-321.
Stern, K. G., and Wyckoff, R. W. G. (1938a). An ultracentrifugal study of

catalase. Science, 87: 18.

500 BIBLIOGRAI'HY

Stern, K. G., and Wyckoff, R. W. G. (1938b). An ultracentrifugal study of

catalase. J. Biol. Chem., 124: 573-583.
Stone, I. (1940). Stable 2,6-dichlorobenzenoneindophenol solutions. Ind. Eng.

Chem., Anal. Ed., 12: 415.
Stoneburg, C. A. (1939). Lipids of the cell nuclei. J. Biol. Chem., 129: 189-196.
Stowell, R. E. (1942). Photometric histochemical determination of thymonucleic

acid in experimental epidermal carcinogenesis. J. Natl. Cancer Inst., 3:

111-121.
Stowell, R. E. (1945a). Feulgen reaction for thymonucleic acid. Stain Technol.,

20: 45-58.
Stowell, R. E. (1945b). The photometric histochemical determination of sub-
stances in the skin. J. Investigative Dermatol., 6: 183-189.
Stowell, R. E. (1946). The specificity of the Feulgen reaction for thymonucleic

acid. Stain Technol, 21: 137-148.
Stowell, R. E., and Albers, V. M. (1943). A spectrophotometric analysis of

tissue staining. Stain Technol., 18: 57-71.
Sumner, J. B. (1925). A more specific reagent for the determination of sugar in

urine. J. Biol. Chem., 65: 393-395.
Sumner, J. B. (1944). A method for the colorimetric determination of phos-
phorus. Science, 100: 413-414.
Sure, B., and Wilder, V. M. (1941). Simplified photoelectric procedures for

microdetermination of nonprotein nitrogen, urea nitrogen, creatine, and

creatinine of blood. J. Lab. Clin. Med., 26: 874-878.
Sutro, C. J. (1936). Fluorescent microscopy. Arch. Path., 22: 109-1 xb.
Svedberg, T., and Pedersen, K. O. (1940). The Utracentrifuge. Clarendon

Press, London.
Sylven, B. (1941). tjber das Vorkommen von hochmolekularen Esterschwelfel-

sauren im Granulationsgewebe und bei der Epithelregeneration. Acta Chir.

Scand., 86: Suppl. 66, 151 pp.
Sylven, B. (1945). Ester sulphuric acids of high molecular weight and mast

cells in mesenchymal tumors. Histochemical studies on tumorous growth.

Acta Radiol., Suppl. 59: 100 pp.

Tada, K. (1926). Ueber eine histochemische Nachweismethode von Blei.
Verhandl. japan, path. Ges., 16: 128.

Takamatsu, H. (1939). Histologische und biochemische Studien iiber die
Phosphatase. Histochemische Untersuchungsmethodik der Phosphatase und
deren Verteilung in verschiedenen Organen und Geweben. Trans. Soc. Path.
Japan. 29: 429-498.

Tannenholz, H., and Muir, K. B. (1933). Methods for microchemical demon-
stration of arsenic in tissues. Arch. Path., 15: 789-795.

Telford Govan, A. D. (1944). Fat-staining by Sudan dyes suspended in watery
media. J. Path. Bact., 56: 262-264.

Thomas, J. A., and Lavollay, J. (1935). reaction histochimique du fer a
la 8-hydroxyquinoleine. Bull, histol. appl. physiol. et path, et tech. micro-
scop., 12: 400-402.

Thomas, L. E. (1946). A histochemical test for arginine-rich proteins. J. Cell.
Cornp. Physiol, 28: 145-157.

BIBLIOGRAPHY 501

Tobias, J. M. (1942). Membrane interferometer manometer. Rev. Sci. Instru-
ments, 13: 232-233.
Tobias, J. M. (1943). Microrespiration techniques. Physiol. Revs., 23: 51-75.
Tobias, J., and Gerard, R. W. (1941). An improved capillary microrespirometer.

Proc. Soc. Exptl. Biol. Med., 47: 531-533.
Tompkins, E. R., and Kirk, P. L. (1942). Quantitative drop analysis. XVI.

An improved diffusion method for total nitrogen. J. Biol. Chem., 142:

477-485.
Tonutti, E. (1938). Ergebnisse histochemischer Vitamin C Untersuchungen.

Protoplasma, 31 : 151-158.
Treadwell, A. de G., Low-Beer, B. V. A., Friedell, J. L., and Lawrence, J. H.

(1942). Metabolic studies on neoplasm of bone with the aid of radioactive

strontium. Am. J. Med. Sci., 204: 521-530.
Trevan, J. W. (1925). The micrometer syringe. Biochem. J., 19: 1111-1114.
Tschopp, E. (1929). Die Lokalisation anorganischer Substanzen in Geweben

(Spodographie), in Mollendorff, W. von, Handbuch der mikroskopischen

Anatomic des Menschen, Springer, Berlin, 1 : 569-600.
Tswett, M. (1911). Uber Reicherts Fluoreszenz Mikroskop und einigen damit

angestellten Beobachtungen iiber Chlorophyll und Cyanophyll. Ber. deut.

botan. Ges., 29: 744-746.
Turchini, J., Castel, P., and Kien, K. V. (1943). De quelques colorations

nucleaires par des fluorones et leur rapprochement avec la reaction

nucleale de Feulgen pour la distinction des acides thymo et zymonucleique,

Arch. soc. sci. med. biol. Montpellier et Lanquedoc, 23-24: 599-600.
Turchini, J., Castel, P., and Kien, K. V. (1944). Contribution a I'etude

histochimique de la cellule. Une nouvelle reaction nucleale. Bull, histol.

appl. physiol. et path, et tech. microscop., 21: 124—127.
Turchini, J., and GosseHn de Beaumont, L. A. (1945). AppUcation d'une

reaction nucleale nouvelle a I'etude de divers noyaux de cellules vegetales.

Com.pt. rend. soc. biol., 139: 584—585.
Tyler, A., and Berg, W. E. (1941). A new type of micro-respirometer. Science,

94: 397-398.

Umbreit, W. W., Burris, R. H., and Stauffer, J. F. (1945). Manometric
Techniques and Related Methods jar the Study of Tissue Metabolism.
Burgess, Minneapolis.

Van Slj^ke, D. D., and Cullen, G. E. (1914). A permanent preparation of urease

and its use in the determination of urea. /. Biol. Chem., 19: 211-228.
Van Slyke, D. D., and Hiller, A. (1933). Determination of ammonia in blood.

J. Biol. Chem., 102 : 499-504.
Van Wagtendonk, W. J., Simonsen, D. H., and Hackett, P. L. (1946). A rapid

microdetermination of glycogen in tissue slices. J. Biol. Chem., 163:

301-306.
Verne, J. (1928). Demonstration histochimique de la formation de corps a

fonctions aldehydiques aux depens des enclaves graisseuses et lipidiques,

Compt. rend. soc. biol., 99: 266-269.

502 BIBLIOGRAPHY

Volk, B. W., and Popper, H. (1944a). Factors influencing the persistence of
vitamin A fluorescence in tissue sections. Arch. Path., 38: 90-92.

Volk, B. W., and Popper, H. (1944b). Microscopic demonstration of fat in
urine and stool by means of fluorescence microscopy. Am. J. Clin. Path.,
14: 234-238.

Wachstein, M., and Zak, F. G. (1946). Bismuth pigmentation. Its histo-
chemical identification. Am. J. Path., 22: 603-611.

Waldschmidt-Leitz, E. (1925). Zur Kenntnis der Enterokinase. Z. physiol.
Cheyn., 142: 217-244.

Walker, A. M. (1933). Quantitative studies of the composition of glomerular
urine. X. The concentration of inorganic phosphate in glomerular urine
from frogs and Necturi determined by an ultramicromodification of the
Bell-Doisy method. J. Biol. Chem., 101: 239-254.

Walker, A. M., and Hudson, C. L. (1937). The role of the tubule in the
excretion of urea by the amphibian kidney. With an improved technique
for the ultramicro-determination of urea nitrogen. Am,. J. Physiol., 118:
153-166.

Walker, A. M., and Reisinger, J. A. (1933). Quantitative studies of the com-
position of glomerular urine. IX. The concentration of reducing sub-
stances in glomerular urine from frogs and Necturi determined by an
ultramicroadaptation of the method of Sumner. J. Biol. Chem., 101:
223-237.

Warburg, 0. (1910). tJber Beeinflussung der Sauerstoffatmung. Z. physiol.
Chem., 70: 413^32.

Waterhouse, D. F. (1945). Studies of the physiology and toxicology of blow-
flies. A histochemical examination of the distribution of copper in
Lucillia cwprina. Council for Scientific and Industrial Research, Common-
wealth of Australia, Melbourne, Bull. No. 191, 20 pp.

Weil, L. (1936). Investigations in enzymatic histochemistrj^. II. A micro-method
for the determination of tryptic activity. Biochem. J., 30: 5-10.

Weil, L., and Russell, M. A. (1940). Studies on plasma phosphatase activity in
relation to fat metabolism in rats. J. Biol. Chem., 136: 9-23.

Westenbrink, H. G. K. (1940). Determination of cocarboxylase and aneurin.
A micro-modification of Ochoa and Peter's method. Com,pt. rend. trav.
lab. Carlsberg, Ser. chim., 23: 195-211; Enzymologia, 8: 97-107.

Westfall, B. B., Findley, T., and Richards, A. N. (1934). Quantitative studies
of the composition of glomerular urine. XII. The concentration of chloride
in glomerular urine of frogs and Necturi. J. Biol. Chem., 107: 661-672.

Whitaker, T. W. (1939). The use of the Feulgen technic with certain plant
materials. Stain Tcchnol., 14: 13-16.

Widmark, E. M. P., and Orskov, S. L. (1928). Eine neue Mikrobiirette.
Biochem. Z., 201: 15-21.

Wigglesworth, V. B. (1937). A simple method of volumetric analysis for small
quantities of fluid: Estimation of chloride in 0.3 /il. of tissue fluid.
Biochem. J., 31 : 1719-1722.

Williams, P. S., and Scott, G. H. (1935a). An electrode arrangement for spark
spectrography. Rev. Sci. Instruments, 6: 277-278, 361.

- BIBLIOGRAPHY 503

Williams, P. S., and Scott, G. H. (1935b). Apparatus for darkfield photometry

and densitometry. J. Optical. Soc. Am., 25: 347-349.
Williamson, M. B., and Gulick, A. (1944). The calcium and magnesium content

of mammalian cell nuclei. /. Cell. Comp. Physiol., 23: 77-82.
Willstatter, R., and Rohdewald, M. (1934). tJber den Zustand des Glykogena

in der Leber, im Muskel und in Leukocyten. Z. physiol. Chem., 225:

103-124.
Wilschke, A. (1914). Uber die Fluoreszenz der Chlorophyllkomponenten. Z.

wiss. Mikroskop., 31: 338-361.
Wolf, A., Kabat, E. A., and Newman, W. (1943). Histochemical studies on

tissue enzymes. III. A study of the distribution of acid phosphatases with

special reference to the nervous system. Am. J. Path., 19: 423-439.

Yamasaki, K. (1931). Ueber die Farbenreaktion auf Cholesterin und Oxycholes-
terin, sowie iiber die Methoden des mikrochemischen Cholesterin-Nach-
weises an Gewebsschnitten. Fukuoka Acta Med., 24: 79-81.

Yin, H. C. (1945). A histochemical study of the distribution of phosphatase in
plant tissues. New Phytologist, 44: 191-195.

Zemecnik, P. C., Lavin, G. I., and Bergmann, M. (1945). Microphotometric
determination of the rate of enzj-matic proteolysis. J. Biol. Chem., 158:
537-545.

Zeuthen, E. (1943). A Cartesian diver micro respirometer with a gas volume
of 0.1 fii. Compt. rend. trav. lab. Carlsberg, Sir. chim., 24: 479-518.

Zittle, C. A., and O'Dell, R. A. (1941). Chemical studies of bull spermatozoa.
Lipid, sulfur, cystine, nitrogen, phosphorus, and nucleic acid content of
whole spermatozoa and of the parts obtained by physical means. J . Biol.
Chem., 140: 899-907.

BIBLIOGRAPHY APPENDIX^

MICROSCOPIC TECHNIQUES

Chemical Methods:

1. Bartelmez, G. W., and Bensley, S. H. (1947). "Acid phosphatase" reactions

in peripheral nerves. Science, 106: 639-641.

2. Bondy, P. K., and Sheldon, W. H. (1947). Histochemical demonstration

of liver glycogen in human diabetic acidosis by liver biopsy. Proc. Soc.
Exptl. Biol. Med., 65 : 68-70.

3. Fawcett, D. W. (1947). Differences in physiological activity in brown and

white fat as revealed by histochemical reactions. Science, 105: 123.

4. Ferguson, R. L., and Silver, S. D. (1947). Demonstration in skin of active

chlorine from chlorine-liberating ointments. Am. J. Clin. Med., 17:
35-36.

5. Ferguson, R. L., and Silver, S. D. (1947). A method for the visual demon-

stration of lewisite in skin. Am. J. Clin. Path., 17: 37-38.

6. Gillman, T., and Ivy, A. C. (1947). A histological study of the participa-

tion of the intestinal epithelium, the reticulo endothelial system and
the lymphatics in iron absorption and transport. Gastroenterology, 9:
162-169.

7. Creep, R. 0., Fischer, C. J., and Morse, A. (1947). Histochemical demon-

stration of alkaline phosphatase in decalcified dental and osseous tissues.
Science, 105: 666.

8. Hess, M., and Hollander, F. (1947). Permanent metachromatic staining

of mucus in tissue sections and smears. /. Lab. Clin. Med., 32: 905-909.

9. Lassek, A. M. (1947). The stability of so-called axonal acid phosphatase

as determined by experiments on its "stainability." Stain Technol., 22:
133-138.

10. Leach, E. H. (1947). Bismarck brown as a stain for mucoproteins. Stain

Technol, 22: 73-76.

11. Lieb, E. (1947). Permanent stain for amyloid. Am. J. Clin. Path., 17:

413-414.

12. Lilhe, R. D., and Greco, J. (1947). Malt diastase and ptyalin in place of

saliva in the identification of glycogen. Stain Technol., 22: 67-70.
-13. McManus, J. F. A. (1946). Histological demonstration of mucin after
periodic acid. Nature, 158: 202.

14. Moree, R. (1947). A confirmation of Rafalko's Feulgen method. Stain

Technol, 22: 63-65.

15. Oster, K. A. (1947). Tissue aldehydes. Exptl Med. and Surg. 5: 219-235.

16. Richards, A. G. (1947). Studies on arthropod cuticle. I. The distribution

* Publications that appeared subsequent to completion of the manuscript.

504

filBLlOGRAPHY APPENDtX 505

Chemical Methods (continued) :

of chitin in lepidopterous scales and its bearing on the interpretation
of arthropod cuticle. Ann. Entomol. Sac. Am., 40: 227-240.

17. Silver, S. D., and Ferguson, R. L. (1947). A method for the visual demon-

stration of mustard (dichlorodiethyl sulfide) in skin. Am. J. Clin. Path.,
17: 39-43.

18. Stowell, R. E., and Zorzoli, A. (1947). The action of ribonuclease on fixed

tissues. Stain Technol., 22: 51-61.

19. Yin, H. C, and Sun, C. N. (1947). Histochemical method for the detec-

tion of phosphorylase in plant tissue. Science, 105: 650.

Physical Methods:

20. Axelrod, D. J., and Hamilton, J. G. (1947). Radio-autographic studies of

the distribution of lewisite and mustard gas in skin and eye tissues.
Am. J. Path., 23: 389-411.

21. Bayley, S. T. (1947). Autoradiography of single cells. Nature, 160: 193-195.

22. Endicott, K. M., and Yagoda, H. (1947). Microscopic historadiographic

technique for locating and quantitating radioactive elements in tissue.
Proc. Sac. Exptl. Biol. Med., 64: 170-172.

23. Engstrom, A. (1947). A new differential x-ray absorption method for

elementary chemical analysis. Rev. Sci. Instruments, 18: 681-682.

24. Engstrom, A. (1947). Quantitative cytochemical determination of nitrogen

by x-ray absorption spectrography. Biochim. et Biophys. Acta., 1: 428-
433.

25. Engstrom, A. (1947). Qualitative microchemical analysis by microradiog-

raphy with fluorescent screen. Experientia, 3: 208.

26. Engstrom, A., and Lindstrom, B. (1947). The photographic action of

x-rays of wavelengths 2.5-25 A. Experientia, 3: 1-3.

27. Engstrom, A., and Lindstrom, B. (1947). Histochemical analysis by x-rays

of long wavelengths. Experientia, 3: 191.

28. Evans, T. C. (1947). Radioautographs in which the tissue is mounted

directly on the photographic plate. Proc. Soc. Exptl. Biol. Med., 64:
313-315.

29. Henkes, H. E. (1947). An investigation into secondary thiochrome fluores-

cence in tissue sections. Acta Anat., 2: 321-350.

30. Marton, L., and Abelson, P. H. (1947). Tracer micrography. Science,

106: 69-70.

31. Symposia of the Society for Experimental Biology, No. I. Nucleic Acid.

Cambridge Univ. Press, London, 1947.

CHEMICAL TECHNIQUES

Colorimetric and Titrimetric Methods:

32. Bell, P. H., and Stryker, C. R. (1947). Thermostated cell compartment

for the Beckman spectrophotometer. Science, 105: 415-416.

33. Bessey, O. A., Lowry, O. H., and Brock, M. J. (1947). The quantitative

determination of ascorbic acid in small amounts of white blood cells
and platelets. J. Biol. Chem., 168: 197-205.

506 BIBLIOGRAPHY APPENDIX

Colorimetric and Tiinmeiric Methods (continued) :

34. Bolomey, R. A., and Kemmerer, A. R. (1947). Spectrophotometric studies

of the Roe method for the determination of dehydroascorbic acid. J.
Biol. Chem., 167: 781-785.

35. Engel, M. G., and Engel, F. L. (1947). The colorimetric microdetermina-

tion of urea nitrogen by the xanthydrol method. J. Biol. Chem., 167:
535-541.

36. Kalckar, H. M. (1947). Differential spectrophotometry of purine com-

pounds by means of specific enzymes. I. Determination of hydroxypurine
compounds. J. Biol. Chem., 167: 429-443.

37. Kalckar, H. M. (1947). Differential spectrophotometry of purine com-

pounds by means of specific enzymes. II. Determination of adenine
compounds. J. Biol. Chem., 167: 445-459.

38. Kalckar, H. M. (1947). Differential spectrophotometry of purine com-

pounds by means of specific enzymes. III. Studies of the enzymes of
purine metabolism. J. Biol. Chem., 167: 461-475.

39. Kirk, P. L., Craig, R., Gullberg, J. E., and Boyer, R. Q. (1947). Quartz

microgram balance, hid. Eng. Chem.., Anal. Ed., 19: 427-429.

40. Kirk, P. L., Rosenfels, R. S., and Hanahan, D. J. (1947). Capillary absorp-

tion cells in spectrophotometry. Ind. Eng. Chem., Anal. Ed., 19: 355-357.

41. Lazarow, A. (1947). Microprecipitating unit for blood and tissue proteins.

J. Lab. Clin. Med., 32: 213-214.

42. Lazarow, A. (1947). A simple apparatus for quantitative microcolorimetric

analysis in final volumes of 0.15 cc. /. Lab. Clin. Med., 32: 215-219.

43. Mills, M. B., and Roe, J. H. (1947). A critical study of proposed modifica-

tions of the Roe and Kuether method for the determination of ascorbic
acid, with further contributions to the chemistry of this procedure.
J. Biol. Chem., 170: 159-164.

44. Natelson, S., and Zuckerman, J. L. (1947). Burette for microtitration ;

Vernier applicable to the capillary micro burette. J. Biol. Chem., 170:
305-311.

45. Richter, K. M. (1947). A precision micro-pipette trimmer. Science, 106:

598-599.

Gasometric and Gradient Tube Methods:

46. Anfinsen, C. B., and Claff, C. L. (1947). An extension of the Cartesian

diver micro respirometer technique. J. Biol. Chem., 167: 27-33.

47. Claff, C. L. (1947). "Braking" pipettes. Science, 105: 103-104.

48. Grant, W. C. (1947). Determination of O2 capacity on 39.3 cubic milli-

meters of blood. Proc. Soc. Exptl. Biol. Med., 66: 60-62.

49. Levi, H., and Zeuthen, E. (1946). Micro-weighing in the gradient tube.

Compt. rend. trav. lab. Carlsberg, Sir. chim., 25: 273-287.

50. Scholander, P. F. (1947). Analyzer for accurate estimation of respiratory

gases in one-half cubic centimeter samples. /. Biol. Chem., 167: 235-250.

51. Scholander, P. F., and Evans, H. J. (1947). Microanalysis of fractions of

a cubic millimeter of gas. J. Biol. Chem., 169: 551-560.

52. Scholander, P. F., Flemister, S. C, and Irving, L. (1947). Microgasometric

BIBLIOGRAPHY APPENDIX 507

Gasometric and Gradient Tube Methods (continued):

estimation of the blood gases. V. Combined carbon dioxide and oxygen.
J. Biol. Chem., 169: 173-181.

53. Scholander, P. F., and Irving, L. (1947). Micro blood gas analysis in frac-

tions of a cubic millimeter of blood. /. Biol. Chem., 169: 561-569.

54. Zeuthen, E. (1947), A sensitive "Cartesian Diver" balance. Nature, 159:

440-441.

MECHANICAL SEPARATION OF CELLULAR COMPONENTS

55. Hogeboom, G. H., Schneider, W. C, and Pallade, G. E. (1947). The isola-

tion of morphologically intact mitochondria from rat liver. Proc. Soc.
Exptl. Biol. Med., 65: 320-321.

LIST OF MAISVFACTVRERS

Agfa-Ansco Corp., Binghamton, New York

A. S. Aloe Co., St. Louis, Missouri

S. Ash, 3044 Third Ave., New York, N.Y.

Askania Werke, BerUn, Germany

Atlas Powder Co., Wilmington, Delaware

British Drug Houses Ltd., London, England

Corning Glass Works, Corning, New York

Difco Laboratories, Inc., Detroit, Michigan

Distillation Products Co., Rochester, New York

Eastman Kodak Co., Rochester, New York

W. Edwards and Co., London, England

Eimer and Amend, New York, N.Y.

Eli Lilly and Co., Indianapolis, Indiana

Fisher Scientific Co., Pittsburgh, Pennsylvania

J. B. Ford Co., Wyandotte, Michigan

General Biochemicals Inc. (Smaco), Chagrin Falls, Ohio

General Electric Co., Schenectady, New York

J. D. Graham, Department of Physiology, University of Penn-
sylvania Medical School, Philadelphia, Pennsylvania

Emil Greiner Co., New York, N.Y.

Gulf Oil Co., Pittsburgh, Pennsylvania

Hanovia Chemical and Alanufacturing Co., Newark, New
Jersey

Hartmann-Leddon Co., Philadelphia, Pennsylvania

0. Hebel, Edward Martin Biological Laboratory, Swarthmore
College, Swarthmore, Pennsylvania

Hengar Co., Philadelphia, Pennsylvania

Hoffmann-La Roche Inc., Nutley, New Jersey

K. Hollborn & Sons, Leipzig, Germany

International Equipment Co., Boston, Massachusetts

Lusteroid Container Co., Inc., South Orange, New Jersey

Macahster Bicknell Co., Cambridge, Massachusetts

Merck and Co., Inc., Rahway, New Jersey

Microchemical Specialties Co., Berkeley, California

National Aniline Division, AUied Chemical and Dye Corp.,
40 Rector Street, New York, N.Y.

Newark Wire Cloth Co., Newark, New Jersey

Onyx Oil and Chemical Co., Jersey City, New Jersey

E. Peterson, Carlsberg Laboratory, Copenhagen, Denmark

Pfaltz and Bauer, Inc., New York, N.Y.

Pyrocell Mariufacturing Co., 207 East 84 Street, New York, N.Y.

508

LIST 'OF MANUFACTURERS 509

Radio Corporation of America, 30 Rockefeller Plaza, New York, N. Y.

Roller Smith Co., Bethlehem, Pennsylvania

G. Schonander Co., Stockholm, Sweden

G. Frederick Smith Chemical Co., Columbus, Ohio

E. R. Squibb and Sons, New York, N.Y.

A. H. Thomas Co., Philadelphia, Pennsylvania

Unicam Instruments Ltd., Cambridge, England

Western Electric Co., 195 Broadway, New York, N.Y,

Westinghouse Electric Corp., 40 Wall St., New York, N.Y.

SUBJECT INDEX

Acetic-carbol-Sudan III, for localiza-
tion of liquids, 39

Acetone-alcohol titration method for
determination of arginase, 308

Acid groups, by titrimetric techniques.
290

Acidimetric acetone method for pro-
teolytic enzymes, 303

Acidimetric method, for calcium, 274
for cholinesterase, 310
for esterase and lipase, 309

Acid phosphatase, chemical staining
methods for, 80
role of ascorbic acid in detection, 81
role of manganese ions in detection,
81

Acid polysaccharides, chemical stain-
ing methods, 45

"Air pipette," for filling microliter
divers under anaerobic conditions,
373

DL-AIanylglycine, substrate for dila-
tometric measurement of pepti-
dase, 418

Albert and Leblond 2,4-dinitrophenyl-
hydrazine reaction for water-
insoluble aldehydes and ketones,
71

Aldehydes, chemical staining methods
for, 65
"true" and "pseudo" Feulgen re-
actions of, differentiation, 65

Aldolase. See Zymohexase.

Alkali groups, by electrometric titra-
tions, 290
by titrimetric techniques, 290

Alkalimetric alcohol method for pro-
teol.ytic enzymes, 304

Alkaline phosphatase, chemical stain-
ing methods for, 78
effect of decalcification upon, 78

Allantoin, by Borsook method, 243
colorimetric determination by cu-
vette technique, 239
use of urease in determination, 243

Allen-Bourne method for zj^mohexase,
87

Alloxan reaction, for localization of
a-amino acid groups in proteins,
60

Altmann-Gersh technique, in freezing-
drying, 4

Amine oxidase, bisulfite as binding
agent in localization, 93
chemical staining methods for, 93

a-Amino acid groups in proteins,
chemical staining methods, 60

Amino groups, by acidimetric acetone
titration method, 290
titrimetric techniques, 290

Aminonaphtholsulfonic acid reagent,
use in determination of phos-
phatase, 209

Aminopolypeptidase, by Linderstr0m-
Lang-Holter method, 303

Ammonia, microliter constriction pi-
pettes used for acid in determina-
tion of, 174
by phenol-hypochlorite method of
Russell, 232

Ammonia buffers, for alkalimetric
alcohol method for proteolytic
enzj^mes, 305

Ammonia distillation, apparatus, 167

511

512

SUBJECT INDEX

Ammonia distillation (continued) :
by Briiel-Linderstr0m-Lang-RozitP

method, 283
colorimetric determination by cu-
vette technique, 234
by titrimetric techniques, 283

Amylase, by Linderstr0m-Lang-Engel
"method, 302
by titrimetric technique, 302

Anaerobiosis, test in filling microliter
divers under anaerobic conditions,
374

Analytical electron microscopy. See
Electron microscopy, analytical.

Anfinsen method for diphosphopyri-
dine nucleotide, 396

Anfinsen-Lowry-Hastings method for
microscopy and analysis on same
tissue section, 431

Anhydro Vitamin A. See Vitamin A,
anhydro.

Apparatus. See under specific types of
apparatus.

Arbacia punctulata eggs, separation
of components, 452

Argentaffin reaction for localization
of phenols, 74

Arginase, by Linderstrdm-Lang-Weil-
Holter method, 307
Linderstr0m-Lang-Weil-HoIter ap-
paratus for ammonia distillation,
167
by titrimetric techniques, 306

Arginine, Sakaguchi reaction for stain-
ing, 56

Arginine and arginine-containing pro-
teins, chemical staining methods
for, 56

Armitage method for peroxidase in
blood or bone marrow smears, 91

Arsenic, chemical staining methods, 30

Arsenophosphotungstate reagent,
Benedict's, use in determination
of uric acid, 240

Ascorbic acid, chemical staining
methods, 54
glutathione as inhibitor in, 55
colorimetric determination by cu-
vette technique, 245
by GHck method, 301
reduced, localization of, 55

reduced and oxidized, localization

of, 55
by titrimetric technique, 300
Azide, as inhibitor in localization of

cytochrome oxidase, 94
Azo reaction, for localization of
phenols, 74

B

Balances, 189

Ball-tipped pipettes, in Cartesian
diver manometry, 360
in capillary diver technique, 386

Bauer-Feulgen stain for glycogen,
48

Bayliss-Walker cell, 188

Beckman spectrophotometer, Lowry-
Bessey application in measure-
ment of small volumes, 217

Behrens procedure for isolation of
thvmus and Ivmph cell nuclei,

459

Belanger-Leblond technique of radio-
autography, 157

Benedict's arsenophospho-tungstate
reagent, in determination of uric
acid, 240

Bennett phenylhydrazine reaction for
water-insoluble aldehydes and ke-
tones, 70

Bennett semicarbazide reaction for
water-insoluble aldehydes and ke-
tones, 71

Bensley-Hoerr procedure for guinea
pig liver, 462

Benzidene in detection of peroxidase.
90, 91

Berg ninhydrin test for a-amino acid
groups, 60
Serra-Quieroz Lopes modification.
60

Berg simplified gas analyzer, 328
sources of error, 329

Bessey-Lowry-Brook method for phos-
phata.se, 229

Bessey-Lowry-Brock-Lopez method
for vitamin A and carotene, 250

Bessey-Lowry-Brock-Lopez rotating
nail head method of stirring, 179

Best carmine stain for glycogen, 49

Bethe method for staining chitin, 54

SUBJECT INDEX

513

Bile acids and pigments, chemical

staining methods, 63
Bismuth, chemical staining methods

30
Bisulfite, as binding agent in localiza-
tion of amine oxidase, 93
Bodian method, Dublin application

in localization of melanin, 61
Boell method for total nitrogen, 233
Boell microhter burette, 261
Boell-Needham-Rogers method for

construction of microhter diver,

352
Boettiger method for glycogen, 248
Borchardt method, for localization

of gold, 27

Bordley-Hendrix-Richards method for

creatinine, 211
Bordley-Richards method for uric

acid, 213
Borsook method for allantoin, 243
for creatine, 242
for creatinine, 242
for uric acid, 240
Borsook-Dubnoff digestion oven, 181
Borsook-Dubnoff method (s), for

amide, peptide, and nitrate nitro-
gen, 288
for total nitrogen, 231
Bott capillary tube filter, 177
Bott method for sodium, 203

preparation of filters, 204
Bourne adaption of Carr-Price re-
action for locahzation of carotene
42
Bourne method for alkaline phos-
phatase, 79
Bourne nitroprusside test for sulf-

hydryl and disulfide groups, 37
Bourne silver stain for reduced and

oxidized ascorbic acid, 55
"Braking" pipette, use in capillary
diver technique, 385
in Cartesian diver manometry, 359
Broda method for magnesium, 18
Bruel-Holter-Linderstr0m-Lang

method for nitrogen, 233
Briiel-Holter-Linderstr0m-Lang

Rozits method, for coating vessels
with hydrophobic layer, 169
for nitrogen and ammonia, 283
for digestion in total nitrogen deter-
mination, 234

Burettes, microhter, 255

Calcium, by analytical electron mi-
croscopy, 147
calcium sulfate test for, 17
colorimetric determination by cu-
vette technique, 219
evaluation of eerie sulfate proce-
dure, 272
by Lindner-Kirk cerimetric method,

275
localization bj^ chemical staining

methods, 16
by microincineration, 145
by Senderoy colorimetric method,

219
in serum, determination by Evelyn

Photoelectric Colorimeter, 222
by Siwe acidimetric method, 274
by Siwe iodometric permanganate

method, 273
radioactive, by radioautography, 161
Sobel-Kaye procedure, 273
by titrimetric techniques, 272
Calcium salts, insoluble, interferenc(?
in Gomori method for alkaline
phosphatase, 78
Calcium sulfate test for calcium, 17
Carbon dioxide, calculation in Cun-
ningham-Barth-Ivirk differentia 1
respirometric method, 318
by gasometric techniques, 337
by Scholander-Roughton method

337
stoppers to protect solutions from,
170
Carbon dioxide-oxj'gen-nitrogen mix-
ture, sample analj'sis by Scho-
lander micrometer burette gas
analyzer, 327
Carbon monoxide, by gasometrin
methods, 334
by Scholander-Roughton method,
334
Carbonyl compounds, water-insoluble,
chemical staining methods for, 69
Carboxj'l groups, by Glick method,
290
titrimetric techniques, 290
Carboxypolypeptidases, by Linder-
str0m-Lang-Holter method, 303

514

SUBJECT INDEX

Carcinogenic hydrocarbons, localiza-
tion by direct fluorescence, 105
Carotene, Bessej^-Lowry-Brock-Lopez
method, 250'
chemical staining methods, 42
colorimetric determination by cu-
vette technique, 250
differentiation from A vitamins by

fluorescence, 104
Steiger method for staining, 42
Carotenoids, chemical staining

methods, 42
Carr-Price reactions. Bourne adapta-
tion for localization of carotene,
42
Capillary diver, 382-393
Capillary diver technique, calcula-
tions, 391-392
Capillary respirometry, 314
Capillary tube colorimetric tech-
nique (s), advantage of microcu-
vettes over, 195
apparatus, 196

in determination of chloride, 200
of sodium, 203
of phosphatase, 209
of phosphate, 208
of reducing substances, 210
of creatinine, 211
of uric acid, 213
of urea, 214

of hvdrogen ion concentration,
216
manipulations, 197
methods, 198

preparation of protein-free super-
natants, 198
Capillary tube colorimetry, 195
Capillary tube filter, 177
Carbonate-chloride-phosphate, chemi-
cal staining methods for, 32
Carboxyl groups, by alkalimetric al-
cohol titration method, 290
Carere-Comes Siena Orange method

for potassium, 15
Cartesian diver manometry, 342
capillary diver technique, 382
in determination of cholinesterase,
393
of cocarboxylase, 394
of diphosphopyridine nucleotide,
396

Cartesian diver manometry (con-
tinued) :

of thiamine, 394
methods other than for respiration,

393
microliter diver technique, 343
principles, 342
Caspersson in situ technique of vis-
ible and ultraviolet absorption
histospectroscopy, 114-116
Caspersson photoelectric apparatus,

film density, 138
Castel cupric salt method for arsenic,

30
Castel method for localization of bis-
muth, 32
Castel reagent, modified, for localiza-
tion of bismuth, 31
Castel sulfide method for arsenic, 30
Catalase, by Holter-Doyle method,
310
by titrimetric techniques, 310
Catheptic proteases by Linderstr0m-

Lang-Holter method, 303
Cell nuclei, Dounce-Lan procedure for
isolation, 455
isolation, 454
from avian erythrocytes, 454
from chicken erythrocytes, 454
from leucocytes, 454
from liver, 456, 458
from muscle tissue, 456
from other cells, 456
from thymus and lymph cells,

459
from tumor tissue, 458
isolation of choromatin threads,

461
Laskowski procedure for isolation
of, 454
Cellular components, mechanical

separation, 447
Cellulose, chemical staining methods

for, 43
Centrifugation techniques, sepa-
ration of cellular components,
447
Centrifuge (s), air-driven, 448-450
electrically powered, 451
hydrogen-driven, 449
vacuum-type, 450-451

SUBJECT INDEX

515

Centrifuge (s) (continued):
high-speed types, air-driven "spin-
ning-top," 448
electric motor, 448
electrically powered, magnetically

supported vacuum-tjT^e, 451
hydrogen-driven "spinning top,"

449
plastic-rotor air-driven, 449
for separation of cellular compo-
nents, 448
vacuum-type air-driven, 450
ultra-. See Ultracentrifuge.
vacuum vs. nonvacuum type, 450
Centrifuge microscope, for separation

of cellular components, 447
Cerimetric method for calcium, 275

for reducing sugars, 297
Chemical staining methods in micro-
scopic technicjue, 11
Chemical staining methods, 11. See
also under specific compounds to
be determined,
freezing-drying techniciue, 11
general requirements 11
localization of enzymes, 77
localization of inorganic elements

and radicals by, 14
mounting media, 13
requirements for enzyme methods,
12
Chemical techniques, 165
apparatus, 166

for determination of amount of a
biological sample by nitrogen
content, 426
by nucleic acid content, 426
dilatometric techniques, 413
methods of isolation used in, 165
Chemotherapeutics, interference of
tissue fluorescence in locatization,
108
localization by direct fluorescence,
108

Chevremont-Comhaire method for

riboflavin, 43
Chibnall method for chloroplasts, 468
Chitin, softening, Diaphanol method,
53
Murray method, 53
staining methods, 52-54

Chloride,
colorimetric determination, by capil-
lary tube technique, 200
by cuvette technique, 224
by electrometric titration method
281

by Linderstr0m-Lang-Palmer-Holter
method, 282

by permanganate oxidation method.

281
by Sender oy colorimetric method.

225
by silver nitrate method, 281
by titrimetric techniques, 281
Chloride- phosphate-carbonate, chemi-
cal staining method, 32
Chlorophyll, localization by direct

observation of fluorescence, 105
Chloroplasts, Chibnall method for
isolation. 468
Granick method for isolation, 469
isolation from leaf cells, 468
Menke method for isolation, 468
Neish procedure for isolation, 469
Chloroplatinic acid as precipitant in
determination of potassium, 269
Cholesterol, chemical staining meth-
ods, 38
Schultz test for staining, 41
Cholinesterase, by Cartesian diver
manometric technique, 393
by Glick acidimetric method, 310
by Linderstr0m-Lang-Glick method,
393
Chondroitin sulfuric acid proteins,

Hempelmann method, 46
Christeller method, for localization

of gold, 27
Chromaffin reaction, for localization

of phenols, 74
Chromate method for localization of

lead, 22
Chromatin elements., detection by
Rafalko modification of Feulgen
reagent, 65
Chromatin threads, isolation from

cell nuclei, 461
Claff-Swenson glass capillary elec-
trodes, 184
Clark-Levitan-Gleason-Greenberg ti-
trimetric procedure for sodium,
270

516

SUBJECT INDEX

Claude procedure for isolation of
"large granules," from liver.
464
preparation of extract, 464

Claude procedure for isolation of
"microsomes," 466

Claude procedure for neoplastic cells
of rat, 463

Claude-Potter procedure for isolation
of chromatin threads, 461

Cocarboxylase, determination by Car-
tesian diver manometry, 394
Ochoa-Peters method, 394
Westenbrink method, 395

Coleman preparation of Feulgen re-
agent, 67

Colorimetric methods in determina-
tion of nitrogen and ammonia.
234

Colorometric techniques, 195. See also
under specific compounds to be
determined,
capillary tube technique, 195
cuvette technique, 216

Condensers, in fluorescence micro-
scopy, 100

Conductivity apparatus, 188

Constriction pipettes, 172

Contraction constant(s), for alanyl-
glvcine hvdrolysis by peptidase,
418
in dilatometric techniques, 413

Convection currents, detection in mi-
croliter diver technique, 345

Conway permanganate oxidation
method for chlorides, 281

Conway-Byrne diffusion method for
ammonia, 230

Conway-Byrne glass diffusion cells
for ammonia distillation, 168

Copper, chemical staining methods.
22
removal in titanometric method for
iron, 279

Cowdry modification of Feulgen tech-
nique, 68

Cramer method for nitrate, 35

Creatine, Borsook method, 242

colorimetric determination by cu-
vette technique, 239
Sure-Wilder method, 243

Creatinine, Borsook method, 242

Creatinine (continued) :
colorimetric determination by
capillary tube method, 211
by cuvette technique, 239
Sure-Wilder method, 243
Cretin color test for calcium and
other minerals, 17
gallic acid reagent in, 17
Cretin-Pouyanne method for nickel,

22
Cryostat, in cutting and handling

frozen tissue sections, 428
Cunningham-Barth-Kirk differential

respirometer, 314
Cunningham-Kirk open-tube respir-
ometer, 319
Cunningham-Kirk-Brooks method for

chorides, 281
Cunningham-Kirk-Brooks sintered

glass filter, 176
Cunningham-Kirk-Brooks titrimetric

method for potassium, 268
Cupric salt method of Castel, for

arsenic, 30
Cuvette (s), quartz, use in colorimetric
analysis, 216
Zeiss, use in colorimetric analysis,
216
Cuvette technique (s), in colorimetric
analysis, 216
in determination of allantoin, 239
in determination of ammonia and
nitrogen, 234
of ascorbic acid, 245
of chloride, 224
of creatine, 239
of creatinine, 239
of glycogen, 247

of phosphate and phosphatase, 226
of uric acid, 239

of vitamin A and carotene, 250
methods, 219
Cytochrome oxidase, azide as inhibitor
in localization of, 94
chemical staining methods, 94
phenylurethan in localization of, 94
Cytoplasmic particulates, isolation, 462

Deane-Nesbett-Hastings method for
preservation of glycogen, 49

-SUBJECT INDEX

517

Decalcification, effect upon alkaline

phosphatase, 78
Deductive methods in chemical tech-
niques, 435
Dehydroascoibic acid by titrimetric

method, 301
Density of photographic film, meas-
urement by Caspersson photo-
electric apparatus, 138
by Gunther-Wilcke procedure,

140
by self-recording microphotom-
eter, 138
Density and "reduced weight" by

dilatometric techniques, 420
Deproteinization of blood in determi-
nation of reducing sugars, 298
Desmoglycogen, by Heatley-Lindahl
method, 300
separation from lyoglycogen, 300
Desox>'ribonucleic acid, estimation
by Stowell technique of absorption
histospectroscopy of Feulgen re-
action, 126
Diaphanol method for softening

chitin, 53
Differential centrifugation in isolation

of mitochondria, 462
Differential respirometer of Cunning-
ham-Barth-Kirk, 314
Scholander micrometer burette
type, 324
Digestion oven, Borsook-Dubnoff

type, 181
Digitonin, in detection of free cho-
lesterol, 39
Dilatometric techniques and appara-
tus, 413-417
in measurement of peptidase, 417
p-Dimethylaminobenzylidene rhoda-
nine, for localization in detec-
tion of copper, 23
of mercury, 24
of silver, 26
2,4-Dinitrophenylhydrazine, in deter-
mination of ascorbic acid, 245
Diphenylamine reagent, in determin-
ation of glycogen, 248
use in method for chlorides, 201
Diphosphopyridine nucleotide, An-
finsen method, 396

Diphosphopyridine nucleotide, (con-
tinued) :

determination by Cartesian diver

manometry, 396
extraction from tissue sections, 396
Disulfide groups, chemical staining
methods, 36

Diver:

^artesian. See Cartesian
See Microliter

diven
Divers, i licroliter

diven

Divers, 0 microliter. See Capillary
diver;

Dopa oidase, chemical staining
meth is for, 91
necessit for control runs in detec-
tion, 2

Dounce rocedure for isolation of
nucle 456

Dounce-In procedure for isolation
of ce nuclei, 455

Drugs, (knge of fluorescent color
with miperature, 107

Dublin etUcation of Bodin method
for :3aIization of melanin, 61

DuBois 3vice for drawing micro-
pipe !S, 176

Dubos-Bchet method for staining
ribojcleic acid, 66

Dunn nihod for hemoglobin, 63

E

Edwardi- Scholander - Roughton
mettl for nitrogen, 336

EhrUch saction for localization of
ind(3 and related compounds, 73

Electro(, 183
glass, 3co-Cunningham-Kirk type,
183

glass
184
open.

)illary, Claff-Swenson type,

r measurement of oxj^gen
teni in tissues, 411

for measurement of oxy-

^n tension in tissues, 407

mtages, 410

ixise in blood, 411

sealea capillary glass, Pickford

tvJl86

518

SUBJECT INDEX

Electrodes (continued) :
silver, Linderstr0m-Lang-Palmer-
Holter type, 183

"Electromagnetic flea" in stirring, 179
ferrum reductum, 179
steel ball bearings, 180

Electrometric titration method for
acid or alkali groups, 2')0
for chlorides, 281

Electron microscope, Scoft-Packer
type, 148

Electron microscopy, analytcal, 147

Elements, identification by smission
spectrography, 109

Elftman-Elftman method fr local-
ization of gold, 28

Emission histospectroscopy, 39

Engstrom technique employii; Roent-
gen absorption histospecoscopy,
127

Enzymes, chemical staining lethods,

77

Erythrocytes, avian, isolatio of cell
nuclei from, 454

Esterases, by Glick adimetric
method 309
by titrimetric techniques, )8

Evelyn photoelectric colorimer, de-
termination of serum calum by,
222

Extraction of lipids, methodor, 292
Schmidt-Nielsen method, 93

Ferritin, isolation from hoi liver,

471
Feulgen reaction, Cowdry jdifica-

tion, 68
in detection of aldehydeSS
Oster modification, 68
specificity for aldehydes,
Whitaker use for plant tiies, 67
Feulgen reagent, Coleman (repar-
ation, 67
for localization of polysaarides,

43

in Oster-Schlossman meti for

amine oxidase, 93
Rafalko modification for ection
of chromatin elements,
Feulgen technique, specificitJS, 66

Fiber balance. See Balances.
Filters, in chemical techniques, 176
in fluorescence microscopy, 100
preparation for determination of
sodium by capillary tube tech-
nique, 204
Fixed pipettes, 170
Flotation medium, in microliter diver

technique, 346
Flotation vessel (s), for capillary diver
technique, 385
in microliter diver technique, 345
Fluorescence, change of tissue color
with temperature, 106
direct, of certain parasites, 108
of compounds, 107
of various tissues, 106
direct observation by fluorescence

microscopy, 104
localization of riboflavin, 104
of uranium salts, 108
of vitamin A, 104
"primary," 99
"secondary," 99
spectroscopic analysis, 108
Fluoresence color of drugs, change

with temperature, 107
Fluorescence microscope, set-up, 100
Fluorescence microscopy, 99
characterization of substances, 104
condensers used, 100
filters used, 100

frozen dried tissue vs. formalin-
fixed tissue, 102
immersion media, 101
mounting medium, 101
preparation of tissues, 102
Fluorescence photomicrography,
equipment prerequisites, 103
Fluorescent dyes. Popper method for

localizing lipids, 105
"Fluorochromes," role in fluorescence

microscop5'^, 99
Folin reagent, use in Bordley-Hend-
rix-Richards method for creatin-
ine, 212
"Forerunner," use in fiUing divers, 362
Formol method for tiyptic activity,

306
Forsgren test for bile acids, 64
Fractionation of lipids, method, 292
Schmidt-Nielsen method, 293

SUBJECT INDEX

519

Freezing-drying, 3

in microincineration, 142
Furnaces. See Ovens.

Gallic acid reagent, for localization

of calcium, 17
Gas analysis, 326
Gas analyzer, Berg, 328
Scholander micrometer burette

type, 326
syringe type, of Scholander-Rough-
" ton, 329
Gasometric techniques, 313
for carbon dioxide, 337
for carbon monoxide, 334
for cholinesterase, 393
for cocarboxylase, 394
for diphosphopyridine nucleotide,

396
factors governing selection, 313
for nitrogen, 335
optical lever respirometer, 399
for oxygen, 331
for respiration, 314, 342, 392
for thiamine, 394
Gasometric-volumetric techniques,

313
Gasometry, manometric methods, 342

polarographic methods, 404
Gas volume, total, measurement in

microliter diver technique, 378
Gentian violet stain for starch, 51
Gerard-Cordier adaptation of Pruss-
ian Blue method for iron to
localization of uranium, 29
Gersh method for localization of
chloride-phosphate-carbonate, 33
Gersh-Baker modification of Caspers-
son in situ technique of visible
and ultraviolet absorption histo-
spectroscopy, 116-118
Gersh-Macallum method for local-
ization of potassium, 14
Glick acidimetric method for ester-
ase and lipase, 309
for cholinesterase, 310
Glick method for ascorbic acid, 301
for carboxyl groups, 290

Glick-Fischer adaptation of Gomori
acid phosphatase method to
grains and sprouts, 82-84
Glick-Fischer hanging-drop tech-
nique, 13
Glick-Linderstr0m-Lang-Holter

method for inorganic phosphate,
280
Glutathione, as inhibitor in detection

of ascorbic acid, 55
Glycogen, Boettiger method, 248
chemical staining method, 43
colorimetric determination by cu-
vette technique, 247
Lazarow procedure for isolation

from liver, 466
by titrimetric techniques, 298
total, Heatley titrimetric method.

299
van Wagtendonk-Simonsen-Hack-
ett method, 249
Gmelin test for bile pigments, 63
Gold, chemical staining methods for,

27
Gomori method for glycogen and

mucin, 50
Gomori revised method for acid phos-
phatase, in animal tissues, 81
Gomori revised method for alkaline
phosphatase, 79. See also Phos-
phataseis).
for lipase, 89
Goulliart method for hemoglobin, 62
Graff method for cytochrome oxidase
in fixed tissue, 94
in fresh tissue, 95
Granick procedure for isolation of

chloroplasts, 469
Guinea pig liver, Bensley-Hoerr pro-
cedure, 462
Gunther-Wilcke procedure for meas-
urement of density of photo-
graphic film by Roentgen ab-
sorption histospectroscopy, 140

Hackmann method for sulfonamides,

76
Hale method for acid polysaccharides.

45

520

SUBJECT INDEX

Hamilton-Soiey-Eifhorn inoceduip

for preparation of radioauto-

graphs, 156
Hammett-Chapinan niiroprusside test

for sulfhydryl and disulfide

groups. 87
Hand-Edwards-Caley method for

mercurous and mercuric mercury.

25
Hand pipettes, 172
Hanging-drop technique oi GHck

and Fischer. 13
Harrison-Thomas-Hill procedure for

preparation of ladioautographs.

156
Harvey method for s(>paration of

components of A. punctulnfa

eggs. 452
Hawes-Skavinski diffusion method foi-

nitrogen. 232
Heart muscle, isolation of cytochrome

oxidase particulates. 471
Heating devices, 180
Heatley imi)roved apjiaratus for

optical lever resi)iiometry. 400
Heatlev method for total glycogen.

299
Heatley microliter burette. 260
Heatley-Beienblum-Chain apparatus

for optical lever respiiometry, 399
Heatley -Lindahl metliod for desmo-

and lyoglycogen, 300
Heck-Brown-Kirk cerimetric method

for reducing sugars, 296
Hematoxylin, in Mallory-Parker

method for lead and copper, 23
Hemicelluloses. chemical staining

methods, 43
Hemoglobin, chemical staining

methods. 62
Hempelmann method for differentia-
tion of chondroitin and mucoitin
sulfuric acid proteins, 46
Histospectroscopy. iSee Enns.sion

histospectrascopy, Roentgen ab-
sorption histospectroscopy , Spec-
troscopy, and Ultraviolet absorp-
tion histospectroscopy.
Hollande modification of Courmont-
Andre method for uric acid and
urates, 73

Holmgren and Wilander method for

staining of mucoproteins, 46
Holter flotation medium for micro-
liter diver technique, 347
Holter method of construction of

microliter diver, 352
Holter method for measurement of
volume of small, irregular bio-
logical samples, 432
Holter moist chamber, 182
Holter-Dovle method for catalase

310
Holter-Doyle method for coating
vessels with hydrophobic layer,
169
Holter-Doyle microliter jacketed con-
striction pipettes, 174
Holter-Doyle modification of Linder-
str0m-Lang-Holter method for
reducing sugars, 296
Holter-Doyle vessel for iodometric

titration, 167
Hotchkiss method for polysacchar-
ides, 44
Humphrey dinitrosoresorcinol test

for iron, 21
Hyaluronic acid, chemical staining

methods, 45
Hyaluronidase, in localization of hy-
aluronic acid, 45
Hydrogen ion concentration, meas-
urement by capillary tube tech-
nique. 216. See also Electrodes,
glass.
Hydrophobic layer. Briiel-Holter-
Linderstr0m-Lang-Rozits method
for coating vessels. 169
Holter-Doyle method for coating
vessels, 169
8-Hydroxyquinolates. in fluoroscopic
localization of certain metals, 108
Hydroxyquinoline reagent in detec-
tion of iron, 21
Hypobromite method of Le^y-Palmer
for nitrogen, 231

Immersion media, in fluorescence
microscopy, 101

-SUBJECT INDEX

521

Incineration oven, Linderstr0m-Lang

type, 181. See also Ovens.
Indole and related compounds, chemi-
cal staining methods, 73
Indophenin reaction for localization
of indoles and related compounds,
74
"Indophenol oxidase." See Cyto-
chrome oxidase.
Indo reaction, for localization of

phenols, 74
Iodide, locahzation, 34
Iodine method for staining cellulose,
51
for staining starch, 51
Iodine number of lipids, by Kretch-
mer-Burr method, 294
by Schmidt-Nielsen method, 295
in titrimetric techniques, 294
Iodine, radioactive, b.v radioauto-

graphy, 161
lodometric permanganate method for

calcium, 273
Iron, chemical staining methods, 19
Iron, dinitrosoresorcinol test, 21
ferric, test, 20
ferrous, test, 20
hydroxj'quinoline test, 21
inorganic, locahzation, 20
by Kirk-Bentley method, 277
by microincineration, 145
organic, localization, 20
by Ramsay method, 278
reductor for estimation, 170
by titrimetric technique, 276
Isomerase. See Zymohexase.
Isotopes, radioactive, as tracer ele-
ments, 154

Jackson acetic-carbol-Sudan III

method for staining lipids. 39
Johnson-Kirk ultrafilters, 177

Kay- Whitehead Sudan IV method for
staining lipids, 39

Kinsey-Robison apparatus for am-
monia distillation, 168

Kinsey-Robison method for urea, 286

Kirk microliter burette, 259

Kirk stirring method for open shallow
vessels, 179

Kirk-Bentley method for iron, 277

Kirk-Bentley micro muffle furnace,
180

Kirk-Bentley titration dish, 169

Kjeldahl tube. Levy, 167

Kretchmer-Holman-Burr method for
iodine number of lipids, 294

Kabat and Furth modification for
alkaline phosphatase, 79

Lactobacillus casei culture and ino-
culum, in determination of ribo-
flavin, 440

Laidlaw method for dopa oxidase, 92

"Large granules," Claude procedure
for isolation from liver, 464
definition, 462

Laskowski procedure for isolation of
cell nuclei, 454

Lazarow procedure for isolation of
lipoprotein and glycogen from
liver, 466
for isolation of liver nuclei, 458

Lead, chemical staining methods, 22
by microincineration, 145
radioactive, by radioautography, 161

Leschke procedure for localization of
urea, 75

Leucocytes, isolation of cell nuclei,
454

Levy Kjeldahl tube, 167

Levy Nesslerization method for ni-
trogen, 231, 237

Levy-Palmer hypobromite method for
nitrogen, 231

Linderstr0m-Lang incineration oven,
181

Linderstr0m-Lang titrimetric method
for sodium and potassium, 266

Linderstr0m-Lang-Duspiva alkalimet-
ric alcohol method for proteolytic
enzymes, 304

522

SUBJECT INDEX

Linderstr0m-Lang-Duspiva method

for carboxyl groups, 290
Linderstr0m-Lang-Engel method for

amylase activity, 302
Linderstr0m-Lang-Ghck method for

cholinesterase, 393
Linderstr0m-Lang-Holter acidimetrir
acetone method for proteolj'tic
enzymes, 303
Linderstr0m-Lang-Holter automatic
microliter pipettes, 174
diffusion method for ammonia, 230
direct method for amino groups, 290
"electromagnetic flea" for stirring,

179
method for reducing sugars, Holter-

Doyle modification, 296
method for total nitrogen, 233
method for urea, 286
microliter burettes, 256
microliter constriction pipettes, 172
reaction vessel, 167
Linderstr0m-Lang-Lanz dilatometric

method for peptidase, 419
Linderstr0m-Lang-Mogensen method
for cutting and handling tissue
sections, 428
Linderstr0m-Lang-Palmer-Holter

electrometric method for chlo-
rides, 282
silver electrode, 183
Linderstr0m-Lang-Weil-Holter appa-
ratus for ammonia distillation in
measurement of arginase, 167
method for arginase, 307
Lindner-Kirk, cerimetric method for
calcium, 275
method for phosphorus, 280
titrimetric method for sodium, 271
Lipase (s), chemical staining methods,
88
by Glick acidimetric method, 309
by titrimetric techniques, 308
"Tweens" as substrates, 89
Lipid, chemical staining methods, 38
extraction and fractionation of
saponifiable fraction, 294
by Schmidt-Nielsen method, 293
of unsaponifiable fraction, 293
localization by fluorochromes, 105
by Schmidt-Nielsen method, 291
titrimetric techniques, 291

Lipoprotein, Lazarow procedure for
isolation from liver, 466

Lison method for polysaccharide sul-
fate compounds, 47

Lison modification of chromaffin re-
action for localization of phenols,
74

Liver, Bensley-Hoerr procedure, 462
Claude procedure for isolation of
"large granules," 464
of "microsomes," 467
Lazarow procedure for isolation of
lipoprotein and glycogen from, 466

Lloyd reagent in determination of
creatine and creatinine, 242

Loele method for a-naphthol oxidase,
96

Loscalzo-Benedetti-Pichler microliter
burette, 263

Lowry quartz fiber balance, 189

Lowry cjuartz torsion balance, 191

Lowry-Bessey adaptation of Beckman
spectrophotometer in measure-
ment of small volumes, 217

Lowry-Bessey method for riboflavin,
440

Lowry-Lopez method for inorganic
phosphate in presence of labile
phosphate esters, 228

Lowry-Lopez-Bessey method for
ascorbic acid, 245
charring in, 247

Lundsteen-Vermehren method for in-
organic phosphate and phospha-
tase, 227

Lymph cells, isolation of nuclei, 459

Lyoglycogen, by Heatley-Lindahl
method, 300
separation from desmoglycogen, 300

Lysolecithin in hemolysis, 454

M

Macallum method for localization of

potassium, Gersh's modification,

14
Macallum technique in determination

of iron, 20
MacKee-Herrmann-Baker-Sulzberger-

method for sulfonamides, 76

SUBJECT INDEX

523

Magnesium, by analytical electron
microscopy, 147
by microincineration, 145
chemical staining methods, 18

Magnesium uranium acetate reagent,
use in method for sodium, 204

Mallory-Parker hematoxylin method
for localization of lead and cop-
per, 23

Mallory-Parker methylene blue
method for lead and copper, 24

Manometer, for microliter diver tech-
nique, 348
use of ])ressure regulator vs. sy-
ringe for fluid adjustment, 348

Manometer fluid in microliter diver
technique, 348

Manometric methods of gasometiy,
342

Manometric techniques of gasometrj',
compared with volumetric tech-
niques, 313

Manometry, Cartesian diver, 342

McJunkin method for peroxidase in
tissue sections, 90

Melanin, chemical staining methods,
61
isolation, 472

Melanoblasts, dopa oxidase in identi-
fication, 91

Membrane interferometer volumetrv,
340-341

Mendel-Bradley method for zinc, 19

Menke method for chloroplasts, 468

Menten-Junge-Green coupling method
for localization of alkaline phos-
phatase, 79

Mercurv, chemical staining methods,
24 '
mercuric, staining method, 25
mercurous, staining method, 25

Metals, localization by direct obser-
vation of fluorescence, 108

Metaphosphoric acid as extraction
medium in determination of as-
corbic acid, 301

Methylene blue, in localization of
succinic dehydrogenase, 96
in Mallory-Parker method for lead
and copper, 24

Methyl green stain for starch, 51

Microbiological techniques, 439
Microcuvettes, advantage over capil-
lary tube colorimetry, 195
Microelectrode measurement of local

oxygen tension in tissue 404
Microelectrodes. platinum, for meas-
urement of oxygen tension in
tissue, 404
Microincineration, 140-146
Microliter burettes, 255
Microliter diver, 350
Microliter diver constant, calculation,
378
measurement of total gas volume,
378
Microliter diver techniques, ball-
tipped pipettes, 360
"braking" pipette, 359
calculations, 378

calculations for change in gas vol-
ume, 379
Cartesian diver manometry, 342
connecting manifold, 347
experimental procedure, 376
filling the diver, 362

technique for large biological ob-
jects, 363
measurement, 374
saturation of flotation medium
with gas, 374
pipettes, 356
calibration, 361

for dilute aqueous solutions, 357
for precise deposition of drop in

diver neck, 360
for transfer of cells and tissue

pieces, 359
for viscous liquids, 359
holding devices, 361
removal of objects from divers, 363
spreading bottom drop in diver,

means, 362
temperature control apparatus, 344
Microliter pipettes, 170
Micrometer, use in microliter burettes,

255
Microphotometer, self-recording, for
measurement of photographic
film, density, 138
Micropipettes. See Pipettes.
Microscope, polarizing. See Polarizing
microscope.

524

SUBJECT INDEX

Microscopy. See Electron microscopy
and Fluorescence microscopy.

"Microsomes," Claude procedure for
isolation, 466
of cytoplasm, 465

Microtomes, for preparation of tissuo
sections of accurate thickness, 427

"Millimole unit" of phosphatase ac-
tivity, definition, 230

Millon reaction for tyrosine in pro-
teins, 59

Milovidov method for starch, 51

Minerals, total distribution by micro-
incineration, 140

Mitochondria, isolation, 462

Moist chambers, 181

Molybdate staining method for phos-
phate, Serra-Queirez-Lopez modi-
fication, 34

Molybdic-sulfuric acid reagent in
determination of phosphatase,
209
in determination of phosphate, 208

Montgomery method for hydrogen
ion concentration, 216

Mounting of flotation vessels in micro-
liter diver technique, 346

Mounting media, in fluorescence mi-
croscopy, 101
for tissue sections, 13

Mucin, chemical staining methods, 43,
47

Mucoitin sulfuric acid proteins, Hem-
pelmann method, 46

Mucoproteins, chemical staining
method, 43, 46

Muffle furnace, micro, Kirk-Bentley
type, 180

Murexide test for certain purines, 72

Murray method for softening chitin,
53

Muscle, elementary composition by
radioautography, 131

N

"Nadi oxidase." See Cytochrome oxi-
dase.

"G. Nadi Oxidase." See Graff method
for cytochrome oxidase.

"M. Nadi Oxidase." See Graff method
J or cytochrome oxidase.

Nadi reagent in detection of cyto-
chrome oxidase, 95
a-Naphthol oxidase, Loele method, 96
Needham-Boell method for total ni-
trogen, 231
Neish procedure for chloroplasts, 469
Neoplastic cells of rat, Claude pro-
cedure, 463
Nessler reagent in determination oi

nitrogen by Levy method, 237
Nesslerization in determination of

total nitrogen, 231
Nickel, chemical staining methods, 22
Nicotinic acid and amides, question
of detection by fluorescence, 104
Nitrate, chemical staining method, 35
Nitrogen,
amide, by Borsook-Dubnoff method,

288
by Boell method, 233
by Borsook-Dubnoff method, 231
by Bruel-Holter-Linderstr0m-Lang-

Rozits method, 233
colorimetric determination, 230
digestion mixtures, 237
digestion procedure, 234
by Edwards-Scholander-Roughton

method, 336
gasometric determination, 335
by Hawes-Skavinski method, 232
by Levy method, 237
by Levy-Palmer method, 231
by Linderstr0m-Lang-Holter

method, 233
microliter constriction pipettes used

in determination, 174
by Needham-Boell method, 231
nitrate, by Borsook-Dubnoff

method, 288
peptide, by Borsook-Dubnoff

method, 288
titrimetric determination, 283
by Tompkins-Kirk method, 232
Nitrogen-carbon dioxide-oxygen mix-
ture, sample analysis by Scho-
lander micrometer burette ga?
analyzer, 327
p-Nitrophenyl phosphate as substrate
in detection of phosphatase, 229
Nitroprusside in detection of sulfhy-
dryl and disulfide groups, 36

)ii w n \^ ^ ^ I ^ \

1 t D

- SUBJECT INDEX

525

Nitro reaction for localization of in-
doles and related compounds, 74

Nitrosamino reaction for localization
of indoles and related compounds
74

Norberg centrifugation tubes, 167

Norberg method for phosphorus, 124

Norberg technique of visible and
ultraviolet absorption histospec-
troscopy, 120

Norberg titrimetric method for po-
tassium, 268

Nucleic acids, chemical staining
methods, 65

Oxygen, by gasometric methods, 331
by Roughton-Scholander method,

331
calculation in Cunningham-Barth-
Kirk differential respirometric
method, 318
Oxygen-nitrogen-carbon dioxide mix-
ture, sample analysis by Scho-
lander micrometer burette gas
analyzer, 327
Oxygen tension iu tissue, calibration
and measuring instruments, 406
local, microelectrode measurement,
404

O

Ochoa-Peters method for thiamine

and cocarboxylase, 394
Okamoto method for localization of

mercury, 24
Okamoto-Mikami-Nishida method for

localization of palladium, 29
Okamoto-Utamura method for locali-
zation of copper, 23
Okamoto-Utamura-Akagi method for
localization of gold, 28
of palladium, 29
of platinum, 29
of silver, 26
Okkels method, for localization of

gold, 27
Open-tube respirometer. See iJes-

pirometer.
Optical lever respirometr>', 399
Organic substances and groups, chemi-
cal staining methods, 38
Osazone reagent in determination of

ascorbic acid, 245
Oster modification of Feulgen tech-
nique, 68
Oster-Mulinos method for differentia-
tion of "true" and "pseudo" Feul-
gen reactions of aldehydes, 65
Oster-Schlossman method for amine

oxidase, 93
Ovens, Borsook-Dubnoff type, 181
Kirk-Bentley type, 180
Linderstr0m-Lang type, 181
microincineration, 142

Packer-Scott method for freezing-
drying, 5

Palladium, chemical staining methods,
29

Pap ammoniacal silver nitrate method
for glycogen, 48

Pectins, chemical staining method, 43

Pepsin by Linderstr0m-Lang-Holter
method, 303

Peptidase, by dilatometric techniques,
417
by Linderstr0m-Lang-Lanz dilato-
metric method, 419
in determination of peptide nitro-
gen, 289

Peptides, racemic, as substrates in
determination of proteolytic en-
zymes, 303, 304

Permanganate. See lodometric per-
ynanganate method for calcium.

Permanganate oxidation method for
chlorides, 281

Peroxidase, chemical staining methods
for detection, 90
use of benzidine in detection, 90

Phenol (s), alkaline, as reagent in
Russell method for ammonia, 238
chemical staining methods for, 74

Phenol-hypochlorite method of Rus-
sell, for ammonia, 232

Phenylurethan in localization of cy-
tochrome oxidase, 94

Phosphatase (s), acid, chemical stain-
ing methods, 80-85

•<v \

520

SUBJECT INDEX

Phosphatase (s), (continued) :

Glick-Fischer modification, 82
Gomori revised method, 81
Moog modification, 81
Wolf-Kabat-Newman modifica-
tion, 80
alkaline, chemical staining methods,
78-80
Bourne modification, 78
Gomori revised method, 79
Kabat-Furth modification, 78
Menten-Junge-Green method, 79
Takamatsu method, 78
Yin modification, 79
by Bessey-Lowry-Brock method, 229
colorimetric determination by capil-
lary tube technique, 209
by other chemical staining methods,

85
by Weil-Russell method, 209
Phosphatase and inorganic phosphate,
Lundsteen-Vermehren method,
227
Phosphatase and phosphate colori-
metric determination by cuvette
technique, 226
Phosphate, chemical staining methods,
34
colorimetric determination b.y capil-
lary tube technique, 208
Glick-Linderstr0m-Lang-Holter

method, 280
in presence of labile phosphate
esters, Lowry-Lopez method.
228
and phosphatase, Lundsteen-Ver-
mehren method, 227
Phosphate-carbonate-chloride, chemi-
cal staining methods, 32
Phosphate and phosphatase, colori-
metric determination by cuvette
technique, 226
Phosphorus, by colorimetric tech-
niques, 208, 226
by Lindner-Kirk method, 280
by Siwe method, 226
by titrimetric techniques, 280
Norberg microscope photometric

method, 124
radioactive, by radioautography, 161

Photographic emulsion, preparation
by Belanger-l^eblond technique,
157

Photometric endpoints in titrimetric
techniques, 265

Photomicrography, 102

Pickford sealed-in capillary glass elec-
trodes, 186

Pigments, localization by direct ob-
servation of fluorescence, 105

Pipettes, microliter, 170

"Plasmal," chemical staining methods,
65

Platinum, chemical staining method,
29

Polarizing microscope, in detection
of cholesterol, 39

Polarographic methods of gasometry,
404
measurement of oxygen tension in
tissue, 404

Policard technique of emission histo-
spectroscopy, 109

Pollack method for calcium, 16

Polysaccharides, acid, chemical stain-
ing methods, 45
chemical staining methods, 43-44

Polysaccharide sulfate compounds,
Lison method, 47

Porphyrin, localization by direct ob-
servation of fluorescence, 105

Post and Laudermilk method for cell-
ulose, 52

Potassium, by Cunningham-Kirk-
Brooks titrimetric method, 268
by microincineration, 144
by titrimetric technique, 268
localization bj^ chemical staining

methods, 14
use of chloroplatinic acid as pre-
cipitant, 269

Potassium and sodium (combined).
by titrimetric technique, 265
Linderstr0m-Lang method, 266

"Primary" fluorescence, 99

"Product 81" as substrate for lipase.
89

Prolinepeptidase by Linderstr0m-
Lang-Holter method, 303

' SUBJECT INDEX

527

Proteases, catheptic, by Linderstr0m-
Lang-Holter method, 303
peptic, by Linderstr0m-Lang-Holter

method, 303
tryptic, by Linderstr0m-Lang-Dus-
piva method, 304
by Weil method, 306
Protein reactions, brief survey, 56
Proteins, arginine and arginine-con-
taining, chemical staining meth-
ods, 56
Proteolytic enzymes, by acidimetric
acetone method, 303
by alkalimetric alcohol method, 304
by formol method, 306
by titrimetric methods, 302
Prussian Blue test for ferric iron, 20
Purines, chemical staining methods,
72

Quartz torsion balance. See Balances.
Quinalizarin reagent in determination
of magnesium, 18

R

Rachele, device for drawing micro-
pipettes, 176

Radioactive elements, detection by
radioautography, 153

Radioactive isotopes as- tracer ele-
ments, 154

Radioautographs, 156

Radioautography, 153. See also
Radioautographs and under spe-
cific compounds to be determined,
advantages, 156

Belanger-Leblond preparation of
photographic emulsion for use in.
157
limitations on radioactive isotopes

studied, 153
role in use of tracers 153

Rafalko modification of Feulgen re-
agent for detection of chromatin
elements, 65

Ralph method for hemoglobin, 62

Ramsay method for iron, 278
Ramsay titanometric method for iron,

removal of copper, 279
Rapkine nitroprusside test for sulf-

hydryl and disulfide groups, 36
Reaction rate, mea.surement by dila-

tometric techniques, 417
"Reduced weight," definition, 421
by dilatometric techniques, 420
Reducing substances, colorimetric de-
termination by capillary tube
technique, 210
Reducing sugars. See Sugars.
Redactor, in chemical techniques, 170

for estimation of iron, 170
Respiration, use of Capillary diver

technique in, 382
Respiration rates, factors governin,q;
choice of capillary diver for, 392
Respirometer, differential. See Dif-
ferential respirometer.
open-tube, 319
Respirometry, optical lever. See Opti-
cal lever respirometry.
Rhodanine, p-dimethylaminobenzyli-
dene, for localization of copper,
23
for localization of mercury, 24
of silver, 26
Riboflavin, chemical staining methods,
43
localization by direct observation of
fluorescence, 104
by spectroscopic analysis of fluor-
escence, 109
by Lowry-Bessey method, 440
by microbiological techniques, 439
factors influencing determination.
439
Ribonuclease, use in Dubos-Brachet
method for detection of ribonu-
cleic acid, 66
Ribonucleic acid, Dubos-Brachet
method, 66
Turchini, Castel, and Kien method
for differentiation from thymonu-
cleic acid, 66, 69
Roberts method for localization of

gold, 27
Roe-Kuether method for ascorbi-o
acid, 245

528

SUBJECT INDEX

Roentgen absorption histospectros-

copy, 127-138
Roentgen tube, for primary excitation,
in Roentgen absorption histospec-
troscopy, 134
for secondary excitation, in Roent-
gen absorption histospectroscopy,
135
Romieu reaction for tryptophane in

proteins, 59
Roughton-Scholander method for oxy-
gen, 331
Russell phenol-hypochlorite method
for ammonia, 232, 238

S

Sakaguchi reaction for staining argi-

nine, 56
Saponin for hemolysis, 454
Schmidt-Nielsen centrifuge for mixing
liquids in sealed capillary tubes,
178
Schmidt-Nielsen method for extrac-
tion and fractionation of lipids,
293
for iodine number, 295
for lipids, 291
Schneider method for localization of

uranium, 29
Scholander microliter burette, 262
Scholander micrometer burette dif-
ferential respirometers, 324
Scholander-Roughton method for car-
bon monoxide, 334
for carbon dioxide, 337
Scholander-Roughton syringe gas ana-
lyzer, 329
collecting blood samples, 331
Schultz cholesterol test, 41
Schulze method for staining chitin,

54
Scott incineration furnace, nitrogen-
oxygen mixture in, 143
Scott incineration procedure, 143
Scott-Packer analytical electron mi-
croscope, 148-151
Scott- Williams technique of emission

histospectrography, 110
"Secondary" fluorescence, 99

Selenium-acid mixtures, microliter
constriction pipettes used for, 174
Semenoff method for succinic dehy-
drogenase, 96
Senderoy method for calcium, 219

for chloride, 225
Sen method for localization of urease

77
Separation, of components of A. punc-
tulata eggs, 452
mechanical, of cellular components,
447
Serra method for arginine and argi-

nine-containing proteins, 57
Serra-Queiroz Lopes modification of
Berg ninhydrin test for a-amino
acid groups, 60
Serra-Queiroz Lopes modification of
molybdate method for phosphate.
34
Siena Orange, in Carere-Comes

method for potassium, 15
Silicon by microincineration, 145
Silver, chemical staining methods, 25
Silver nitrate method for chlorides, 281
Sintered glass filter, 176
Sisco-Cunningham-Kirk glass elec-
trode, 183
Siwe acidimetric method for calcium,

274
Siwe iodometric permanganate method

for calcium, 273
Siwe method for inorganic phos-
phorus, 226
Sobel-Kaye procedure for calcium,

273
Sodium, by Clark-Levitan-Gleason-
Greenberg titrimetric procedure,
270
colorimetric determination by cap-
illary tube technique, 203
determination by difference from
sodium-potassium and potassium,
271
by Lindner-Kirk titrimetric method,

271
by microincineration, 144
by titrimetric technique, 270
use of zinc uranyl acetate as pre-
cipitant, 271
Sodium alizarin sulfonate for local-
ization of calcium, 16

SUBJECT INDEX

529

Sodium and potassium, Linderstr0m-

Lang method, 266
Sodium and potassium (combined) by

titrimetric technique, 265
Sonic vibrations for disruption of

spermatozoa, 471
Spectrograph for microanalysis in

Roentgen absorption histospec-

troscopy, 136
Spectrograph, vacuum. See Vacuum

spectrograph.
Spectroscopy. See also Histospectros-

copy.
in analysis of fluorescence, 108
Spermatozoa, disruption bj' sonic vi-
brations, 471
"Spinning top" centrifuges. See

Centrifuges.
Staining methods. See Chemical stain-
ing methods and under specific

compounds to be determined.
Starch, chemical staining methods,

43, 51
detection by polarized light, 51
Steiger method for carotene in leaves,

42
Stein test for bile pigments, 63
Stirring devices, 178
Stoneburg procedure for isolation of

cell nuclei, 456
Stoppers, 169
Stowell technique of absorption histo-

spectroscopy, 125
of Feulgen stain for desoxyribo-

nucleic acid, 66, 126
Streptococcus pyogenes, isolation of

particulate from, 471
Strontium, radioactive, by radioautog-

raphy, 161
Submicroscopic particulates of cyto-
plasm, 465
Succinic dehydrogenase, chemical

staining methods, 96
Sudan dye, Telford Govan technique

for staining lipids in aqueous

media, 40
Sudan IV for localization of lipids, 39
Sugars, reducing, determinution by
Heck-Brown-Kirk method, 297

determination by Holter-Doyle
modification of Linderstr0m-
Lang-Holter method, 296

Sugars (continued) :

by titrimetric techniques, 296

Sulfate compounds, polysaccharide,
Lison method for, 47

Sulfhydryl groups, free, chemical
staining method, 36
protein-bound, chemical staining

method, 37
total, chemical staining method, 37

Sulfide method of Castel, for arsenic,
30

Sulfonamides, chemical staining meth-
ods, 75

Sulfur, radioactive, by radioautog-
raphy, 161

Sumner reagent in determination of
reducing substances, 211

Supernatants, trichloroacetic acid,
preparation, 199
tungstic acid, preparation, 198
zinc sulfate-sodium hydroxide, prep-
aration, 200

Sure-Wilder method for creatine and
creatinine, 243

Syringe pipettes, 176

T

Telford Govan technique for Sudan
dye staining of lipids in aqueous
media, 40

Thiamine, determination by Cartesian
diver manometry, 394
localization by spectroscopic analy-
sis of fluorescence, 109
Ochoa-Peters method for, 394
Westenbrink method for, 395

Thomas method for arginine and ar-
ginine-containing proteins, 58

Thomas-Lavollay hydroxyquinoline
test for iron, 21

Thymonucleic acid, Turchini, Castel,
and Kien method for differenti-
ation, from ribonucleic acid, 66.
69

Thymus cells, isolation of nuclei from,
459

Tissues, changes of fluorescence color
with temperature, 106

530

SUBJECT INDEX

Tissiu'is {cimlinued) :

preparation for fluorescence micros-
copy, 102
Tissue sections, frozen, preparation,
427

method for cutting and handling.
428

microscopic examination and chemi-
cal analysis on same section, 430
Titanometric method for iron, 276
Titration. See also Electrometric ti-
trntion.

holders for tubes in, 170
Titration vessel of Kirk and Bentley,

169
Titrimetric techniques, 255

for acid groups, 290

for alkali groups, 290

for amide, peptide, and nitrate
nitrogen, 288

for amino groups, 290

for amylase, 302

for arginase, 306

for ascorbic acid, 300

for calcium, 272

for catalase, 310

for chlorides, 281

for esterases, 308

for glycogen, 298

for iodine number of lipids, 294

for iron, 276

for lipases, 308

for lipid, 291

for nitrogen and ammonia, 233, 283

for phosphorus, 280

for proteolytic enzymes, 302

for reducing sugars, 296

for urea, 286

for urease, 287

methods, 265

photometric endpoints, 265
Tobias-Gerard respirometer, 322
Toluidene blue stain for starch, 51
Tompkin.s-Kirk diffusion method for

nitrogen, 232
Torsion balances, 189, 191
Tracers, role of radioautography in

use of, 153
Trichloroacetic acid supernatants,

preparation, 199
Tryptic activity, formol method for,

" 306

Tryptophane in proteins, chemical
staining methods, 58

Tube and pestle, in chemical tech-
niques, for grinding tissue, 170

Tube holders, 170

Tungstic acid supernatants, prepara-
tion, 198

Turchini, Castel, and Kien method
for differentiation of ribo- and
desoxyribonucleic acids, 66, 69

Turchini, Castel, and Kien method
for nucleic acids, 66, 69

Turnbull's blue test for ferrous iron,
20

"Tweens" as substrates for localiza-
tion of lipase, 89

TjTosine in proteins, chemical stain-
ing methods, 59

U

Ultracentrifuge, Svedberg oil-driven,

448
Ultrafilters, 177
Ultraviolet absorption histospectros-

copy, 113
Uranium, chemical staining methods,
29
by microincineration, 145
Uranium salts, detection by fluores-
cence, 108
Urates, Hollande modification of
Courmont-Andre method for
localization of, 73
Urea, chemical staining methods, 75
colorimetric determination by

capillary tube method, 213
determination by use of urease, 286
by Kinsey-Robinson method, 286
by Linderstr(7(m-Lang-Holter

" method, 286
by titrimetric techniques, 286
Urease, chemical staining methods, 77
in determination of allantoin, 243
in determination of arginase, 307
in determination of urea, 286
titrimetric techniques for, 287
Uric acid, by Borsook method, 240
colorimetric determination by capil-
lary tube method, 213

SUBJECT INDEX

531

Uric acid {continued) :

by cuvette technique, 239
Hollande modification of Cour-
mont-Andre method for localiza-
tion of, 73

Vacuum spectrograph for tissue analy-
sis in Roentgen absorption histo-
spectroscopy, 137
van Wagtendonk-Simonsen-Hackett
method for glycogen, 248

Vessels used in chemical techniques,
166

Visible and ultraviolet absorption
histospectroscopy, 113

Vitamin A, chemical staining methods,
42
colorimetric determination by cuv-
ette technique, 250
determination by method of Bessey-

LowTy-Brock-Lopez, 250
differentiation from carotene by

fluorescence, 104
localization by direct observation
of fluorescence, 104

Vitamin Ai, localization by direct ob-
servation of fluorescence, 104

Vitamin A2, localization by direct ob-
servation of fluorescence, 104

Vitamin K, localization by fluores-
cence, 105

Volume, Holter method of measure-
ment, 432

Volumetric techniques of gasometrj',
313

Von Kossa silver test for calcium, 17

W

Wachstein-Zak method for bismuth,

31
Walker method for phosphate, 208
Walker-Hudson method for urea, 214

Walker-Reisinger method fir deter-
mination of reducing substances,
211

Waterhouse method for localization
of copper, 23

Wavelength and analysis lines of ab-
sorption edges of certain elements.
129

Weight, apparatus for determination,
423

Weil formol method for tryptic
activity, 306

Weil-Russell method for phospha-
tase, 209

Westenbrink method for thiamine
and cocarboxylase, 395

Westfall-Findley-Richards method for
chlorides 200

Whitaker, Feulgen technique for
plant tissues, 67

Wigglesworth silver nitrate method
for chlorides, 281

Williams-Scott photoelectric appara-
tus, for dark-field photometry and
densitometry, 113
for estimation of ash in micro-
incineration 146

Windaus digitonin test for free cho-
lesterol, 39

Winkler solution, in carbon monoxide
determination, 334

Xanthydrol method for localization
of urea, 75

Zander method for staining chitin, 54
Zeiss cuvettes. See Cuvettes, Zeiss.
Zinc, chemical staining method, 18
Zinc sulfate-sodium hydroxide super-

natants, preparation, 200
Zinc uranyl acetate as precipitant in

determination of sodium, 271
Zymohexase, chemical staining

methods, 87

1

r