673 
 G55m Ghflmberlain 
 
 Southern Branch 
 of the 
 
 University of California 
 
 Lo8 Angeles 
 
 
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 This book is DUE on the last date stamped below. 
 
 MAY 3 1921 
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 MAR 7 1930 
 
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 MAY 1 ^^34 
 
 Tf 6 6 1935 
 
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STATE nOSriAL SCHOOL, 
 
 METHODS IN PLANT HISTOLOGY 
 
METHODS 
 
 IN 
 
 PLANT HISTOLOGY 
 
 BY 
 
 CHARLES J. CHAMBERLAIN, Ph.D. 
 
 Instructor in Botany in the University of Chicago 
 
 e/3 2. 
 
 CHICAGO 
 
 Wqz "ClnlversftB of Cbfcaoo press 
 1901 
 
 8\3^ 
 
COPYRIGHT, I9OI 
 
 BY THE UNIVERSITY OF CHICAGO 
 
 CHICAGO, ILLINOIS 
 
^^ 
 
 y 
 
 STATE NORMAL SCHOOL, 
 
 QK 
 C35.-VV 
 
 PREFACE. 
 
 This book has grown out of a course in histological tech- 
 nique conducted by the author at the University of Chicago. 
 The course has also been taken by non-resident students through 
 the Extension Division of the University. The Methods were 
 published over a year ago as a series of articles in the Journal 
 of Applied Microscopy, and have called out numerous letters of 
 commendation, criticism, suggestion, and inquiry. The work 
 has been thoroughly revised and enlarged by about one-half. It 
 is hoped that the criticism and suggestion, and also the expe- 
 rience gained by contact with both resident and non-resident 
 students, have made the directions so definite that they may be 
 followed, not only by those who work in a class under the 
 supervision of an instructor, but also by those who must work 
 in their own homes without any such assistance. 
 
 More space has been devoted to the paraffin method than to 
 any other, because it has proved to be better adapted to the 
 needs of the botanist. The celloidin method, the glycerine 
 method, and free-hand sectioning are also described, and their 
 advantages and disadvantages are pointed out. 
 
 The first part of the book deals with the principles of fixing 
 and staining, and the various other processes of microtechnique, 
 while in the later chapters these principles are applied to spe- 
 cific cases. This occasions some repetition, but the mere pres- 
 entation of general principles will not enable the beginner to 
 make good mounts. 
 
 The illustrations and notes in the later chapters are not 
 intended to afford a study of general morphology, but they 
 
vi Methods in Plant Histology 
 
 merely indicate to students with a limited knowledge of plant 
 structures the principal features which the preparations should 
 show. The photomicrographs were made from the author's 
 preparations by Dr. W. H. Knap, and figs. 52, 57, and 59 were 
 drawn by Miss Eleanor Tarrant ; all other figures of plant 
 structures were made from the author's drawings. 
 
 Corrections and suggestions will be heartily appreciated. 
 
 Charles J. Chamberlain. 
 Chicago, 
 June I, igoi. 
 

 CONTENTS. 
 
 PART I. 
 
 Pagb 
 
 Chapter I. Apparatus i 
 
 Chapter II. Reagents 7 
 
 Killing and Fixing Agents 7 
 
 Stains 7 
 
 Formulae for Alcohols 9 
 
 Miscellaneous Reagents 10. 
 
 Arrangement of the Outfit 10 
 
 Chapter III. Temporary Mounts 11 
 
 Chapter IV. The General Method 13 
 
 Killing and Fixing 13 
 
 Washing 14 
 
 Hardening and Dehydrating 14 
 
 Clearing 16 
 
 The Transfer from Clearing Agent to Paraffin 16 
 
 The Paraffin Bath 17 
 
 Imbedding 18 
 
 Cutting 19 
 
 Fixing Sections to the Slide 20 
 
 Removal of Paraffin 21 
 
 Removal of Xylol 21 
 
 The Transfer to the Stain 21 
 
 Clearing 22 
 
 Mounting in Balsam 22 
 
 A Tentative Schedule for Paraffin Sections 23 
 
 Chapter V. Killing and Fixing Agents , . . 25 
 
 The Alcohols 25 
 
 The Chromic Acid Group 26 
 
 Picric Acid 30 
 
 Corrosive Sublimate 30 
 
 Formalin - 31 
 
 General Hints on Fixing 31 
 
 Chapter VI. Staining 33 
 
 The Hsematoxylins 34 
 
 The Carmines 39 
 
 The Anilins 41 
 
viii Methods in Playit Histology 
 
 Chapter VII. General Remarks on Staining 47 
 
 Chapter VIII. Practical Hints on Staining 53 
 
 Chapter IX. The Celloidin Method 55 
 
 Chapter X. The Glycerine Method 58 
 
 PART II. 
 
 Chapter XI. Thallophytes — Alg^e 63 
 
 Cyanophycese ,.,.... 63 
 
 Chlorophycese 65 
 
 Phseophyceae 73 
 
 Rhodophycese 75 
 
 Chapter XII. Thallophytes — Fungi 77 
 
 Schizomycetes 77 
 
 Myxomycetes 78 
 
 Phycomycetes 79 
 
 Ascomycetes 80 
 
 ^cidiomycetes 84 
 
 Basidiomycetes 87 
 
 Lichens 88 
 
 Chapter XIII. Bryophytes — Hepatic^e 89 
 
 Chapter XIV. Bryophytes — Musci 97 
 
 Chapter XV. Pteridophytes — Filicine^e 103 
 
 Chapter XVI. Pteridophytes — Equisetine^e 115 
 
 Chapter XVII. Pteridophytes — LycopodinejE 117 
 
 Chapter XVIII. Spermatophytes — Gymnosperms 119 
 
 Chapter XIX. Spermatophytes — Angiosperms 129 
 
 Chapter XX. Labeling and Cataloguing Preparations ... 141 
 
 Chapter XXI. A Class List of Preparations . 143 
 
 Chapter XXII. Formula for Reagents 149 
 
 Index . 157 
 
Part I. 
 CHAPTER I. 
 
 The following list of apparatus includes a fair equipment for 
 histological work : a microscope magnifying at least 400 diam- 
 eters ; a hand microtome; a sliding microtome ; a razor; a hone 
 and a good razor strop ; a paraffin bath and lamp ; a turn-table ; 
 a scalpel ; a pair of needles ; a pair of scissors ; a pair of for- 
 ceps ; stender dishes; minots or watch-glasses ; a wash bottle ; 
 a graduate (50 or 100 cc); pipettes; slides, 1X3 inches; round 
 covers, 18 mm. or ^ inch in diameter; and 
 square covers, 7^ inch. Long covers, 22 X 
 50 mm,, will be needed for some of the serial 
 sections. 
 
 A convenient and effective microscope 
 should have a rack and pinion coarse adjust- 
 ment, a fine adjustment, two eyepieces (about 
 one-inch and two-inch preferred), a low-power 
 objective of two-thirds of an inch or a one-inch 
 focus, a high-power objective of one-fifth or 
 one-sixth of an inch focus, a double nose- 
 piece, an iris diaphragm, and an Abbe con- 
 denser. A cheap and practical form is shown 
 in fig. /, and similar instruments are for sale 
 by all the leading companies. 
 
 The hand microtome {fig. 2) will be found 
 extremely useful, especially by the busy teacher 
 who has large classes. Any sliding microtome, if kept in good 
 order, will be sufficient for the work to be described in this book, 
 but those of medium size are to be preferred. The student's 
 microtome {figs, j and j A) is quite inexpensive and does good 
 
 Fig. 2. A convenient form 
 of hand microtome. 
 
Methods in Plant Histology 
 
 Fig. I. A compound microscope, with rack and pinion coarse adjustment, micrometer screw fine 
 adjustment, triple nosepiece, iris diaphragm, and condenser. 
 
Apparatus 
 
 work. Where ex- 
 pense is not an 
 objection the Jung 
 Thoma.the Minot, 
 and other micro- 
 tomes of similar 
 grade are to be 
 preferred. 
 
 The stout ra- 
 zors our grand- 
 fathers used to 
 shave with are ex- 
 cellent for free- 
 hand sectioning, 
 for hand-micro- 
 tome work, and 
 even for cutting 
 
 Fig. 3. The student's microtome. 
 
 Fig. 3 A . Clamp to hold an ordinary razor in the student's microtome. 
 
Methods ifi Pla?it Histology 
 
 paraffin sections on the sliding microtome. The blade should 
 be flat on one side [fig. 4 A). Modern razors [fig. 4 B) with 
 delicate blades, though good to shave with, are worthless for 
 cutting sections of plants. The razor is a necessity; if a micro- 
 tome knife is 
 wanted in addi- 
 tion, it should 
 have a bevel 
 about like that 
 shown in fig. 4 A. 
 F'G-4- Ashortblade.two 
 
 or three inches in length, is to be 
 preferred to the longer ones, which 
 are much more troublesome to 
 sharpen. 
 
 There are numerous forms of 
 the paraffin bath. Those with a 
 water-jacket, a thermometer, and a 
 thermostat to maintain an even 
 temperature are the most con- 
 venient where gas is available 
 
 Fig. 5. A paraffin bath, with water-jacket, 
 designed to be used with a thermostat. 
 
 Fig. 6. ^, top view; >B, sidev.ew; C", 
 tain the paraffin. 
 
 id view ; D, box to c 
 
 {fig. j). As a rule it 
 is easier to keep the 
 temperature constant 
 in the larger baths. 
 A bath which, if care- 
 fully watched, gives 
 the very best results 
 can be made by any 
 tinner, and is very in- 
 expensive. The ac- 
 companying figures 
 show the form and 
 dimensions [fig. 6). 
 
 It is made of cop- 
 per one thirty-second 
 
Apparatus 
 
 of an inch thick, but thicker copper is as good or better. There 
 should be two boxes to contain the paraffin ; the covers to the 
 boxes should fit loosely. Any kind of a lamp may be used. 
 
 Stender dishes are now very generally used for staining on 
 the slide. The form shown \xi fig. y A is made just large enough 
 to hold two slides, placed 
 back to back, and . hence 
 requires only a minimum of 
 the reagent. The cap in 
 this form does not fit closely 
 enough to keep absolute 
 alcohol and xylol, but does 
 very well for the other alco- 
 hols and stains. We do not 
 believe that the convex 
 cover is as good as a flat 
 one. The form shown in fig. y B is 
 hoi and xylol, but even with this 
 
 Fig. 7. A, Naples jar; Z>, stender dish. 
 
 the best for absolute alco- 
 it is better to put a little 
 vaseline or glycerine on the cover to prevent any evaporation. 
 
 Wide-mouthed 
 bottles, though 
 not so conveni- 
 ent, give just as 
 good results. 
 
 A serviceable 
 form of turn-table 
 for glycerine 
 mounts is shown 
 in fig. 8. 
 
 The other 
 pieces of appa- 
 ratus mentioned 
 need no com- 
 ment. By consulting a catalogue, which will be furnished by any 
 dealer, the beginner can determine what he needs to buy, and what 
 he can find substitutes for, if it is necessary to be very economical. 
 
CHAPTER II. 
 
 REAGENTS. 
 
 It would require entirely too much space even to enumerate 
 the reagents which are occasionally used in a fully equipped 
 university laboratory. The following list includes only those 
 which are used constantly. The Microtomist' s Vade-Mecum by 
 Lee contains very complete formulae for stains and other reagents. 
 The quantities mentioned below indicate about what the average 
 student uses in a three-months' course in methods. Nearly all 
 the stains, however, would last for a year, if properly used. 
 
 KILLING AND FIXING AGENTS. 
 Commercial alcohol (about 95 per cent.), 2 liters; absolute 
 alcohol, 300 cc; ether, 50 cc; chromic acid, 10 g.; corrosive 
 sublimate, 10 g.; glacial acetic acid, 25 cc; hydrochloric acid, 
 50 cc; picric acid, 5 g.; chloroform, 50 cc; [osmic acid, i per 
 cent, solution in water, 25 cc. This is extremely expensive, and 
 not necessary except for the most delicate work]. Formulae for 
 making killing and fixing agents from these materials will be 
 given later. 
 
 STAINS. 
 
 Only a few of the most important stains are given in this 
 list. In general one should have enough of a stain to stand 
 about two inches high in the stender dish or bottle in which the 
 staining is to be done. The theory and practice of staining will 
 be discussed later. 
 
 Delafield's Haematoxylin. — To 100 cc of a saturated solution of 
 ammonia alum add, drop by drop, a solution of i g. haematoxylin 
 dissolved in 6 cc. of absolute alcohol. Expose to air and light 
 for one week, then filter. Add 25 cc of glycerine and 25 cc of 
 methyl alcohol. Allow to stand until the color is rather dark. 
 Filter, and keep in a tightly stoppered bottle. The solution 
 
 7 
 
8 Methods m Plant Histology 
 
 should stand for two months before it is ready for use, but, if 
 needed immediately, the " ripening," which is brought about by 
 the oxidation of hsematoxylin into haematin, may be secured in 
 a few minutes by a judicious addition of peroxide of hydrogen, 
 
 Mayer's Haem-Alum. — Dissolve with gentle heat i g. of haema- 
 toxylin in 50 cc, of 95 per cent, alcohol; add a solution of 50 g. 
 of alum in a liter of distilled water. Allow the mixture to cool 
 and settle ; filter ; add a crystal of thymol to preserve from 
 mold. The stain is ready for use as soon as made, and it keeps 
 well. 
 
 Haidenhain's Iron Alum-Haematoxylin. — Two solutions are 
 used, and they are never to be mixed : 
 
 (a) A I ^ to 4 per cent, aqueous solution of ammonia sulphate of iron. 
 
 {b) A ^2 per cent, aqueous solution of haematoxylin. 
 
 Cyanin, Erythrosin, Safranin, Gentian Violet. — Numerous 
 formulae are given for these and other anilin stains, but the fol- 
 lowing general formula gives excellent results : 
 
 Make a 3 per cent, solution of anilin oil in distilled water; 
 shake thoroughly and frequently for a day; add enough alcohol 
 to make the whole mixture about 20 per cent, alcohol. Add i g. 
 of cyanin or erythrosin, etc., as the case may be, to 100 cc. of 
 the solution. Safranin is often used in a strong alcoholic solu- 
 tion, and even with the above formula it is better to dissolve the 
 safranin in strong alcohol before adding it to the mixture. 
 
 Acid Fuchsin. — Use a i to 2 per cent, solution in water, or 
 70 per cent, alcohol. 
 
 Iodine Green. — Use a i to 4 per cent, solution in water or 
 alcohol. A 3 per cent, solution in 70 per cent, alcohol is very 
 good for the vascular system of plants. 
 
 Mixtures of Fuchsin and Iodine Green. — The following formula 
 is often used for karyokinetic figures : 
 
 (a) A }i per cent, solution of fuchsin in water. 
 
 {b) A %, per cent, solution of iodine green in water. 
 
 Just before using mix {a) and (<^) in various proportions until 
 what is needed for the particular case is found. 
 Orange G. — Use a saturated aqueous solution. 
 
Reagetits 
 
 Eosin. — A I to 5 per cent, solution in water or alcohol. A 
 2 per cent, aqueous solution is good for material to be mounted 
 in glycerine, but a 2 per cent, solution in 70 per cent, alcohol is 
 better for balsam mounts. The stronger solution may be diluted 
 as needed for special cases. 
 
 FORMULAE FOR ALCOHOLS. 
 
 The grades of alcohol in most common use are 35 per cent., 
 50 per cent., 70 per cent., 85 per cent., 95 per cent., and 100 per 
 cent. The 100 per cent, is expensive, and great care should be 
 taken to keep the bottle well corked or the stender dish closely 
 covered. The following formulae will enable anyone to make 
 the other grades of alcohol from 95 per cent, alcohol and water: 
 95 35 95 50 95 70 95 85 
 
 60 45 25 
 
 The above are the formulae for 35 per cent., 50 per cent., 70 
 per cent., and 85 per cent, alcohol. Any other grade can be 
 gotten in the same way. In the first formula, subtract 35 from 
 95 ; the result, 60, is the number of cubic centimeters of water 
 which must be added to 35 cc. of 95 per cent, alcohol 
 in order to obtain 35 per cent, alcohol. The mixture 
 contains 95 cc. of 35 per cent, alcohol. If more or 
 less than 95 cc. of the mixture is needed, take 
 proportional parts of 35 and 60. This simple method 
 is a time-saver, but if the bottles or stender dishes 
 are to be filled frequently, it will be a still further 
 saving of time to use a long label [fig. 9) , and, after 
 pouring in the 95 per cent, alcohol, draw a line 
 showing how high it reaches, and then, after pouring 
 in the water, draw another line. The next time it is 
 necessary to fill the bottles merely pour in 95 per 
 cent, alcohol until it reaches the first line, and then pour in water 
 until it reaches the second line. It is not necessary to use dis- 
 tilled water, if pure drinking water is available. 
 
 Fig. 9. 
 
10 . Methods in Plant Histology 
 
 CLEARING AGENTS. 
 
 Xylol is the most generally useful clearing agent yet known. 
 Clove oil, cedar oil, bergamot oil, carbolic acid, and turpentine 
 are all necessary for special purposes. About 200 cc. of xylol, 
 50 cc. of clove oil, and 25 cc. of each of the others makes a fair 
 outfit to begin with. 
 
 MISCELLANEOUS. 
 
 Canada balsam, 25 cc; glycerine, 50CC.; glycerine jelly, 25 cc; 
 2 per cent, celloidin, 50 cc; 10 per cent, celloidin, 50 cc; hard 
 and soft paraffin, 500 g. each ; gold size, 25 cc; and a small soft 
 brush for ringing glycerine mounts. 
 
 ARRANGEMENT OF THE OUTFIT FOR STAINING AND MOUNTING. 
 
 It is best to keep the various reagents in definite positions in 
 order that no time may be lost in hunting for anything. The 
 
 UiJ^Vj (Uiyr\ U>^^ (h^Wj U%H«tJ (^.^nftU Ur^2\ 
 
 TcWOa.J (Orony&J L^\J^ \%<if^^^ /atiUU. J /f,TjtWuvj HUA-HootJ 
 UsiloU Moo J MS j (^85 J ^70^ (^50 j Cli\ 
 
 UL^ (%.\j^ 
 
 accompanying diagram {jig. 10) of a part of the top of a table 
 shows a convenient arrangement. 
 
 The alcohols are in front and the stains are placed behind. 
 The eosin, fuchsin, iodine green, cedar oil, and clove oil may be 
 kept in bottles; the rest should be in stender dishes. 
 
CHAPTER III. 
 
 TEMPORARY MOUNTS. 
 
 Before considering the complicated methods involved in 
 making permanent preparations a word should be said about 
 temporary mounts. A preliminary examination of almost any 
 botanical material may be made without any fixing, imbedding, 
 or staining. If a little starch be scraped from a potato, and a 
 small drop of water and a cover-glass be added, a very good 
 view will be obtained, and if a small drop of iodine solution be 
 allowed to run under the cover, the preparation, while it lasts, 
 could hardly be improved. The unicellular and filamentous 
 algae can be studied quite satisfactorily from such mounts. 
 The protonema of mosses and the prothallia of ferns should be 
 studied in this way, even if a later study from sections is 
 intended. If the top of a moss capsule be cut off at the level 
 of the annulus, a beautiful view of the peristome may be obtained 
 by simply mounting in a drop of water, or, in a case like this 
 where no collapse is to be anticipated, the object may be mounted 
 in a small drop of glycerine — just enough to come to the edge 
 of the cover without oozing out beyond — and the preparation 
 made permanent by sealing with gold size or any good cement. 
 The antheridia and archegonia of mosses may be examined if 
 the surrounding leaves are carefully teased away with needles. 
 Free-hand sectioning with a sharp razor and judicious teasing 
 with a pair of needles will give a fair insight into the anatomy 
 of the higher plants without demanding any further knowledge 
 of technique. This rough work is a very desirable antecedent to 
 the study of microtome sections, because most students see in a 
 series of microtome sections only a series of sections when, in 
 the mind's eye, they ought to see the object building itself up in 
 length, breadth, and thickness as they pass from one section to 
 another. 
 
 II 
 
CHAPTER IV. 
 
 THE GENERAL METHOD. 
 
 We shall now consider the routine of mounting an object in 
 Canada balsam. While the outline refers more particularly to 
 the paraffin method, the principles are general in their applica- 
 tion and must be mastered by everyone who desires to make 
 first-class preparations. Several of the topics, like killing and 
 fixing, staining, etc., will be treated in detail when considering 
 the various reagents. 
 
 I. KILLING AND FIXING. 
 
 Usually the same reagent is used for both killing and 
 fixing. The purpose of a killing agent is to bring the life- 
 processes to a sudden termination, while a fixing agent is used 
 to fix the cells and their contents in as nearly the living condi- 
 tion as possible. The fixing consists in so hardening the 
 material that the various elements may retain their natural con- 
 dition during all the processes which are to follow. This step 
 is one of extreme importance. Take the killing and fixing 
 fluids into the field. If one waits until the material is brought 
 to the laboratory, there may be some fixing, but it will, 
 in many cases, be too late to do much killing. Material like 
 Spirogyra, however, may be brought from the field into the 
 laboratory before fixing, if considerable water be brought with 
 it. Branches with developing buds may be brought in and kept 
 in water. Always have the material in very small pieces, in 
 order that the reagents may act quickly on all parts of the 
 specimens. Pieces larger than one-fourth inch cubes shpuld be 
 avoided whenever possible. For very fine work no part of the 
 specimen should require the reagent to penetrate more than 
 one-sixteenth of an inch. In general, the volume of the reagent 
 should be ten to fifty times that of the material. The time 
 
 13 
 
14 Methods in Plant Histology 
 
 required for this process varies with the reagent, the character 
 of the tissue, and the size of the piece. About twenty-four 
 hours is a commonly recommended period for chromic acid 
 solutions. While this might suffice in some cases, we should 
 recommend two or three days, and even a longer period would 
 probably do no harm. 
 
 II. WASHING. 
 
 Nearly all fixing agents, except the alcohols, must be 
 washed out from the material as completely as possible before 
 any further steps are taken, because some reagents leave annoy- 
 ing precipitates which must be removed, and others interfere 
 with subsequent processes. Aqueous fixing agents with chromic 
 acid as their principal ingredient are washed out with water ; 
 aqueous solutions of corrosive sublimate are also washed out 
 with water ; but alcoholic solutions should be washed out with 
 alcohol of about the same strength as the fixing agent ; picric 
 acid, or fixing agents with picric acid as an ingredient, must not 
 be washed out with water, but with alcohol, whether the picric 
 acid be in aqueous or alcoholic solution. Running water is best, 
 and where this is not convenient the water should at least be 
 changed quite frequently. The washing-out process usually 
 takes from twelve to twenty-four hours, but it can be shortened 
 about one-half by keeping the fluid lukewarm. 
 
 III. HARDENING AND DEHYDRATING. 
 
 After the material has been washed, it is necessary to con- 
 tinue the hardening and also to remove the water. Alcohol is 
 used almost entirely for these purposes. It completes the 
 hardening and at the same time dehydrates, that is, it replaces 
 the water in the material, an extremely important considera- 
 tion, for the least trace of moisture, a trace so slight as to be 
 almost imaginary, is nevertheless sufficient to make a prepara- 
 tion poor or indifferent when it might have been excellent. 
 
 The process of hardening and dehydrating must be gradual. 
 If the material should be transferred directly from water to 
 absolute alcohol, the hardening and dehydrating would be 
 
General Method 15 
 
 brought about in a very short time, but the violent osmosis 
 would cause a ruinous contraction of the more delicate parts. 
 Therefore, transfer from water to 35 per cent, alcohol, which 
 should act for six to twenty-four hours. Then use 50 per cent. 
 for; a similar period. Material may now be placed in 70 per 
 cent, alcohol, where it may remain until ready for use, since 70 
 per cent, alcohol is a good preservative. Various devices, like 
 constant drips and osmotic apparatus, have been proposed to 
 secure a more gradual transfer. Whether these have any real 
 advantages still remains to be proved. The writer has taken 
 well-fixed fern prothallia through the series 35 per cent., 50 per 
 cent., 70 per cent., without the slightest plasmolysis. Such 
 things as fern prothallia, filamentous algae, etc., can be watched 
 under the microscope as the transfer is made, and, if plasmolysis 
 results, the series of alcohols may be made closer, e.g., 10 per 
 cent., 20 per cent., 30 per cent., etc. It is said that material 
 left for some time in 70 per cent, alcohol will shrink in spite of 
 good killing and fixing, and it is also claimed that its capacity 
 for staining is diminished. Some recommend that glycerine be 
 added to the alcohol; others prefer to complete the dehydrating 
 process and leave the material in an essential oil ; while still 
 others would imbed it and keep it in paraffin. The last is doubt- 
 less best of all, but requires such an immense amount of labor 
 that it is impracticable for general purposes. Nearly all of our 
 own material which is not needed for immediate use is in 70 
 per cent, alcohol, unless, of course, the material has been put 
 into formalin or some such reagent which kills, fixes, and pre- 
 serves all at once. 
 
 After the 70 per cent, alcohol, use 85 per cent, and 95 
 per cent., allowing six to twenty-four hours for each. Then 
 use 100 per cent, alcohol for one or two days. The 70 per 
 cent, would probably complete all the hardening which is neces- 
 sary, but the other three must be used to complete the removal 
 of water. 
 
 Up to this point the processes are exactly the same, whether 
 the material is to be imbedded in paraffin or celloidin. 
 
1 6 Methods in Pla?it Histology 
 
 IV. CLEARING. 
 
 Let us suppose that the material has been thoroughly 
 dehydrated, so that not the slightest trace of water remains. If 
 the supposition chances to be contrary to fact, all the work 
 which has preceded, as well as all which is to follow, is only an 
 idle waste of time. The purpose of a clearing agent is to make 
 the tissues transparent, but clearing agents also replace the 
 alcohol. At this stage the latter process is the essential one, 
 the clearing which accompanies it being incidental. The clear- 
 ing, however, is very convenient, since it shows that the alcohol 
 has been replaced and that the material is ready for the next step. 
 
 Various clearing agents are in use. Xylol is the most gen- 
 erally employed, and for most purposes it seems to be the 
 best. Bergamot oil, cedar oil, clove oil, turpentine, and chloro- 
 form are all necessary for special purposes. 
 
 The transfer from absolute alcohol to the clearing agent 
 should he gradual, like the hardening and dehydrating processes. 
 The following is a good method : 
 
 3 parts loo per cent, alcohol and i part xylol, i to lo hours. 
 2 parts 100 per cent, alcohol and 2 parts xylol, i to lo hours. 
 I part I GO per cent, alcohol and 3 parts xylol, i to 10 hours. 
 
 This transfer is best accomplished by adding the xylol to the 
 alcohol. It is not necessary to measure, since anyone can guess 
 with suflficient accuracy. Shake gently each time the xylol is 
 added. Pour off the mixture and add pure xylol, which should 
 cause the material to become transparent. This may require 
 only a moment, but may require hours. Other clearing agents 
 may be used in the same way instead of the xylol. 
 
 V. THE TRANSFER FROM CLEARING AGENT TO PARAFFIN. 
 This should also be a gradual process. The most con- 
 venient method is to place a small block of parafifin in the pure 
 clearing agent with the material. The parafifin dissolves gradu- 
 ally and produces the same result as if a small shaving of 
 paraffin had been added every few minutes for a day or so. 
 During this process the bottle or dish should be kept lukewarm. 
 Six to ten hours, or over night, is usually sufficient for this step, 
 
General Method ly 
 
 although it would seem that material may be kept here for a 
 much longer time without injury, and in case of such refractory 
 material as the megaspores of the heterosporous pteridophytes 
 four or five days seem necessary. Excellent preparations of the 
 embryo-sac of Aster have been made from material which had 
 remained in the xylol and paraffin for nearly three years. No 
 more paraffin should be added than will go into perfect solution. 
 The temperature may be gradually increased, so that a much 
 greater amount of paraffin will go into solution, but the paraffin 
 must not be allowed to crystallize. 
 
 VI. THE PARAFFIN BATH. 
 
 This step is usually called infiltration, but when the trans- 
 fer from the clearing fluid to paraffin is made gradually, as has 
 just been indicated, the process of infiltration is already begun. 
 It is now necessary to get rid of the xylol or other clearing 
 agent. This may be done by simply pouring off the mixture of 
 xylol and paraffin and replacing it with pure melted paraffin. 
 The bath should be kept at a temperature about i° C. higher 
 than the melting-point of the paraffin. Fifty-three degrees C. 
 is a good temperature for general purposes, but this may be 
 reduced from i° to 3° C. in winter, and must often be raised 
 in summer. For special purposes it is sometimes necessary to 
 use a temperature as high as 70° C. If the xylol or other 
 clearing agent is not thoroughly removed, the paraffin will be 
 granular or mealy, and will not cut well. The paraffin should be 
 changed once or twice to make sure that the clearing agent 
 is all removed. Do not waste this paraffin, for the clearing 
 agent can be driven off by prolonged heating, and the paraffin 
 will be better than ever. Most people use soft paraffin (about 
 45° C.) for the first half of the time necessary for infiltration, 
 and a harder grade (48° C. to 54° C.) for the latter part of the 
 process. This is a good plan, for soft paraffin melts at a lower 
 temperature, and it is always well to minimize heat. 
 
 Some think that it is better not to pour off the mixture of the 
 clearing agent and paraffin, but rather to evaporate the clearing 
 
1 8 Methods i7i Plant Histology 
 
 agent by keeping the temperature just high enough to prevent 
 the crystallization of the paraffin. This method has certainly 
 given good results with small, delicate objects. 
 
 The time required for infiltration varies with the character 
 of the tissue and the size of the piece. Few things can be well 
 infiltrated in less than an hour. Lily ovaries require one to four 
 hours ; heads of Aster at the fertilization period, six to twelve 
 hours. Some claim that even delicate objects, like fern pro- 
 thallia and the filamentous algae and fungi, are not injured by 
 a bath of several days, if care be taken not to let the tempera- 
 ture rise above 48° C. to 50° C. No one seems to know how 
 long a certain object should remain in the bath, but many 
 competent investigators are now using more prolonged periods. 
 
 VII. IMBEDDING. 
 
 Material may be imbedded in paper trays, watch crystals, 
 or in apparatus made for the purpose. Imbedding L's consist- 
 ing of two L-shaped pieces {^fig. 11) of brass, 
 type metal, or lead are very convenient. 
 We use a pair-of L-shaped pieces with arms 
 three inches long. These furnish a box of 
 almost any required size. A piece of glass 
 serves for a bottom. The tray, minot, or 
 whatever is used, should be slightly smeared 
 with glycerine, to prevent sticking. If several objects are to be 
 imbedded in one dish, it is best to have the dish as near the 
 temperature of melted paraffin as possible ; otherwise the objects 
 may stick to the bottom, and it will be impossible to arrange 
 them properly. Great care should be taken, however, not to 
 have the dish too hot, since too high a temperature not only 
 injures the material, but also prevents a thorough imbedding. 
 Pour the paraffin with the objects into the imbedding dish and 
 cool as rapidly as possible. If paraffin cools slowly, it crystal- 
 lizes and does not cut well. The layer of paraffin should be 
 just thick enough to cover the objects, not only as a matter of 
 economy, but because a thick layer retards the cooling. Very 
 
General Method 
 
 19 
 
 small objects, like the megaspores of Marsilea, ovules of Silphium, 
 etc., may simply be poured out upon a cool piece of glass. In 
 this way very thin cakes are made, which harden very i^apidly. 
 
 VIII. CUTTING. 
 Twenty minutes after an object is imbedded it is ready for 
 cutting. Trim the paraffin containing the object into a conveni- 
 ent shape, and fasten it upon a block of wood. Blocks of pine 
 three-fourths of an inch long and three-eighths of an inch square 
 are good for general purposes. Put paraf^n on the end of the 
 block so as to form a firm cap about one-eighth of an inch thick. 
 Warm the cap and the bottom of the piece containing the object, 
 and press them lightly together ; then touch the joint with a hot 
 needle, put the whole thing into cold water for a minute, and it 
 is ready for cutting. Cutting can be learned only by experience, 
 but a few hints may not come amiss : (^) Keep the knife sharp. 
 If expense is not too serious an objection, it is well to have two 
 hones, one rather soft, for use when the knife is dull, and the 
 other quite hard, for putting on an even edge. Flood the stone 
 with water, and rub it with the small slip which accompanies all 
 high-grade hones ; this not only makes a lather which facilitates 
 the sharpening, but it also keeps the surface of the hone flat. 
 As soon as the edge of the knife appears smooth and even 
 under a magnification of thirty or forty diameters, the sharpen- 
 ing is completed with a good strop. It is better to sharpen the 
 knife every time you use it. {b) Keep the microtome well oiled 
 and clea?i. {c) Trim the block so that each section shall be a 
 perfect rectangle. 
 
 B 
 Fig. 12. 
 
20 Methods in Plant Histology 
 
 A ribbon of sections like that shown in fig. 12 A is much 
 better than one like B of the same figure, because sections will 
 usually come off in neater ribbons if the knife strikes the longer 
 edge of the rectangle, so that the sections are united by their 
 longer sides rather than by the shorter. Crooked ribbons are 
 caused by wedge-shaped sections, and are always to be avoided, 
 because they make it difficult to economize space, and also 
 because they present such a disorderly appearance. The knife, 
 which should be placed at a right angle to the block and not 
 obliquely, should strike the whole edge of the block at once, and 
 should leave in the same manner. 
 
 IX. FIXING SECTIONS TO THE SLIDE. 
 
 Sections must be firmly fixed to the slide, or they will be 
 washed off during the processes involved in staining. Mayer's 
 albumen fixative is excellent for this purpose. Formula : 
 White of egg, 50 cc. (Active principle.) 
 Glycerine, 50 cc. (To keep it from drying up.) 
 Salycilate of soda, i g. (Antiseptic, to keep out bacteria, etc.) 
 
 Shake well and filter. It will keep from two to six months. 
 The fixative may be used alone or in connection with the water 
 method. Put a small drop of fixative on the slide, smear it 
 evenly over the surface, and then wipe it off with a clean finger 
 or piece of linen until only a scarcely perceptible film remains ; 
 then add several drops of distilled water and float the sections 
 or ribbons on the water. Warm gently until the paraffin 
 becomes smooth and free from wrinkles. Be careful not to 
 melt the parafifin, for the albumen of the fixative coagulates 
 with less heat than is required to melt the paraffin. It is a very 
 good plan to put the slide on the top of the water bath for a 
 moment and then, after the sections have become smooth, 
 remove the surplus water and leave them on the bath with a 
 couple of thicknesses of blotting paper under them for three or 
 four hours, or, better, over night. If the fixative is used alone, 
 as is often the case when sections are very thick, none of this 
 delay is necessary, since the sections are merely laid upon the 
 fixative and pressed down gently with the finger. 
 
Ge7ieral Method 2i 
 
 X. REMOVAL OF THE PARAFFIN. 
 
 To remove the paraffin it is very customary to heat the 
 slide gently until the paraffin melts, and then place the slide in 
 xylol for thirty seconds or a minute. I believe, however, that 
 it is far better merely to warm the slide a little (not warmer than 
 30° C). Good xylol will then remove the paraffin in two or 
 three minutes. Even if the slide should not be warmed at all, 
 good xylol should remove the paraffin in five minutes. 
 
 XI. REMOVAL OF XYLOL. 
 The xylol may be removed either by absolute alcohol 
 or by the 95 per cent. The absolute alcohol does not seem to 
 be really necessary. For my own work I have two stender 
 dishes of xylol and two of absolute alcohol. After the xylol 
 has been used for a time I employ it only for removing paraffin, 
 and in the same way use the absolute alcohol only for removing 
 the xylol, while the other two dishes are used only for dehydrat- 
 ing and clearing. In transferring from xylol to absolute alcohol, 
 or vice versa, it is well to drain off the superfluous liquid b^ 
 resting the corner of the slide upon a piece of blotting paper. 
 These rather expensive reagents will last much longer when this 
 precaution is taken. 
 
 XII. TRANSFER TO THE STAIN. 
 
 Stains are aqueous or alcoholic, and alcoholic stains are 
 of various strengths. If an aqueous stain be used, the slide 
 should be passed successively through the alcohols 95 per cent., 
 85 per cent., 70 per cent., 50 per cent., and 35 per cent., allow- 
 ing each to act for about thirty seconds, after which the slide is 
 put into the stain. (In many cases it is sufficient to put a few 
 drops of the stain on the slide with a pipette.) From the stain 
 the slide is passed back through the various grades of alcohol, 
 allowing it to remain about thirty seconds in each as before. If 
 the stain be alcoholic of about 70 per cent, strength, the process 
 is somewhat shorter, for the slide goes into the stain from 70 
 per cent, alcohol, and goes back into 70 per cent, alcohol from 
 the stain. The rule is to transfer to the stain from the alcohol 
 
22 Methods in Plant Histology 
 
 which is nearest the strength of the stain. These directions are 
 based upon a rather wide experience and certainly give excellent 
 results with stains which do not forbid their application. How- 
 ever, evidence has long been accumulating which indicates that 
 in case of very thin sections (sections so thin that no whole cells 
 are included) only the absolute alcohol is necessary and that 
 the rest may be omitted. Whether it is better to omit the 
 alcohols from 35 per cent, to 95 per cent, when staining on the 
 slide is a question still waiting for a definite answer, although 
 careful and systematic experimenting would soon settle it. 
 Even if it should be shown that it is as well or better to omit 
 the alcohols from 35 per cent, to 95. per cent, in case of thin 
 sections, it must not.be inferred that these alcohols maybe 
 omitted in preparing a piece of epidermis stripped from a leaf, 
 for, in this case, a ruinous plasmolysis occurs, and in sections 
 thick enough to include whole cells the same damage is done. 
 In the directions given in this book it is assumed that the series 
 35 per cent., 50 per cent., 70 per cent., 85 per cent., 95 per cent., 
 and 100 per cent, is to be used, unless otherwise mentioned. 
 XIII. CLEARING. 
 
 After the sections have been stained, passed back through 
 the various grades of alcohol, and have been thoroughly dehy- 
 drated in absolute alcohol, they are cleared or made transparent 
 by means of xylol or some other clearing agent. The clearing 
 agent must be a solvent of balsam. From thirty seconds to five 
 minutes will be sufficient for clearing any kind of sections. 
 XIV. MOUNTING IN BALSAM. 
 
 After the sections are cleared, wipe the slide on the side 
 which does riot bear the sections. Put on a drop of Canada 
 balsam and add a clean,* thin cover. Before the cover is put 
 on, pass it through the flame of an alcohol lamp to remove 
 moisture, for it would be a pity indeed to injure a preparation 
 
 'Slides and covers should be treated with hydrochloric acid, or equal parts of 
 bydrochloric acid and water, for several hours. They should then be thoroughly rinsed 
 in water and placed in 95 per cent, alcohol. They should be wiped with a cloth per- 
 fectly free from lint. 
 
Geiieral Method 23 
 
 at this stage of the process. Add a label, and the mount is 
 complete. 
 
 A TENTATIVE SCHEDULE FOR PARAFFIN SECTIONS. 
 
 It will be useful to give several tentative schedules for the 
 use of beginners. It cannot be too strenuously insisted that 
 these schedules are only tentative, their sole object being to give the 
 beginner a start. The following is a tentative schedule for the 
 ovary of a lily at any period before fertilization. The pieces 
 should not be more than half an inch in length. 
 
 1. Chromo-acetic acid, 2 days. 
 
 2. Wash in water, i day. 
 
 3. Thirty-five per cent., 50 per cent., 70 per cent., 85 per cent., 95 per cent, 
 alcohol, 6 to 24 hours each, as convenient. 
 
 4. One hundred per cent, alcohol, 24 hours. This should be changed once 
 or twice. The volume should be at least ten times that of the material. 
 
 5. Transfer from absolute alcohol to xylol, allowing at least 2 hours in each 
 of the three mixtures, and 2 hours in pure xylol. 
 
 6. Add paraffin to the xylol and keep warm for 12 to 24 hours. 
 
 7. Melted paraffin in the bath, 2 to 24 hours, as convenient. The paraffin 
 should be changed once or twice. 
 
 8. Imbed. 
 
 9. Section; about lOyu is a good thickness, 
 
 10. Fasten to the slide. 
 
 11. Dissolve off the paraffin in xylol. 
 
 12. One hundred per cent., 95 per cent., 85 per cent., 70 per cent., 50 per 
 cent, alcohol, 30 seconds each. An hour in each would do no damage 
 if other duties should interfere, 
 
 13. Delafield's hematoxylin, 10 minutes, 
 
 14. Rinse in water, 5 minutes. A couple of hours does no harm. 
 
 15. Thirty-five per cent., 50 per cent., and 70 per cent, alcohol, 30 seconds 
 each. An hour would do no harm, 
 
 16. Acid alcohol, i second, 
 
 17. Seventy per cent, alcohol, i minute. An hour would do no harm. 
 
 18. Erythrosin, 30 seconds to i minute. 
 
 19. Eighty-five per cent., 95 per cent., 100 per cent., about 5 seconds each. 
 This step must not be too prolonged, lest the erythrosin wash out. 
 
 20. Xylol, at least i minute. The slide may be left here for an hour without 
 injury. 
 
 21. M.)unt in balsam. 
 
24 Methods in Plant Histology 
 
 On the whole, it is not a good plan to use protracted periods 
 in the processes from No. 1 1 to No. 20 inclusive, because there 
 is danger that a prolonged soaking may loosen some of the more 
 delicate contents of the cells, even if whole sections do not 
 come off bodily. 
 
CHAPTER V. 
 
 KILLING AND FIXING AGENTS. 
 
 In this short account only the reagents which are at present 
 considered most valuable for botanical work will be considered. 
 Probably no process in microtechnique is in more urgent need 
 of improvement than this first step of killing and fixing. Nearly 
 all of our formulae are merely empirical, for very few botanists 
 are expert chemists, and those who have the requisite knowl- 
 edge of chemistry are interested in physiological problems 
 rather than in microtechnique. The principal ingredients of the 
 usual killing and fixing agents are : alcohol, chloroform, chromic 
 acid, acetic acid, osmic acid, formic acid, picric acid, sulphuric 
 acid, platinum chloride, iridium chloride, corrosive sublimate, 
 and formalin. We shall consider first 
 
 THE ALCOHOLS. 
 
 a. Ninety-five per cent. Alcohol. — This is in quite general use 
 for material which is needed only for rough work. It is extremely 
 convenient, since it kills, fixes, and preserves at the same time 
 and needs no changing or washing. It really has nothing to 
 recommend it for fine work. It causes protoplasm to shrink, but 
 cell walls usually retain their position, so that 95 per cent, alco- 
 hol will do for free-hand sections of wood and many herbaceous 
 stems ; but even free-hand sections of tender stems, like young 
 geraniums and begonias, will look better if better reagents are 
 employed. Alcohols weaker than 95 per cent, are not to be 
 recommended as fixing agents, although 70 per cent, alcohol, or 
 even 50 per cent., will preserve material for habit work. 
 
 b. Absolute (100 per cent.) Alcohol. — This is a fair killing and 
 fixing agent, but is rather expensive. It causes but little shrink- 
 ing of the protoplasm, and is a time-saver if material is to be 
 imbedded in paraffin. With 95 per cent, or with absolute 
 
 25 
 
26 Methods in Plant Histology 
 
 alcohol, objects are generally left in the reagent until needed for 
 use, but such material becomes very brittle. The addition of 
 glycerine is an improvement, if material is to be kept long. 
 Acetic acid has been used with alcohols to counteract the 
 tendency to shrink. One of the most successful of the alcohol 
 combinations is 
 
 c. Carnoy's Fluid. — 
 
 Absolute alcohol, 6 parts. 
 
 Chloroform, 3 parts. 
 
 Glacial acetic acid, i part. 
 The penetration of the reagent is excellent, and only a few 
 hours are needed for fixing. Material should be washed in 
 absolute alcohol (perhaps 95 per cent, alcohol would do no 
 harm) until there is no odor of acetic acid. This should not 
 require more than one or two hours. It is better to imbed in 
 paraffin at once, but when this is not convenient the material 
 may be transferred to 85 per cent, alcohol and then to 70 per 
 cent., where it maybe left until needed. Cyanin and erythrosin, 
 fuchsin and iodine green, and similar combinations, give particu- 
 larly brilliant staining after this reagent. 
 
 THE CHROMIC-ACID GROUP. 
 
 Chromic acid, or solutions with chromic acid as a foundation, 
 are the most generally useful killing and fixing agents yet known 
 to the botanist. A i per cent, solution of chromic acid in water 
 gives good results, but it is better to use the chromic in connec- 
 tion with other ingredients, such as acetic acid, formic acid, 
 osmic acid, etc. The proportions of the various ingredients, 
 for the present at least, must be determined by experiment. 
 With favorable objects like fern prothailia, Spirogyra, and other 
 things which can be watched while the fixing is taking place, 
 suitable proportions are rather easily determined, because speci- 
 mens, after being placed in the reagent, may be examined at 
 frequent intervals, and combinations which cause plasmolysis 
 may be rejected and different proportions tried until satisfactory 
 results are secured. For example, fern prothailia might be placed 
 in the following solution : chromic acid, 2 g. ; acetic acid, i cc; 
 
Killing and Fixi?ig Agents 27 
 
 and water, 97 cc. If plasmolysis takes place, weaken the chromic 
 or strengthen the acetic, since the chromic has a tendency to 
 produce contraction, and the acetic to cause swelling. Too 
 large a proportion of acetic acid, however, may cause distortion, 
 and hence it would be better to weaken the chromic. In case 
 of fern prothallia, 3 parts chromic, i part acetic, and 396 parts 
 water will cause practically no plasmolysis, and the fixing is 
 sufficiently thorough to permit imbedding in parafifin. A com- 
 bination may be quite satisfactory for fern prothallia and still 
 fail to give good results with Spirogyra, and a combination which 
 succeeds very well with Spirogyra may not succeed at all with Vau- 
 cheria. For very critical work the most favorable proportions 
 must be determined for the particular plant under investigation. 
 When the effect of the reagent cannot be observed directly, it is 
 well to make a free-hand section and thus determine whether 
 plasmolysis takes place. It is not safe to judge the action of a 
 fixing agent by the appearance of sections cut from material 
 which has been imbedded in parafifin, because shrinking of the 
 cell contents often takes place during the transfer from absolute 
 alcohol to the clearing agent or during infiltration with paraffin, 
 and sometimes during even later processes. When in doubt as 
 to proportions, we should suggest 2 cc. chromic acid, 2 cc. acetic 
 acid, and 296 cc. water as a good formula for most purposes. 
 
 The time required for fixing undoubtedly varies with different 
 plants, but twenty-four hours may be considered a minimum 
 even for the most delicate objects. It is well known that zoolo- 
 gists allow fixing agents like Miiller's fluid and Erlicki's fluid to 
 act for weeks before the material is passed on to the next 
 stage, and it may well be questioned whether botanists have 
 not made a mistake in allowing the chromic solutions to act for 
 so short a time. At present most botanists recommend sixteen 
 to twenty-four hours for material which is to be imbedded in 
 parafifin, but some recent experiments in my laboratory indicate 
 that material which has been in the fixing fluid for two or three 
 days is better able to withstand the subsequent processes. More 
 rapid penetration, and consequently more immediate killing, can 
 
28 Methods in Plant Histology 
 
 be secured if the reagent is kept warm (30° to 40° C). Since 
 chromic acid has a tendency to render objects hard and brittle, 
 it is often better to use some other fixing agent, if much diffi- 
 culty is anticipated in the cutting. 
 
 After fixing is complete, the reagent should be washed out 
 with water. Running water is desirable, and where this is not 
 convenient the water must be changed frequently. Any material 
 should be sufficiently washed in six to twenty-four hours, but 
 the time may be shortened about one-half by using lukewarm 
 water. 
 
 Some of the formulae are as follows : 
 
 a. Strong Chromo-Acetic Solution. — 
 
 I g. chromic acid. 
 
 I cc. glacial acetic acid. 
 
 98 cc. water. 
 
 This solution has been used quite extensively in embryo- 
 logical work upon the higher plants. 
 
 b. "Weak Chromo-Acetic Solution. — (Schaffner's formula): 
 
 0.3 g. chromic acid. 
 0.7 acetic acid. 
 
 99 cc. water. 
 
 This has also been used in embryological work. It causes 
 little or no plasmolysis, but the chromic seems rather weak. 
 Difficult material, like Aster heads and ripe Capsella pods, cuts 
 more readily after this reagent than after the stronger solution. 
 
 c. Medium Chromo-Acetic Solution. — 
 
 0.7 g. chromic acid. 
 
 0.5 cc. glacial acetic acid. 
 
 100 cc. water. 
 
 We are now using this solution, and it seems very successful 
 in nearly all cases. 
 
 d. Flemming's Fluid. — (Weaker solution.) 
 
 ( I per cent, chromic acid, 25 cc. 
 
 A. \ ^ P^^ cent, acetic acid, 10 cc. 
 / Water, 55 cc. 
 
 B. I per cent, osmic acid, 10 cc. 
 
Killing and Fixing Agents 29 
 
 Keep the mixture A made up, and add B as the reagent is 
 needed for use, since it does not keep well. This fluid is quite 
 expensive on account of the osmic acid. For cytological work 
 it gives as good results as any fixing agent which has yet been 
 thoroughly tested. It is especially recommended for chromo- 
 somes, centrosomes, achromatic structures, and mitotic phe- 
 nomena in general. Material should be in very small pieces 
 one-eighth of an inch square, or in thin slices one-eighth of an 
 inch or less in thickness, for the fluid penetrates poorly. The 
 blackening due to the osmic acid may be removed by peroxide 
 of hydrogen just before the slide is passed from the alcohol into 
 the stain. Flemming's safranin-gentian violet-orange combina- 
 tion gives its most brilliant results after this reagent. It seems 
 possible that the traditional superiority of this reagent has been 
 overestimated. 
 
 e. Merkel's Fluid. — 
 
 Equal volumes of a 1.4 per cent, solution of chromic acid and a 1.4 per 
 cent, solution of platinic chloride. 
 
 This is also an expensive reagent. It is recommended for 
 mitotic phenomena, but does not seem to equal Flemming's 
 solution. 
 
 /. Hermann's Fluid. — 
 
 1 per cent, platinic chloride, 15 parts. 
 Glacial acetic acid, i part. 
 
 2 per cent, osmic acid, 4 or 2 parts. 
 
 This is the most expensive fixing agent yet discovered, and 
 for botanical purposes it does not seem to be any better than the 
 cheaper chromic mixtures. It is mentioned here with chromic 
 mixtures because it originated as a variation of Flehiming's 
 fluid, the platinic chloride being substituted for the chromic 
 acid. 
 
 According to Lee, the chief objection to all mixtures into 
 which chromic acid enters is that "it precipitates certain of the 
 liquid albuminoids of the tissues in the form of filaments or net- 
 work, which are often of great regularity and simulate structural 
 elements of the tissues." Nevertheless, the mixtures which have 
 
30 Methods in Plant Histology 
 
 just been described are the best which have yet been thoroughly 
 tested. If material killed in any of the above mixtures is not 
 well washed, the haematoxylins will not stain. It is claimed that 
 the anilins will stain in spite of poor washing, but it is a question 
 whether such preparations are as permanent as those from well- 
 washed material. 
 
 PICRIC ACID. 
 
 Use a saturated solution in water or 70 per cent, alcohol. 
 One gram of picric acid crystals will saturate about 75 cc. of 
 water or alcohol. This reagent penetrates well and does not 
 make the material brittle. It is to be recommended when diffi- 
 culty is anticipated in the cutting. If used cold, the time varies 
 from one to twenty-four hours, depending upon the character of 
 the tissue and size of the specimen. If used hot (85° C), five 
 or ten minutes will be sufficient. Material should be washed in 
 70 or 50 per cent, alcohol. Water is injurious, and some even 
 go so far as to avoid aqueous stains, unless the material has been 
 thoroughly washed. The washing should be continued until the 
 material appears whitish and the alcohol no longer becomes 
 tinged with yellow. Picro-carmine gives its best results after 
 this reagent. Picric acid can be combined with various other 
 fixing agents, and so we have picro-sulphuric acid, picro-nitric 
 acid, picro-chromic acid, picro-chromic-sulphuric acid, and picro- 
 
 osmic acid. 
 
 CORROSIVE SUBLIMATE. 
 
 Use a 2 to 5 per cent, solution in water, or 70 per cent, alco- 
 hol. The addition of about i cc. of glacial acetic acid to lOO cc. 
 of this solution is certainly an improvement. The time required 
 is considerably shorter than for chromic solutions. From one to 
 ten hours will be found to be sufficient. If used hot (85° C), 
 only five or ten minutes is required. Washing may be done 
 with water, but 50 per cent, alcohol is better. If a few drops of 
 an iodine solution be added to the alcohol, the alcohol takes on 
 a brownish color, but soon clears up. If the addition of iodine 
 be continued, the washing is complete when the alcohol no 
 longer clears up, but retains the brown color. If the washing is 
 
Killing and Fixi?ig Agents 31 
 
 incomplete, crystals of corrosive sublimate will be unduly con- 
 spicuous in preparations made from the material. Camphor may 
 be used instead of iodine to hasten the washing. 
 
 The carmines are very brilliant after corrosive sublimate on 
 account of the formation of mercuric carminate, but haematoxy- 
 lin and anilines also give good results. It is claimed, however, 
 that achromatic structures do not stain well with the safranin-gen- 
 tian violet-orange combination after this fixing agent, but it 
 might be worth while to test other stains before discarding this 
 reagent for cytological work. 
 
 Corrosive sublimate material gets very brittle if allowed to 
 remain long in alcohol^ and therefore it is better to imbed it as 
 soon as possible. 
 
 FORMALIN. 
 
 Formalin is a comparatively recent addition to the list of 
 killing and fixing agents. It is an excellent preservative, often 
 preserving the blue and red colors as well as the structure of 
 objects. A 2 or 4 per cent, solution in water is good for fila- 
 mentous algs. The material may simply be put into the reagent 
 and left until needed for use. For class use formalin material 
 should be washed in water for several minutes, because the 
 fumes are irritating to the eyes and mucous membranes. After 
 a thorough washing in water any of the usual stains may be 
 used. 
 
 GENERAL HINTS ON FIXING. 
 
 It is very desirable that the fixing agent should penetrate 
 quickly to all parts of the object. For this reason material 
 should be in small pieces. The best fixing agents do their best 
 work near the surface of the piece. Small objects like Azolla 
 and fern prothallia may be thrown into the fixing agent entire ; 
 even larger objects, which, like the anthers of Lilium, are easily 
 penetrated, may also be put in without any cutting. Most 
 objects larger than quarter of an inch cubes should be trimmed 
 with a sharp knife or razor ; some knowledge of the structures 
 concerned is essential before one can trim material with unvary- 
 ing success. 
 
32 Methods in Plant Histology 
 
 Some objects, although small, cause trouble in various ways. 
 Many buds are hairy and will not sink ; if such things are dipped 
 quickly in strong alcohol, they will usually sink. If rather large 
 air bubbles prevent the material from sinking, as in case of peri- 
 chaetical leaves of some mosses and involucral leaves of liver- 
 worts, a little dissection or a careful snip with the scissors will 
 obviate the difficulty. If an air-pump is available, the bubbles 
 are easily removed. 
 
 It is often asked whether fixing agents really preserve the 
 actual structure of cell contents. It must be admitted that 
 some things, notably the liquid albuminoids, are much modified 
 in appearance, but the most competent observers are now 
 inclined to believe that such delicate objects as chromosomes, 
 centrosomes, the achromatic figure, and even the structure of 
 protoplasm, can be studied with confidence from material which 
 has been fixed, imbedded, and stained. Recent study of these 
 objects in the living condition has strengthened this confidence. 
 
 It is certain that we have not yet found the ideal fixing 
 agent for cell contents. Such an agent must not be a solvent 
 of any of the cell contents, must penetrate rapidly, must pre- 
 serve structures perfectly, and must harden so thoroughly that 
 every detail shall remain unchanged during the subsequent 
 processes of dehydrating, clearing, imbedding, sectioning, and 
 staining. 
 
CHAPTER VI. 
 
 STAINING. 
 
 Staining is one of the most important and most complicated 
 processes of microtechnique. The formulae are largely empirical, 
 and there is still abundant room for experimentation in the use 
 of mordants and in the effect of the same stain or combination 
 after various fixing agents. 
 
 Stains may be classified in various ways ; e.g., there are three 
 great groups of stains — the Carmines, the Haematoxylins, and 
 the Anilins. Stains may be classified as basic and acid, or they 
 may be regarded as general and specific. A general stain affects 
 all the elements, while a specific stain affects only certain 
 elements or stains some elements more deeply than others. 
 Stains which show a vigorous afifinity for the nucleus have been 
 called nuclear stains, and those which affect the cytoplasm more 
 than the nucleus have been termed plasma stains. Of course, 
 such stains are specific. 
 
 We shall consider some of the more important haematoxylins, 
 carmines, and anilins, reserving general directions and theoreti- 
 cal questions until the end of this chapter. Many of the formulae 
 are taken from The Microtomist's Vade-Mecum (Lee), which is 
 easily the most complete compendium of stains and other 
 reagents concerned in microtechnique. It is to be regretted 
 that botanists have no book of this character, but it must be 
 confessed that we have not the material for such an extensive 
 work. Other formulae are from Botanical Microtechnique (Zim- 
 mermann) and from Stirling's Histology. The directions for 
 using a stain apply to stains made up according to the formulae 
 which are given here, and may need modification if other formulae 
 are employed. It is hoped, however, that the directions will 
 give the student sufficient insight into the rationale of staining 
 to enable him to make any necessary modifications. 
 
 33 
 
34 Methods in Plant Histology 
 
 THE HJEMATOXYLINS. 
 
 The most important hematoxylins are Delafield's haema- 
 toxylin, Kleinenberg's haematoxylin, Erlich's haematoxylin, 
 Boehmer's haematoxylin, Mayer's haem-alum, and Haidenhain's 
 iron alum-haematoxylin. 
 
 All the haematoxylins mentioned above contain alum, and, 
 according to Mayer, who has written the most important work 
 on haematoxylin stains ("Ueber das Farben mit Hamatoxylin," 
 Mitth. a. d. Zool. Station zu Neapel, lo : 170-186, 1891, and " Ueber 
 Hamatoxylin, Carmin und verwandten Materien," Zeit. f. wiss. 
 Mikr., 16 1 196-220, 1899). "The active agent in them is a com- 
 pound of haematin with alumina. This salt is precipitated in the 
 tissues, chiefly in the nuclei, by organic and inorganic salts there 
 present [e. g., by the phosphates) and perhaps also by other 
 organic bodies belonging to the tissues." These salts are fixed 
 in the tissues by the killing and fixing agent, and when the stam 
 is applied a chemical combination results, Haematoxylins stain 
 well after any of the fixing agents described in the preceding 
 paper, but they are most effective when used after members of 
 the chromic-acid series. 
 
 Delafield's Haematoxylin. — "To 100 cc. of a saturated solution 
 of ammonia alum add, drop by drop, a solution of i g. of haema- 
 toxylin dissolved in 6 cc. of absolute alcohol. Expose to air 
 and light for one week. Filter. Add 25 cc. of glycerine and 
 25 cc. of meth3''l alcohol. Allow to stand until the color is 
 sufficiently dark. Filter, and keep in a tightly stoppered bottle." 
 (Stirling and Lee.) The addition of the glycerin and methyl 
 alcohol will precipitate some of the ammonia alum in the form 
 of small crystals. The last filtering should take place four 
 or five hours after the addition of the glycerine and methyl 
 alcohol. 
 
 The solution should stand for at least two months before it 
 is ready for using. This "ripening" is brought about by the 
 oxidation of haematoxylin into haematin, a reaction which may 
 be secured in a few minutes by a judicious application of per- 
 oxide of hydrogen. 
 
Stai?ting 3 5 
 
 Transfer to the stain from 35 per cent, alcohol or from water. 
 The length of time required is exceedingly variable. Sometimes 
 sections will stain deeply in three minutes, but it is often necessary 
 to stain for thirty minutes or even longer. This stain may be 
 diluted with several times its own volume of water ; when this is 
 done, the time required is correspondingly long, but the staining 
 is frequently more precise. The length of time required will be 
 fairly uniform for all material taken from the same bottle. This 
 fact indicates that the washing process, which follows killing and 
 fixing, is an important factor ; if the washing has been thorough, 
 the material will stain readily; but if the washing has been insuf- 
 ficient, the material may stain slowly or not at all. The washing 
 is particularly important when the fixing agent contains an acid. 
 Transfer from the stain to water. Distilled water is neither 
 necessary nor desirable. Some writers recommend washing for 
 twenty-four hours, but this seems unnecessary; an hour is usually 
 enough, and a few minutes is often sufficient. Precipitates are 
 often formed when slides are transferred directly to alcohol from 
 this stain ; otherwise, it would be better to transfer from the 
 stain to 35 per cent, alcohol. Pass through the alcohols to 70 
 per cent, alcohol and then give the slide a few dips (two seconds 
 is often sufficient) in acid alcohol (0.5 cc. HCl. to 100 cc. of 
 70 per cent, alcohol). This extracts the stain more rapidly from 
 other parts than from the nuclei, and hence gives a good nuclear 
 stain. Some prefer to stain for a very short time and use no 
 acid alcohol, but, as a rule, it seems best to overstain and then 
 differentiate in this way, because sharper contrasts are obtained. 
 Transfer from acid alcohol to 70 per cent, alcohol and leave here 
 until a rich purple color replaces the red due to the acid. Since 
 small quantities of the acid alcohol are carried over into the 70 
 per cent, alcohol, it is well to add a drop of ammonia now and 
 then to neutralize the effect of the acid. Too much ammonia is 
 to be avoided, for it gives a disagreeable bluish color with poor 
 differentiation, probably on account of the precipitation of 
 alumina. The slide may now be passed through the alcohols, 
 cleared in xylol, and mounted in balsam ; or, if a double stain be 
 
36 Methods in Plant Histology 
 
 preferred, treat for thirty seconds to one minute with eosin, 
 erythrosin, or some other stain affording a good contrast; rinse 
 in 70 per cent, alcohol and proceed as usual. 
 
 Delafield's hsematoxylin is the most generally useful stain in 
 the haematoxylin group. It brings out cellulose walls very 
 sharply, and consequently is a good stain for embryos and the 
 fundamental tissue system in general. With safranin it forms a 
 good combination for the vascular system, the safranin giving 
 the lignified elements a bright red color, while the haematoxylin 
 stains the cellulose a rich purple. It is a good stain for chro- 
 matin, and the achromatic structures show up fairly well, but can 
 be brought out much better by special methods. Archesporial 
 cells and sporogenous tissue are very well defined if proper care 
 be taken. Whenever you are in doubt as to the selection of a 
 stain for general purposes, we should advise the use of Delafield's 
 haematoxylin. 
 
 The following is a general schedule for staining paraffin sec- 
 tions on the slide in Delafield's haematoxylin : 
 
 1. Stain (from water or 35 per cent, alcohol) 5 minutes. 
 
 2. Rinse in water, 5 minutes. 
 
 3. Thirty-five per cent., 50 per cent., and 70 per cent, alcohol, 30 seconds 
 each. 
 
 4. Acid alcohol, i second. 
 
 5. Seventy per cent, alcohol, i minute. 
 
 6. Eighty-five per cent., 95 per cent., and too per cent, alcohol, 30 seconds 
 each. ^ 
 
 7. Xylol, I minute. 
 
 8. Mount in balsam. 
 
 If, after rinsing in water, the stain is evidently too weak, put 
 the slide or section back into the stain until it appears over- 
 stained. For thin sections (lO/^ or less) thirty seconds in each 
 of the alcohols will be unnecessarily long, although thirty minutes 
 would do no damage. Give the slide a single dip into the acid 
 alcohol, transfer it quickly to the 70 per cent, alcohol, and then 
 examine it ; if it still appears overstained, give it another dip in 
 the acid alcohol, and repeat the process until the stain is what 
 you want. After the haematoxylin is just right, apply a contrast 
 
Staining 37 
 
 stain, if you wish to double-stain. It is a good plan to move the 
 slide gently to and fro in the absolute alcohol. Before trans- 
 ferring to the xylol, wipe the alcohol from the back of the slide, 
 or at least rest the corner of the slide upon blotting paper 
 for two or three seconds, in order that you may not carry over 
 so much alcohol into the xylol, and thus impair this rather 
 expensive reagent. The slide may also be moved gently to and 
 fro in the xylol. Add a drop of balsam and a cover. Since the 
 xylol is very volatile, this last step must be taken quickly. If 
 blackish spots appear, they are usually caused by the drying of 
 sections before the balsam and cover are added ; if there are 
 whitish spots or an emulsion-like appearance, the clearing is not 
 thorough ; this may be caused by poor xylol (or other clearing 
 agent); by absolute alcohol which is considerably weaker than 
 its name implies (the absolute alcohol must test at least as high 
 as 99 per cent., and ought to test as high as 99.5 per cent., if 
 xylol is to be used for clearing) ; or the difficulty may be caused 
 by passing too quickly through the absolute alcohol and xylol, 
 or may even be caused by moisture on the cover-glass. 
 
 Kleinenberg's Haematoxylin. — This stain has had a wide use, 
 but it is now largely replaced by better formulae. It is men- 
 tioned here merely because it is on the shelves of so many 
 laboratories! It is doubtful whether it is equal to Delafield's in 
 any kind of botanical work. A good description is given in the 
 Quarterly Journal of the Microscopical Society, 74 : 208 — , 1897. 
 Erlich's Haematoxylin. — 
 
 Distilled water, 50 cc. 
 
 Absolute alcohol, 50 cc. 
 
 Glycerine, 50 cc. 
 
 Glacial acetic acid, 5 cc. 
 
 Haematoxylin, i g. 
 
 Alum in excess. 
 
 Keep it in a dark place until the color becomes a deep red. 
 If well stoppered, it will keep indefinitely. Transfer to the stain 
 from 50 per cent, or 35 per cent, alcohol. Stain five to thirty 
 minutes. Since there is no danger from precipitates and the 
 
1 
 
 38 Methods in Pla?it Histology 
 
 solution does not overstain, it is not necessary to treat with 
 water or with acid alcohol, but the slide may be transferred from 
 the stain to 70 per cent, alcohol. Eosin, erythrosin, or orange 
 G are good contrast stains. 
 Boehmer's Haematoxylin. — 
 
 Hsematoxylin, I g. 
 
 Absolute alcohol, 1 2 cc. 
 
 Alum, I g. 
 
 Distilled water, 240 cc. 
 The solution A must ripen for two months. When wanted 
 for use, add about 10 drops of A to 10 cc. of B. Stain ten to 
 twenty minutes. Wash in water and proceed as usual. 
 
 Mayer's Haem-Alum. — Haematoxylin, i g., dissolved with 
 heat in 50 cc. of 95 per cent, alcohol and added to a solution 
 of 50 g. of alum in a liter of distilled water. Allow the mixture 
 to cool and settle ; filter ; add a crystal of thymol to preserve 
 from mold. (Lee.) 
 
 It is ready for use as soon as made up. Unless attacked by 
 mold, it keeps indefinitely. Transfer to the stain from water. 
 It is seldom necessary to stain for more than ten minutes, and 
 four or five minutes is generally long enough. As a rule, better 
 results are secured by diluting the stain (about i cc. to 10 cc. of 
 distilled water) and allowing it to act for ten hours or over night. 
 This is a good stain for the nuclei of filamentous algae and fungi, 
 since it has little or no effect upon cell walls or plastids. Wash 
 thoroughly in water and transfer to 10 per cent, glycerine. 
 Specimens may be mounted in balsam, if they can be gotten 
 through without shrinking. 
 
 Haidenhain's Iron Alum-Haematoxylin. — This stain was intro- 
 duced by Haidenhain in 1892 and has gained a well-deserved 
 popularity with those engaged in cytological work. Two solu- 
 tions are used, and they are never mixed : 
 
 A. One and a half to 4 per cent, aqueous .solution of ammonia sulphate 
 of iron. (At present we use a 3 per cent, solution.) 
 
 B. One-half per cent, aqueous solution of haematoxylin. 
 
 The first solution acts as a mordant, i. e., it does not stain, 
 but prepares the tissue for the action of the second solution. 
 
Staining 39 
 
 Transfer to the iron-alum from water ; allow this solution to 
 act for two hours ; wash in water five minutes, and then stain 
 in the yi per cent, haematoxylin ten hours or over night. 
 Rinse in water five minutes and treat for a second time with the 
 iron alum, which now rapidly extracts the stain. The action of 
 the iron alum should be watched under a microscope, and when 
 the chromosomes of karyokinetic figures appear sharply defined, 
 the slide should at once be placed in water and washed for at 
 least an hour, since a very little of the iron alum, if left in the 
 tissue, will cause the preparation to fade. If staining for details 
 other than nuclei, the slide must be transferred to water as soon 
 as the desired effect is produced. After the washing in water, 
 the slide is passed through the alcohols, cleared in xylol, and 
 mounted in balsam. This stain is excellent for the filamentous 
 algae and fungi, and it keeps well in glycerine. For preparations 
 to be mounted in balsam, erythrosin, fuchsin, or orange G make 
 good contrast stains. Apply the second stain after the last 
 washing in water. The second stain should always be very light. 
 
 Chromosomes and centrosomes ("of those plants which have 
 centrosomes") take a brilliant black, and other details, though 
 not brightly colored, often show excellent definition. 
 
 The times given above must not be accepted as final. Many 
 prefer to wash in water for several hours after the first immersion 
 in iron alum. A plan which has proved convenient and very 
 successful is to put the slide into the iron alum in the morning, 
 let it wash in water during the afternoon, leave it in the ^ per 
 cent, of haematoxylin over night, and finish the preparation the 
 next morning. It is a long process, requiring care, patience, and 
 judgment, but it is worth the effort. 
 
 THE CARMINES. 
 
 This group of stains, immensely popular a few years ago, has 
 rapidly lost favor among botanists as newer stains and combina- 
 tions have appeared. When it is desirable to stain in bulk, 
 nothing has been found which will serve better than the car- 
 mines. Only three of these stains will be considered : 
 
40 Methods i?i Plant Histology 
 
 Greenacher's Borax Carmine. — 
 
 Carmine, 3 g. 
 Borax, 4 g. 
 
 Distilled water, 100 cc. 
 k 
 
 Dissolve the borax in water and add the carmine, which is 
 
 quickly dissolved with the aid of gentle heat. Add lOO cc. of 
 70 per cent, alcohol and filter. (Stirling.) 
 
 The following is a slightly different method for making this 
 stain from the ingredients mentioned above : Dissolve the 
 borax in water, add the carmine, and heat gently for ten minutes ; 
 after the solution cools, add the alcohol and filter ; let the solu- 
 tion stand for two or three weeks, then decant, and filter again. 
 
 Stain the material in bulk from 50 per cent, alcohol one to 
 three days, then treat with acid alcohol (50 cc. of 70 per cent, 
 alcohol + 2 drops of hydrochloric acid) until the color becomes 
 a clear red ; this may require only a few hours, but may take 
 two or three days. The material may then be passed through 
 the rest of the alcohols (six to twenty-four hours each), cleared, 
 imbedded, and cut. After the sections are thoroughly fastened 
 to the slide, the paraflfin should be dissolved off with xylol. The 
 balsam and cover may be added immediately, or the slide may' 
 be passed down through the alcohols for a contrast stain. 
 
 Alum Carmine. — A 4 per cent, aqueous solution of ammonia 
 alum is boiled twenty minutes with i per cent, of powdered 
 carmine. Filter after it cools. (Lee.) 
 
 Stain from water twelve to twenty-four hours and wash in 
 water. No acid alcohol is needed, since the solution does not 
 overstain. 
 
 Alum Cochineal. — 
 
 Powdered cochineal, 50 g. 
 
 Alum, 5 g. 
 
 Distilled water, 500 cc. 
 
 Dissolve the alum in water, add the cochineal, and boil ; 
 evaporate down to two-thirds of the original volume, and filter. 
 Add a few drops of carbolic acid to prevent mold. (Stir- 
 ling.) 
 
Staining 4 1 
 
 Stain as with alum carmine. A few years ago it was a very 
 common practice to stain in bulk in alum cochineal and counter- 
 stain on the slide with bismark brown. 
 
 THE ANILINS. 
 
 Many of the most brilliant and beautiful stains yet discovered 
 belong to this group. These stains are so numerous that we 
 shall not attempt to mention even their names, but shall con- 
 sider only those which are in most common use by botanists. 
 The following formula has proved to be fairly satisfactory for all 
 the anilins mentioned in this account, but other formulae will be 
 given for most of the stains : 
 
 Make a 3 per cent, solution of anilin oil in distilled water; 
 shake well and frequently for a day; add enough alcohol to make 
 the whole mixture about 20 per cent, alcohol ; add i g. of cyanin, 
 erythrosin, safranin, gentian violet, etc., to each lOO cc. of this 
 solution. 
 
 The anilins keep well in balsam, but not in glycerine. Xylol 
 is a good clearing agent for all of them, but clove oil is very much 
 better in case of gentian violet. Unfortunately, some of them 
 do not give permanent stains. Some are acid, some basic, and 
 some neutral. 
 
 The rapidity with which sections must be transferred from 
 one fluid to another makes many of them more difficult to man- 
 age than the haematoxylins or the carmines, but the stains are so 
 valuable that even the beginner should spend most of his time 
 with the anilins. 
 
 Many anilins stain quite deeply in one to twenty minutes, 
 but if the stain washes out during the dehydrating process, stain 
 longer, even ten to thirty hours, if necessary. If the stains are 
 made up according to the formula mentioned above, transfer to 
 the stain from 35 per cent, alcohol and from the stain to 35 per 
 cent, alcohol, if the stain does not wash out too rapidly; if the 
 stain washes out, try 50 per cent., 70 per cent., 85 per cent., 95 
 per cent., or even directly to absolute alcohol. It will often be 
 found impracticable to transfer from the stain to alcohols weaker 
 
42 Methods ifi Plant Histology 
 
 than the 85 per cent., lest all the stain be washed out before the 
 clearing agent is reached. 
 
 Since the anilins are seldom used as single stains, but almost 
 invariably in combination with other stains, the logical order 
 will be disregarded and the stains will be treated, as they are 
 used, in their most usual combinations. 
 
 Cyanin and Erythrosin. — Stain in cyanin five to ten minutes 
 or longer ; rinse quickly in alcohol, and then stain thirty seconds 
 to one minute in erythrosin. If the cyanin washes out, stain for 
 an hour, and if it still washes out, omit the rinsing in alcohol 
 and transfer directly from the cyanin to the erythrosin. 
 
 The erythrosin may be used first; in this case stain for five 
 minutes in erythrosin, transfer directly to cyanin, and stain for 
 about ten seconds. Try 70 per cent, alcohol, but if the stains 
 are lost, try 85 per cent., 95 per cent., or even transfer directly 
 to 100 per cent. 
 
 This is a good combination for general work, and if properly 
 used is excellent for mitotic phenomena and the most delicate 
 cytological work. Delafield's haematoxylin may be used with 
 erythrosin. Stain first with the haematoxylin, and, after the pur- 
 ple color has replaced the red due to the acid, stain lightly with 
 erythrosin. Eosin may be used instead of erythrosin, but is less 
 transparent. Eosin will be mentioned later in connection with 
 special methods for algae and fungi. 
 
 Flemming's Safranin-Gentian Violet-Orange. — Safranin has 
 long been a famous stain for karyokinesis. This triple combina- 
 tion was published in 1891, but its value in plant cytology was 
 not thoroughly appreciated until five or six years later, when its 
 application was developed to a high degree of perfection by 
 various investigators of the Bonn (Germany) school. 
 
 According to Flemming, stain two to three days in safranin 
 (dissolve 0.5 g. safranin in 50 cc. absolute alcohol, and after 
 four days add 10 cc. distilled water); rinse quickly in water ; 
 stain one to three hours in a 2 per cent, aqueous solution of 
 gentian violet ; wash quickly in water, and then stain one to 
 three minutes in a i per cent, aqueous solution of orange G. 
 
Staining 43 
 
 Tranfer from the stain to absolute alcohol, clear in clove oil, and 
 mount in balsam. 
 
 The following method seems to be better for mitotic phe- 
 nomena in plants : Transfer to safranin from 35 per cent, alcohol 
 and stain sixteen to twenty-four hours. If the stain acts for 
 only a few hours, it washes out too rapidly to be controlled with 
 any precision. (The safranin may be made up according to the 
 formula given in the preceding paragraph or according to the 
 general formula.) Rinse in water for a minute and then in 50 
 per cent, alcohol until only nucleoli and the chromosomes of 
 dividing nuclei retain the red color. If the alcohol does not 
 wash out the stain sufficiently, dip the slide in acid alcohol (not 
 more than 2 or 3 drops of hydrochloric acid to 100 cc. of 
 50 per cent, alcohol). If acid has been used, wash for a 
 moment in pure 50 per cent, alcohol, and then stain for four or 
 five minutes in gentian violet (aqueous solution, or made up 
 according to the general formula). Rinse for a few seconds in 
 water and then stain about thirty seconds in a i per cent, aque- 
 ous solution of orange G. Transfer from the stain directly to 
 absolute alcohol and hasten the dehydrating by gently rinsing 
 the slide in the fluid. One or two quick dips in the 95 per cent, 
 alcohol before transferring to the absolute does not seem to do 
 any harm and certainly saves the more expensive reagent. As a 
 rule, not more than ten seconds can be allowed for dehydrating, 
 because the gentian violet washes out so rapidly. The extreme 
 transfer from the orange G, an aqueous stain, to absolute alco- 
 hol, which must often be made to prevent the gentian violet 
 from washing out, indicates that, for staining very thin sections 
 on the slide, the series of alcohols from 35 per cent, up to 100 
 per cent, is not at all necessary. 
 
 Treat with clove oil for five to ten seconds. The clove oil 
 not only clears, but it rapidly extracts the gentian violet, pro- 
 ducing an elegant differentiation. Replace the clove oil by 
 cedar oil, if you have strictly first-class cedar oil. Such cedar 
 oil is very thin, light yellow in color, and has a faint odor of 
 cedar wood. It costs about the same as clove oil. Do not get 
 
44 Methods hi Pla?it Histology 
 
 the expensive oil used for immersion lenses. If your cedar oil 
 is colorless and has a strong odor, do not use it at all, but drain 
 off the clove oil as thoroughly as possible by resting the edge of 
 the slide on blotting paper for two or three minutes, and then 
 mount in balsam. If any considerable amount of clove oil is 
 left on the slide, the preparation is almost sure to fade. The 
 cedar oil is not a solvent of gentian violet, and consequently 
 there is little danger that the preparation will fade when this oil 
 is used. Some investigators transfer from clove oil to xylol, 
 but it seems to us that the brilliancy of the gentian violet is 
 impaired. Chromosomes should take a clear red and the spindle 
 fibers a bright violet. This combination is generally used after 
 Flemming's solution, but seems to do equally well after other 
 members of the chromic-acid series. Achromatic structures do 
 not seem to stain well after corrosive sublimate or picric acid. 
 
 Fuchsin. — Use a i or 2 per cent, solution in water or in 70 per 
 cent, alcohol. Transfer to the alcoholic solution from 70 per 
 cent, alcohol ; stain one to two hours; differentiate the stain in 
 I per cent, solution of picric acid in 70 per cent, alcohol — this 
 may require thirty seconds or several minutes ; rinse in 70 per 
 cent, alcohol until a bright red replaces the yellowish color due 
 to the acid, and then proceed as usual. 
 
 Iodine Green. — A i per cent, solution in 70 per cent, alcohol 
 is good for the vascular system of plants. This stain resists the 
 washing-out process better than methyl green. Stain in iodine 
 green at least an hour, and it is not a bad plan to stain over 
 night ; rinse in 70 per cent, alcohol ; stain fifteen seconds to one 
 minute in erythrosin, and proceed as usual. Methyl green may 
 be made and used in the same way. 
 
 Fuchsin and Iodine Green Mixtures. — Two solutions are kept 
 separate, since they do not retain their efficiency long after they 
 
 are mixed : 
 
 . ( 0.1 g. fuchsin (acid). 
 I 50 cc. distilled water. 
 „ j 0.1 g. iodine green. 
 50 cc. distilled water. 
 
Staining 45 
 
 iioo cc. absolute alcohol. 
 I cc. glacial acetic acid. 
 0.1 g. iodine. 
 Mix equal parts of A and B. Transfer to the stain from 
 water. The proper time must be determined by experiment. 
 Twenty-four hours might be recommended for a trial. Transfer 
 from the stain directly to solution C and from C to xylol. 
 Another formula : 
 
 A. 0.5 g. acid fuchsin. 
 
 B. 0.5 g. iodine green. 
 
 Mix a pipette full of A with a pipette full of B ; stain two to 
 eight minutes ; transfer to 85 per cent, or 95 per cent, alcohol, 
 dehydrate rapidly, clear in xylol, and mount in balsam. Both 
 these formulae are good for karyokinesis. 
 
 Bismark Brown. — Use a 2 per cent, solution in 70 per cent, 
 alcohol. If material has been stained in bulk in one of the 
 carmines, a few minutes' staining on the slide with bismark brown 
 gives a good contrast. It is particularly good for cell walls. 
 
 Nigrosin. — Use a i or 2 per cent, solution in water. A few 
 drops of this solution to a watch-glass full of water stains fila- 
 mentous algae or fungi in one to three hours. The stain keeps 
 well in glycerine or balsam, but it is hard to get these forms 
 into balsam without more or less shrinking. 
 
CHAPTER VII. 
 
 GENERAL REMARKS ON STAINING. 
 
 It must be remembered that many things may be examined 
 alive without killing, fixing, staining, or any of those processes. 
 A filament of Spirogyra shows the chromatophore nicely if 
 merely mounted in a drop of water; the nucleus may be visible, 
 and the pyrenoids can usually be located. Of course, such a 
 study is necessary if one is to understand anything about the 
 plant, and in an elementary class this might be sufficient, but a 
 drop of iodine solution applied to the edge of the cover would 
 emphasize certain details, e.g., the starch in the pyrenoids would 
 appear blue, the nucleus a light brown, and the cytoplasm a 
 lighter brown. This illustrates at least one advantage to be 
 gained by staining ; it enables us to see structures which would 
 otherwise be invisible, or almost invisible. 
 
 With so many stains at our disposal, it at once becomes a 
 problem just which stain or combination to use in each particu- 
 lar case. Beautiful and instructive preparations occasionally 
 result from some happy chance, but uniform success demands 
 skill and judgment in manipulation, and also a knowledge of the 
 structures which are to be differentiated. Let us take a vascular 
 bundle for illustration. Safranin stains the xylem a bright red, 
 but, with judicious washing, is entirely removed from the cam- 
 bium and cellulose elements of the phloem. A careful staining 
 with Delafield's haematoxylin now gives a rich purple color to 
 the cellulose elements which were left unstained by the safranin, 
 thus contrasting sharply with the lignified elements. If cyanin 
 and erythrosin be used, the xylem takes the blue and the cam- 
 bium and phloem take the red. Many terms have been given 
 to indicate the affinity of certain tissues for certain stains. 
 Auerbach used the terms "erythrophilous" and "cyanophilous" 
 in 1890. This eminent zoologist studied spermatozoa and ova. 
 
 47 
 
48 Methods in Plant Histology 
 
 He found that, if preparations containing both spermatozoa and 
 ova were stained with cyanin and erythrosin, the nuclei of the 
 spermatozoa took the cyanin, while the nuclei of the ova pre- 
 ferred the erythrosin; hence the terms "cyanophilous " and 
 "erythrophilous." Auerbach regarded these differences as an 
 indication of sexual differences in the cells. 
 
 Rosen (1892) supported this theory, and even went so far 
 a$ to regard the,tube nucleus of the pollen grain as female, on 
 account of its erythrophilous staining. In connection with this 
 theory it was suggested that the ordinary vegetative nuclei are 
 hermaphrodite, and that in the formation of a female germ 
 nucleus the male elements are extruded, leaving only the ery- 
 throphilous female elements ; and, similarly, in the formation of a 
 male nucleus the female elements are extruded, leaving only the 
 cyanophilous male elements. 
 
 As long ago as 1884 Strasburger discovered that with a 
 mixture of fuchsin and iodine green the generative nucleus of a 
 pollen grain stains green, while the tube nucleus stains red. In 
 1892 {Verhalten des Pollens) he discussed quite thoroughly the 
 staining reactions of the nuclei. The nuclei of the small pro- 
 thallial cells of gymnosperm microspores are cyanophilous like 
 the male generative nuclei. The nuclei of a nucellus surround- 
 ing an embryo-sac are also cyanophilous, while the nuclei of 
 structures within the sac are erythrophilous. His conclusion is 
 that the cyanophilous condition in both cases is due to poor 
 nutrition, while the erythrophilous condition is due to abundant 
 nutrition. A further fact in support of the theory is that the 
 nuclei of the adventitious embryos which come from the nucellus 
 of Funkia ovata are decidedly erythrophilous, while the nuclei 
 of the nucellus to which they owe their food-supply are cyano- 
 philous. 
 
 In division stages nuclei are cyanophilous, but from anaphase 
 to resting stage cytoplasm is taken into the nucleus, and the 
 cyanophilous condition gradually changes to the erythrophilous. 
 
 An additional fact in favor of this theory is that in Ephedra 
 the tube nucleus which has very little cytoplasm about it is 
 
General Remarks o?i Stainifig 49 
 
 cyanophilous. Strasburger claims that there is no essential dif- 
 ference between male and female generative nuclei, and subse- 
 quent observation has shown that within the oospore the sex 
 nuclei are alike in their reaction to stains. 
 
 Malfatti (1891) and Lilienfeld (1892-3) claim that these 
 reactions are dependent upon the amount of nucleic acid present 
 in the structures. During mitosis the chromosomes consist of 
 nearly pure nucleic acid and are intensely cyanophilous, but the 
 protoplasm, which has little or no nucleic acid, is erythrophi- 
 lous. There is a gradual transition from the cyanophilous con- 
 dition to the erythrophilous, and vice versa, the acid structures 
 taking basic stains and basic structures the acid stains. 
 
 The terms "erythrophilous" and "cyanophilous" are falling 
 into disuse, since the affinity is for basic and acid dyes, rather than 
 for blue or red colors. That the terms are misnomers becomes 
 evident when a combination like safranin (basic) and acid green 
 (acid) is used, for the cyanophilous structures stain red, and the 
 erythrophilous green. 
 
 Probably but few investigators who have attained any pro- 
 ficiency in microtechnique have not asked themselves how 
 much dependence can be placed upon staining reactions as a 
 means of analysis. Do staining reactions enable us to determine 
 the chemical composition of a structure ? If two structures stain 
 alike with Delafield's haematoxylin, does this mean that they 
 have the same chemical composition ; or if, on the other hand, 
 they stain differently, must they necessarily be different in their 
 chemical composition? Delafield's haematoxylin, when care- 
 fully used, gives a rich purple color, but a careful examination 
 will often show that in the same preparation some structures 
 stain purple, while others stain red. Does this mean that the 
 purple and red structures must have a different chemical com- 
 position ? Many people believe that structures which stain dif- 
 ferently with a given stain must be chemically different, but 
 they readily agree that structures which stain alike are not 
 necessarily^ similar in chemical composition. Chromosomes of 
 dividing nuclei and lignified cell walls stain alike with safranin ; 
 
50 Methods in Plant Histology 
 
 chromosomes and cellulose cell walls stain much alike with 
 Delafield's haematoxylin ; but everyone recognizes that the 
 chromosome is very different in its chemical composition from 
 either the cellulose or the lignified wall. 
 
 According to Fischer (1897 and 1900), stains indicate physi- 
 cal but not chemical composition. Fischer experimented with 
 substances of known chemical composition. Egg albumin was 
 shaken until small granules were secured. These were fixed 
 with the usual fixing agents, and then stained with Delafield's 
 haematoxylin. The extremely small granules stained red, while 
 the larger ones became purple. Since the granules are all alike 
 in chemical composition, Fischer concluded that the difference 
 in staining must be due to physical differences. With safranin, 
 followed by gentian violet, the larger granules stain red and the 
 smaller violet ; if, however, the gentian violet be used first, then 
 treated with acid alcohol and followed by safranin, the larger 
 granules take the red and the smaller the gentian violet. In 
 root tips similar results were obtained. Safranin followed by 
 gentian violet stained chromosomes red and spindle fibers violet, 
 while gentian violet followed by safranin stained the chromo- 
 somes violet and the spindle red. One often reads that chro- 
 mosomes owe their strong staining capacity to nuclein, and 
 especially to the phosphorus, but, according to Fischer, this is 
 shown to be unfounded, since albumin gives similar results and 
 yet contains no phosphorus, and is not chemically allied to 
 nuclein. Delafield's haematoxylin is one of the so-called nuclear 
 stains. The nuclei of animals and plants stain deeply with this 
 reagent, but cellulose membranes, the dense protoplasm of 
 embryonic cells, the pyrenoids of green algae, and many other 
 structures resemble nuclei in their staining. The most critical 
 work on this subject has been done by those who are investigat- 
 ing the structure of the Cyanophyceae and Bacteria to determine 
 whether these forms have nuclei or not. Butschli claims that 
 the granules which stain red with haematoxylin are to be identi- 
 fied with chromatin, while Fischer, whose results have just been 
 given, claims that staining indicates merely physical differences. 
 
Ge?ieral Remarks on Staining 51 
 
 The subject cannot yet be regarded as settled, but whatever may 
 be true in regard to these conflicting theories, all agree that 
 stains are of the highest importance in differentiating struc- 
 tures, and in bringing out details which would otherwise be 
 invisible. 
 
CHAPTER VIII. 
 
 PRACTICAL HINTS ON STAINING. 
 
 In later chapters specific directions will be given for making 
 a series of preparations ranging from the lowest algae to the 
 flowering plants, but a few suggestions will be made here. 
 
 The number of stains in the catalogues is becoming so great 
 that it is impossible to become proficient in the use of all of 
 the;m. It is far better to master a few of the most valuable stains 
 than to do indifferent work with many. The beginner, especially 
 if rather unacquainted with the details of plant structure, may 
 believe that he has an excellent preparation when it is really a 
 bad, or at most an indifferent, one. To illustrate, let us suppose 
 that a pollen mother-cell in a late spirem stage has been stained 
 with cyanin and erythrosin. A preparation in which the cell 
 merely shows a differentiation into nucleus and cytoplasm must 
 be classed as bad ; if the nucleus shows a definitely outlined 
 spirem thread, the preparation is better, but is still only indiffer- 
 ent ; if the thread appears as a delicate red ribbon bordered by 
 blue granules, the staining may be regarded as a success. If 
 mitotic figures have been stained with cyanin and erythrosin, a 
 first-class preparation should show blue chromosomes and red 
 spindles ; if stained with safranin and gentian violet, the chromo- 
 somes should be red and the spindles violet. 
 
 In staining growing points, apical cells, young embryos, 
 antheridia, archegonia, and many such things, the cell walls are 
 the principal things to be differentiated, if the preparations are 
 for morphological study. As a rule, it is better in such cases not 
 to use double staining, but to select a stain which stains the cell 
 walls deeply without obscuring them by staining starch, chloro- 
 phyll, and other cell contents. For example, try the growing 
 point of Equisetum The protoplasm of such growing points is 
 very dense. If Delafield's haematoxylin and erythrosin be used, 
 
 S3 
 
54 Methods in Pla?it Histology 
 
 the haematoxylin will stain the walls and nuclei, and will slightly 
 affect the other cell contents, but the erythrosin will give the 
 cytoplasm such a dense stain that the cell walls will be seriously 
 obscured. It would be better to use haematoxylin alone. The 
 same suggestion may well be observed in tracing the develop- 
 ment of antheridia, archegonia, embryos, and similar structures. 
 Permanent preparations are an absolute necessity for the 
 greater part of most advanced work, but let us not imagine that 
 we cannot examine anything until we have made a permanent 
 mount. It would be impossible to make a permanent mount of 
 the rotation of protoplasm. It is better for many purposes to 
 look at motile spores while they are moving. Use Spirogyra 
 while it is fresh and green (if you can), and use permanent prep- 
 arations only to bring out nuclei and other details which are 
 not so easily seen in living material. Examples might be multi- 
 plied. 
 
CHAPTER IX. 
 
 THE CELLOIDIN METHOD. 
 
 "Celloidin is a form of nitro-cellulose." It is very inflam- 
 mable, but does not explode. It may be obtained in the form 
 of tablets or cuttings, which have to be dissolved in a mixture 
 of equal parts of absolute alcohol and ether. It is customary to 
 use two solutions, a "thick" and a "thin." The thick solution 
 (about 10 or 12 per cent.) should have about the consistency of 
 thick syrup. The thin may be made by mixing equal parts of 
 the thick and ether alcohol. 
 
 As mentioned in the chapter on the "General Method," the 
 killing, washing, and dehydrating are the same as for the paraffin 
 method. After dehydrating in absolute alcohol the succeeding 
 steps are as follows : 
 
 1. Ether alcohol, i to 2 days. 
 
 2. Thin celloidin, 2 to 6 days. 
 
 3. Thick celloidin, 3 to 10 days. 
 
 It seems better, however, to begin with about 2 per cent, 
 celloidin and transfer successively through 4 per cent., 6 per 
 cent., etc., to 12 per cent., or to allow the 2 per cent, to concen- 
 trate by removing the cork for a short time each day. 
 
 The material may now be imbedded and mounted upon 
 a block at the same time. The blocks should have surface 
 enough to accommodate the objects, and should be about one- 
 fourth of an inch thick. White pine makes good blocks ; cork 
 is much inferior. Place the block for a moment in ether alcohol 
 and then dip into the 2 per cent, celloidin the end of the block 
 which was left rough by the saw. With the forceps remove a 
 piece of the material from the thick celloidin and place it upon 
 the block, taking care to keep it right side up. Dip the block 
 with its object first in thick celloidin, then in thin, and after 
 exposing to the air for a few minutes drop it into chloroform, 
 
 55 
 
56 Methods in Plant Histology 
 
 where it should remain for about ten to twenty hours. It should 
 then be placed in equal parts of glycerine and 95 per cent, 
 alcohol, where it may be kept indefinitely. If the material is 
 hard, like many woody stems, it will cut better after remaining 
 in this mixture for a couple of weeks. 
 
 In cutting, the knife should be set as obliquely as possible, 
 and both the knife and the object should be kept wet with the 
 mixture of glycerine and alcohol. As fast as they are cut the 
 sections are transferred with a soft brush to 70 per cent, alcohol. 
 The succeeding steps are the same as for free-hand sections, but 
 many stains are not available because they stain the celloidin. 
 Safranin and Delafield's haematoxylin, or Delafield's haematoxy- 
 lin and eosin, are good combinations for celloidin sections. Do 
 not use absolute alcohol for dehydrating, since it dissolves the 
 celloidin, but transfer from 95 per cent, alcohol to Eycleshymer's 
 clearing fluid (equal parts of bergamot oil, cedar oil, and car- 
 bolic acid), which clears readily from 95 per cent, alcohol. 
 Mount in balsam. 
 
 The following schedules for staining celloidin sections will 
 give the student a start. The times given will vary with the 
 thickness of the section and character of the tissue. 
 
 a. For staining in Delafield's haematoxylin and eosin : 
 
 1. Seventy per cent., 50 per cent., and 35 per cent, alcohol, 2 to 5 minutes 
 each. 
 
 2. Delafield's haematoxylin, 5 to 30 minutes. 
 
 3. Wash in water, 5 minutes. 
 
 4. Thirty-five per cent, and 50 per cent, alcohol, 2 to 5 minutes each. 
 
 5. Acid alcohol (i cc. hydrochloric acid -f- 100 cc. of 70 per cent, alcohol) 
 until the stain is extracted from the celloidin, or at least until the celloidin 
 retains only a faint pinkish color. 
 
 6. Seventy per cent, alcohol (not acid) until the purple color replaces the 
 red due to the acid. 
 
 7. Eosin (preferably a i per cent, solution in 70 per cent, alcohol), 2 to 5 
 minutes. 
 
 8. Eighty-five per cent., 95 per cent, alcohol, 2 to 5 minutes each. 
 Remember that 100 per cent, alcohol is not to be used with celloidin 
 
Celloidin Method 57 
 
 q. Eycleshymer's clearing fluid until transparent, usually i or 2 minutes, 
 
 but sometimes 5 or 10. 
 10. Mount in balsam. 
 
 b. For staining in safranin and Delafield's haematoxylin : 
 . I. Seventy per cent., 50 per cent., and 35 per cent, alcohol, 2 to 5 minutes 
 each, 
 
 2. Safranin, 6 to 24 hours, preferably the longer period. 
 
 3. Acid alcohol (try about i drop of hydrochloric acid to 30 cc. of 70 per 
 cent, alcohol) until the stain is removed from the celloidin or at least 
 becomes very faint. 
 
 4. Fifty per cent, and 35 per cent, alcohol, 2 to 5 minutes each. 
 
 5. Delafield's haematoxylin, 2 to 5 minutes. 
 
 6. Wash in water, 5 minutes. 
 
 7. Thirty-five per cent, and 50 per cent, alcohol, 2 to 5 minutes each. 
 
 8. Acid alcohol (the same as was used for the safranin) until the stain is 
 extracted from the celloidin. If it is found that this double immersion 
 in acid alcohol extracts too much safranin, the third step may be omitted 
 or shortened by removing the sections to the 50 per cent, alcohol before 
 the stain is thoroughly removed from the celloidin. 
 
 g. Eighty-five per cent, and 95 per cent, alcohol, 2 to 5 minutes each, 
 
 10. Eycleshymer's clearing fluid until cleared. 
 
 1 1. Mount in balsam. 
 
 The celloidin method has its disadvantages as well as its 
 advantages. It is extremely slow and tedious, and it is rarely 
 possible to cut sections thinner than lO/u,, while, on the other 
 hand, it gives smoother sections. The entire absence of heat 
 makes it very useful for delicate, succulent tissues. Stems and 
 roots which cannot be handled at all in paraffin cut well in 
 celloidin, and much larger sections can be cut than in paraffin. 
 
 When material is to be imbedded, use celloidin as a last 
 resort. Use paraffin when you ca7i, celloidin when you must. 
 
 I am indebted to my friend Mr. W. B. MacCallum for several 
 suggestions in regard to this method. 
 
CHAPTER X. 
 
 THE GLYCERINE METHOD. 
 
 It is hard to get the filamentous algae and fungi into balsam 
 without shrinking ; consequently, these forms are usually mounted 
 in glycerine or glycerine jelly. 
 
 Flemming's fluid and chromo -acetic acid are good fixing 
 agents. Corrosive sublimate in water, or in 70 per cent, alcohol, 
 used hot, is also to be recommended. For general morphology, 
 stain for six hours or over night in a ^ per cent, aqueous solution 
 of eosin, transfer directly to a i per cent, solution of acetic acid 
 in distilled water, and allow it to act for about five minutes ; wash 
 thoroughly in water to remove the acid, and then put the 
 material into a watch-glass in a 10 percent, solution of glycerine 
 in water. The watch-glass should be kept as free from dust as 
 possible, but should not be covered. As soon as the solution 
 appears to be about as thick as pure glycerine, the material is 
 ready for mounting. Place a small quantity of the material on 
 a slide, arrange it carefully, add a small drop of glycerine, and 
 a round cover. Seal with gold size (a varnish used by painters 
 in laying gold leaf). None of the sealing media will stick to 
 moist surfaces, hence it is essential that there should be only 
 enough glycerine to come to the edge of the cover. If it is 
 desired to mount rather large specimens, like the antheridia and 
 oogonia of Chara, it is best to spin a ring on the slide, thus 
 forming a shallow cell. Before the ring becomes hard the 
 material may be mounted and sealed, or the ring may be allowed 
 to harden, and just before a mount is to be made a very thin 
 ring of the varnish may be added. The mount is firmer if the 
 cover is not only sealed in the usual way, but also sticks to the 
 ring underneath. 
 
 If glycerine jelly is to be used, place the bottle in warm 
 59 
 
6o Methods i?i Plant Histology 
 
 water until the jelly becomes liquid, but avoid any unnecessary 
 heat. Take the material from the glycerine, add a drop of the 
 warm jelly, and seal as before. More detailed directions for 
 mounting material in glycerine are given in the chapters on algae 
 and fungi. 
 
Part II. 
 
 In the preceding chapters the principles and methods of tech- 
 nique have been described in a general way. It is often diffi- 
 cult, especially for a beginner, to apply general principles to 
 specific cases, and, besides, the types which he might select for 
 the preparations might not form a symmetrical collection. Con- 
 sequently, a series of forms has been selected which will not 
 merely serve for practice in microscopical technique, but will 
 also furnish the student with preparations for a fairly satisfactory 
 study of plant structures from the algae up to the angiosperms. 
 It is not at all our purpose to discuss general morphology, but 
 rather to answer, by means of sketches and specific directions, 
 the multitudinous questions which confront the instructor in the 
 laboratory. For those who have had a thorough training in 
 general morphology the following suggestions will be in some 
 degree surperfluous. Those who are beginning the study of 
 minute plant structure are referred to the standard text-books for 
 descriptions of the plants mentioned here. 
 
 STAi£ilOR11AL SCHOOL, 
 
CHAPTER XI. 
 
 THALLOPHYTES. ALG^. 
 CYANOPHYCE-ffi. 
 Wasserbliithe. — These forms occur as scums, often irides- 
 cent, on the surface of stagnant or quiet water. Some of the 
 commonest forms are Ccelosphcerium and Anabcena {jig. ij). 
 Some of the Chlorophyceai also 
 occur as Wasserbliithe. Where 
 the material is very abundant, 
 it may be collected by simply 
 skimming it off with a wide- 
 mouthed bottle, but where it is 
 rather scarce, it is better to filter 
 the water through a cloth, and 
 finally rinse the algse off into a 
 bottle. Enough formalin may 
 now be added to the water in 
 the bottle to make a 2 per cent, 
 solution. The material may be 
 kept here indefinitely, but after 
 a few hours it is ready for use. 
 If the forms are small, like 
 Atiabce7ta, smear a slide lightly 
 with Mayer's albumen fixative, 
 as if for paraffin sections, add a drop of the material and allow it 
 to dry, heat the slide gently to coagulate the albumen, or immerse 
 the slide in strong alcohol for a few minutes, and then proceed 
 with the staining. Cyanin and erythrosin is a good combination 
 for differentiating the granules. Delafield's haematoxylin, used 
 alone, stains some granules purple and others red. Iron alum- 
 haematoxylin is excellent for heterocysts. If the forms are large 
 enough to collapse with such treatment, the glycerine method 
 
 may be employed. 
 
 63 
 
 Coelosphaerium Kuetzingianum. B, Anabaena 
 flos-aquse. C, Anabaena gigantea. D and E, a 
 heterocyst and spore of A. gigantea drawn from 
 paraffin sections stained in cyanin and erythrosin. 
 
64 
 
 Methods in Pla?it Histology 
 
 If it is desirable to make paraffin sections, put the material, 
 drop by drop, on a piece of blotting paper until an appreciable 
 layer of sediment is obtained. Get the paper with its material 
 into paraffin by the usual method, taking great care not to wash 
 the algae off. After imbedding, trim away the 
 paper and dip the block in melted paraffin. 
 Sections can now be cut and stained in the 
 usual manner. 
 
 Oscillaria. — For most purposes it is best to 
 study Oscillaria in the living condition. It is 
 readily found in watering troughs, in stagnant 
 water, on damp earth, and in other habitats. 
 The commonest forms have a deep blue-green 
 or brownish color. For the purposes of identi- 
 fication and herbarium specimens, the material 
 may simply be placed on a slip of mica and 
 allowed to dry. When wanted for use, add a 
 drop of water and a cover, and the mount is ready for examina- 
 tion. For sections or 
 for glycerine mounts {\ 
 
 fix in chromo-acetic 
 acid. {Fig- ^4-) 
 
 Rivularia. — This 
 form is readily found 
 on the underside of the 
 leaves of water-lilies 
 [Nupkar, NymphcBa, 
 etc.) , but is also abun- 
 dant on submerged 
 leaves and stems of 
 other plants. It oc- 
 
 FlG. 14. Oscillaria. 
 
 Portions of two fila- 
 ments, the one at the 
 right showing a hormo- 
 gonium, h. 
 
 Fig. 15. 
 A. nodule crushed under 
 
 glass. B, tour filaments more 
 
 curs in the form of Wghly magnified, showing heterocysts at the base. 
 
 translucent, gelatinous nodules of various sizes. Chromo-acetic 
 acid gives beautiful preparations, but good results can also be 
 secured from formalin or picric-acid material. 
 
 The most instructive preparations for morphological study 
 
Thallophytes. AlgcB 
 
 65 
 
 Fig. 16. Tolypothrix. 
 b, a false branch, h, hetero- 
 
 can be obtained by the glycerine method. 
 Stain in eosin or Mayer's haem-alum. When 
 ready for mounting, crush a small nodule 
 by pressing on the cover-glass. Fig. i§ is 
 drawn from such a preparation. The par- 
 affin method is easily applied, since the 
 gelatinous matrix keeps the plants in 
 place. Glceotrichia, Nostoc, and forms of 
 similar habit may be prepared in the same 
 way. 
 
 Tolypothrix. — This form occurs as small 
 tufts, either floating in stagnant water or 
 attached to plants and stones. It furnishes 
 an excellent example of false branching. 
 {Fig. 16.) Scytojiema is a similar form 
 which is fairly common. The glycerine 
 method should be employed for perma- 
 nent preparations, but this, like all small '^^^' 
 filamentous algae, may be dried on mica for herbarium pur- 
 poses. 
 
 CHLOROPHYCEiE. 
 
 The ponds, ditches, and rivers of any locality will yield an 
 abundance and variety both of the unicellular and multicel- 
 lular members of this group. The unicellular and filamentous 
 members, together with such forms as Volvox, are best prepared 
 by the glycerine method. The structure is so much more com- 
 plicated than in the Cyanophyceae that it demands far more care 
 and skill to make good preparations, Chromo-acetic acid is a 
 good killing and fixing agent for the whole group, but Flem- 
 ming's fluid (weaker solution) seems to be a little better in some 
 instances. Very good results have been obtained by adding 
 about 5 cc. of I per cent, osmic acid to 100 cc. of chromo- 
 acetic acid (Schaffner's formula). A formula which gives satis- 
 factory results with Spirogyra may cause plasmolysis with 
 Cladophora. The given filament should be placed under the 
 microscope in the fixing agent, and, if plasmolysis occurs, the 
 
66 
 
 Methods in Plant Histology 
 
 Fig. 17. 
 A , Vaucheria geminata. 
 
 B 
 
 Vaucheria. 
 ^.V.sessilis. 
 
 chromic should be weakened or the acetic strengthened until the 
 suitable proportions are determined. This is a slow process, but 
 Cladophora and Vaucheria are almost sure to shrink without it. 
 About twenty-four hours in any of the chromic series and a four 
 to ten hours washing in water will be. sufficient for members of 
 
 this group. Only a few 
 o of the most familiar forms 
 will be mentioned. 
 
 Vaucheria. — This form 
 can always be obtained in 
 greenhouses, especially in 
 the fernery, where it forms 
 3 a green felt on the pots. 
 '''*^^°'"'- The greenhouse form is 
 
 likely to be Vaucheria sessilis. Another species, V. geminata, is 
 very common in the spring, when it may be found in ponds and 
 ditches. {^Fig. ly.^ It is extremely difficult to get mounts show- 
 ing the nuclei. The following method is sometimes successful : 
 
 1. Chromo-acetic acid (Schaffner's formula), 24 hours. 
 
 2. Wash in water, 4 to 10 hours. 
 
 3. Iron alum, 2 to 4 hours. 
 
 4. Water, 15 to 30 minutes. 
 
 5. One-half per cent, haematoxylin, over night. 
 
 6. Wash in water, 5 to ro minutes. 
 
 7. Iron alum until details become clear. This may take only a few minutes, 
 but may take an hour. 
 
 8. Wash thoroughly in water, i to 4 hours. 
 
 g. Ten per cent, glycerine and allow the glycerine to thicken. 
 10. Mount and seal. 
 
 Cladophora. — This is found attached to sticks and stones in 
 quiet or running water. It is easily recognized by its character- 
 istic branching. {^Fig. 18.^ The nuclei of the ccenocytic seg- 
 ments are readily brought out by the method just described for 
 Vaucheria. Alum carmine and Mayer's haem-alum are also good 
 stains for the nuclei. 
 
 Hydrodictyon. — This is popularly known as the "water-net." 
 Nets of all sizes should be selected for study. The segments 
 
Thallophytes. Algce 
 
 67 
 
 are coenocytic, and the nuclei are hard to differentiate except in 
 the younger segments. The method given for Cladophora yields 
 good results. The young nets forming within the older seg- 
 ments are easily demonstrated by Mayer's haem-alum. Paraffin 
 sections stained in cyanin and erythrosin, iron 
 alum-hgematoxylin, or the safranin-gentian violet- 
 orange combinationwill repay the trouble, especially 
 in case of the oldest segments within which new 
 nets are forming. The habit is beautifully shown 
 in preparations stained with eosin. The eosin ( i per 
 cent, aqueous solution) should act for about twenty- 
 four hours. Then transfer to i per cent, acetic 
 acid for a few minutes. If the stain comes out 
 rapidly in the acid, one minute may be sufficient, 
 but if the stain does not wash out, it is better to 
 let the acid act for four or five minutes. Wash 
 thoroughly in water to remove all trace of acid, 
 or the preparation will 
 fade. Transfer to 10 per 
 cent, glycerine and pro- Fixed. in chromo- 
 
 di / 7-"' \ acetic acid, Stained in 
 
 as usual. [Ftg.ig.) HaidenhaiVs iron 
 _ . rr^, . . alum-h3ematoxylin. 
 
 Spirogyra. — This is 
 probably the most widely known of all 
 the algae, and, fortunately, it is rather 
 easy to obtain beautiful and instructive 
 preparations. The following is a good 
 fixing agent for most Spirogyras; 
 
 Chromic acid, -^^ g. 
 
 Glacial acetic acid, -i\ g. 
 
 Water, 99 cc. 
 
 The addition of i cc. of i per cent, osmic acid seems to 
 improve it without causing any blackening. Flemming's weaker 
 solution is excellent. If it causes too much blackening, as it 
 probably will, the material, after being washed, should be placed 
 in weak peroxide of hydrogen (one part H^Oa to three parts 
 H2O) until the blackening due to osmic acid disappears. After 
 
 Fig 
 
 Cladophora. 
 
 Fig. 19. Hydrodictyon. 
 A small portion of a young net 
 
68 
 
 Methods i?i Plant Histology 
 
 a moment's washing in water it is then ready for the stain. The 
 iron alum-haematoxylin, as just described, brings out the nuclei 
 and pyrenoids with great distinctness, {^Fig. 20.) A few min- 
 utes in aqueous eosin after the last washing in water often gives 
 a beautiful differentiation, but the preparations will be quite 
 
 Fig. 20. Spirogyra. 
 
 From material fixed in chromo- acetic acid and stained in iron alum-haematoxylin. A 
 single cell showing nucleus, chromatophore, and pyrenoids 
 
 division. C, ; 
 
 , a nucleus undergoing 
 ng nucleus, D, zygospores, each showing two nuclei. 
 
 inferior if the eosin is allowed to stain too deeply, Mayer's 
 haem-alum is a better stain for stages in conjugation. 
 
 It is difficult to get Spirogyra into paraffin without shrinking, 
 .but it can*be done. Watch carefully and note where plasmolysis 
 
Thallophytes. Algm 
 
 69 
 
 occurs. There will probably be little or no trouble until 
 the transfer from 100 per cent, alcohol to the clearing agent. 
 Make this transfer as gradual as may be necessary. After the 
 pure xylol or other clearing agent is reached, add a lump of 
 paraffin large enough to saturate the clearing fluid at a tempera- 
 ture of 40 to 45° C. Allow the xylol to evaporate at this 
 temperature and imbed as usual, taking care to 
 keep the filaments as nearly parallel as possible. 
 Many elegant combinations, like cyanin and 
 erythrosin, fuchsin and iodine green, safranin- 
 gentian violet-orange, and others not available 
 for glycerine preparations, can be used with 
 paraffin sections. It is comparatively easy to 
 get any such alga into celloidin. Safranin and 
 Delafield's haematoxylin then make a good 
 combination. 
 
 Zygnema — Use the same methods as for 
 Spirogyra. In staining conjugating material the 
 stain should be extracted until the four chroma- 
 tophores of the zygospore become distinct. The 
 nuclei are comparatively small and unsatisfac- 
 
 . #-!-., , 11 , 1 ,1 11 The filament at the left 
 
 tory. Ihe stellate chromatophores are well shows three zygospores and 
 
 , , , , . -t r • 1 0"C parthenogenetic spore 
 
 brought out by alum carmme. If iron alum- which is distinguished by 
 
 having only two chromato- 
 
 haematoxylin and eosin are used, the eosin may phores. The fiiamem on the 
 
 •' •' right shows two cells, each 
 
 well be much deeper than in case of Spirogyra. with a pair of steiiate chro- 
 
 1 c oy matophores. Drawn from 
 
 (pier 2T\ material fixed in 2 per cent. 
 
 \J. i-^ . ^i.j formalin and stained in iron 
 
 Diatoms.— Diatoms and desmids have been -'""^h^-^-'o-y'*"- 
 variously classified, and their position is not yet fully determined. 
 Living diatoms are often found clinging in great numbers to fila- 
 mentous algse, or forming gelatinous masses on various submerged 
 plants. It is difficult to get really good preparations showing the 
 nucleus and chromatophores. If the diatoms are clinging to fila- 
 mentous algae, the algae with the diatoms attached may be put 
 into chromo-acetic acid (twenty-four hours), washed in water, 
 stained, passed up through the alcohols, and cleared in xylol, 
 or, better, in clove oil or bergamot oil, which do not dry up so 
 
 Fig. 21. Zygnema. 
 
70 
 
 Methods in Plant Histology 
 
 rapidly. Here the diatoms may be picked or scraped off from 
 the other algae, which will probably have become much shrunken 
 by this treatment. Mount in balsam. Haidenhain's iron alum- 
 haematoxylin is recommended for the nucleus and the centro- 
 some, which is quite prominent in diatoms. Delafield's haema- 
 toxylin and erythrosin give a good view of the nucleus and 
 
 chromatophore. If a glycer- 
 ine mount is preferred, the 
 iron alum-haematoxylin is a 
 good stain. 
 
 When the material is in 
 gelatinous masses, it may be 
 fixed in chromo-acetic acid 
 and imbedded in paraffin. 
 There will, of course, be some 
 difficulty in cutting, and many 
 frustules will be broken, but 
 there will, nevertheless, be 
 occasional views which show 
 details better than when the 
 diatoms are mounted whole. 
 
 Lriceratmm sp. ThC SlHcioUS Shclls Of dia- 
 
 toms are among the most beautiful objects which could be exam- 
 ined with the microscope {fig. 22) . To obtain perfectly clean 
 mounts requires considerable time and patience, but when the 
 ^material is once cleaned, preparations may be made at any time 
 with very little trouble. Diatom enthusiasts have devised numer- 
 ous methods for cleaning diatoms, and separating the various 
 forms from each other, but we shall give here only a few simple, 
 practical methods. 
 
 Material for mounts of frustules of living forms " may be 
 obtained by skimming off the brownish scum found on ponds, 
 by squeezing out water weeds, by scraping sticks and stones 
 which are covered at high water, or from the mud of filter beds 
 at pumping works, or in other places. The material is put in a 
 dish of water, and after it has settled the water is decanted. 
 
 Diatoms. X 255- 
 
 A^ Pleurosigma angulatum. B, Navicula dactylis. 
 C, Synedra biceps. D, Gomphonema sphaerophorum. 
 E,'tx\(. 
 
Thallophytes. Algce yi 
 
 This is repeated until the water will clear in about one-half hour.- 
 The sediment is then treated with an equal bulk of sulphuric 
 acid, after which bichromate of potash is added until all action 
 ceases. .After a couple of hours the acid is washed out. To 
 separate the diatoms, place the sediment in a glass dish with 
 water, and when the water becomes clear give the dish a slight 
 rotary motion. This will bring the diatoms to the top, when 
 they may be removed with a pipette and placed in alcohol. To 
 mount, place a number in distilled water, evaporate a few drops 
 of the mixture on a cover-glass, which is then mounted on a 
 slide in balsam." (From a review of Dr. Wood's paper on 
 " Diatoms," /^«^- App. Mic, March, 1899.) 
 
 Many scouring soaps and silver polishes contain large quan- 
 tities of fossil diatoms, and the diatomaceous earths are particu- 
 larly rich. Break up a small lump of such material and boil it 
 in hydrochloric acid. A test-tube is very convenient for this 
 process. Let the diatoms settle, pour off the acid, and then 
 wash in water. As soon as the diatoms settle, the water should 
 be poured off. The washing should be continued until the 
 hydrochloric acid has been removed. When the washing is 
 complete, pour on a little absolute alcohol, and after a few 
 minutes pour off the alcohol and add equal parts of turpentine 
 and carbolic acid. The material will keep indefinitely in this 
 condition and may be mounted in balsam at any time. In mak- 
 ing a mount, put a little of the material on a slide and allow it 
 to become dry, or nearly dry, and then add the balsam and cover. 
 If the balsam should be added too soon, the diatoms are likely 
 to move to the edge of the cover. 
 
 Desmids. — When these forms are very abundant, they may be 
 treated like the filamentous algs, except that extreme care must 
 be taken lest the desmids be lost while changing fluids. It 
 often happens that a single desirable desmid appears when 
 examining field collections. In such a case, remove it with a 
 fine pipette, and get it into a drop of water on a clean slide, 
 invert it over a bottle of i per cent, osmic acid for a minute, 
 leave the slide exposed to the air until the water has almost all 
 
72 
 
 Methods in Plant Histology 
 
 the 
 
 Evaporated, and then add a drop of lo per cent, glycerine. In 
 a few hours (six to twenty-four) put on a cover and seal. It 
 requires more time, care, and patience than it is worth to 
 attempt staining in such a case. {^Fig. 2j.) 
 
 Oedogonium. — In selecting material it will be better for teach- 
 ing purposes to choose the larger monoecious forms. The nuclei, 
 pyrenoids, and chromatophores are 
 easily differentiated. Mayer's haem- 
 alum is a good stain, especially for 
 the antheridia. Alum carmine or 
 eosin will bring out 
 {Fig. 24.) 
 
 Chara. — This form 
 is so large and coarse 
 that it hardly pays to 
 mount it in glycerine. 
 If a glycerine mount is ^y. 
 desired to show the an- 
 theridia and oogonia in 
 Fig. 23. Desmids. X 255. positiou, Spin a ring of 
 
 From glycerine preparations. Not stained. 
 A, Cosmarium pectinoides. B, Closte- CemCnt On thc slidc, 
 rium cucumis. C, Staurastrum comutum. 
 A Arthrodesmus octocornis. thuS making 3. Ccll in 
 
 which small portions of the plant may be mounted. 
 For parafifin sections select the tip of the plant, 
 a piece about half an inch in length. Sections of 
 this may show, not only the large apical cell, but 
 also various stages in the development of antheri- „ r, j 
 
 ° c Fig. 24. Oedogonium 
 
 dia and oogonia. Delafield's haematoxylin is a "oduiosum. 
 
 , . . , .- ,, 1 r 1 '^> antheridia. 0, 
 
 very good stam for the apical cell and for the oogonium. Drawn 
 
 . . rr^, horn material fixed in 
 
 development of antheridia and oogonia. The later i per cent, chromic 
 
 ^ " acid, and stained in 
 
 stages in the development of antherozoids are Mayer's hsem-aium. 
 brought out more clearly by the safranin-gentian violet-orange, 
 or by cyanin and erythrosin, but here unusual care must be taken 
 not to stain too deeply. 
 
 Good preparations showing shield, manubrium, capitula, and 
 filaments may be obtained by staining in bulk in alum carmine 
 
Thallophytes. AlgcB 
 
 73 
 
 and then crushing the antheridium under the cover-glass after 
 the specimen is in balsam. {^Fig. 2j.) 
 
 PHJEOPHYCEiE. 
 
 The brown algae are almost exclusively marine. The slime, 
 so prevalent in the group, often makes the technique difficult. 
 
 Ectocarpus. — Fix in 
 chromo-acetic acid 
 (twenty-four hours), 
 wash in fresh water, 
 since the salt of sea 
 water may cause incon- 
 venience in subsequent 
 processes. Stock mate- 
 rial should be passed up 
 
 A, portion of a branch showing an antheridium, a, and an 
 oogonium, o. B, median longitudinal section of an apical cell. 
 Drawn from a preparation fixed in chromo-acetic acid, and 
 stained in Delafield's haematoxylin. 
 
 to 70 per cent, alcohol for safe keeping. 
 Eosin or Mayer's haem-alum are good for 
 glycerine mounts. If paraffin sections are 
 to be made, the material must be brought 
 very gradually from absolute alcohol into 
 the clearing agent, and from the clearing 
 agent into the paraffin. {^Fig. 26.) 
 
 Other filamentous members of the 
 group, as well as the more delicate mem- 
 branous form.s, may be treated like Ecto- 
 carpus. 
 
 Fucus. — Fucus may be fixed, washed, and preserved like 
 Ectocarpus. It is difficult to get paraffin sections across the whole 
 fertile branch, but elegant sections may be obtained by cutting 
 
 Fig. 26. Ectocarpus confervoides. 
 
 From a preparation stained in 
 Mayer's haem-alum, and mounted in 
 glycerine. X 255. in, multilocu- 
 lar sporangium, w, unilocular spo- 
 
74 
 
 Methods in Pla?it Histology 
 
 narrow strips containing a few conceptacles. The safranin- 
 gentian violet-orange combination is good for such sections. For 
 such views as are represented in fig. 2y, C and D, the material 
 
 should be stained 
 in bulk in borax 
 carmine or alum 
 carmine. The 
 process for borax 
 carmine is as fol- 
 lows : 
 [. Borax carmine, 24 
 
 hours. 
 3. Acid alcohol (2 
 drops of HCl in 
 50 cc. of 70 per 
 eent.alcohol),until 
 the color becomes 
 a clear red. This 
 may take an hour 
 or even a day. 
 
 3. Seventy to loo per 
 cent, alcohol, 2 
 hours each. 
 
 4. Clear in cedar oil, 
 bergamot oil, or 
 oil of cloves. 
 
 5. Tease out the con- 
 tents of the con- 
 ceptacles suffi- 
 ciently to show 
 details, and mount 
 in balsam. 
 
 Fig. 27. Fucus vesiculosus. 
 
 A, small portion of plant showing bladders and fruiting branches. 
 One-half natural size. B, transverse section of fruiting branch showing 
 oogonial conceptacles. X 6. C, antheridia and paraphyses. From a 
 preparation fixed in chrorao- acetic acid, stained in borax carmine, teased 
 out and mounted in balsam. X 255. D, oogonium showing five of the 
 eight oospheres. Prepared as in C. 
 
 The process for alum carmine is the same, except that no 
 acid alcohol is used. 
 
 Sections like that shown in B are easily cut in celloidin. 
 After staining in borax carmine or alum carmine, imbed in 
 celloidin in the usual way. After hardening the celloidin in 
 chloroform, put the block into 95 per cent, alcohol for two or 
 three hours, and then into Eycleshymer's clearing fluid (equal 
 parts bergamot oil, cedar oil, and carbolic acid), until thoroughly 
 cleared. The block may be left here indefinitely, and sections 
 
Thallophytes. AlgcB 
 
 75 
 
 B 
 
 may at any time be mounted in balsam as soon as they are 
 cut. 
 
 Chorda, Lamifiaria, and similar forms may be treated like Fucus. 
 
 RHODOPHYCEiE. 
 
 The red algae belong almost exclusively to salt water, -but a 
 few genera are found only in fresh water, usually in running water, 
 and a few forms occur both ^ 
 in salt and in fresh water. 
 
 The technique is more 
 difficult than in the case of 
 the brown algae. Until some- 
 thing better is suggested, the p 
 same method of fixing and 
 washing may be used as for 
 the brown algae. Picric acid, 
 corrosive sublimate, and ab- / 
 solute alcohol have been 
 tried, but do not give as good 
 results as the chromo-acetic ^ 
 acid or Flemming's fluid. 
 
 Batrachospermum. — This 
 is a green, fresh-water mem- 
 ber of the red algae. It is 
 not very uncommon in small 
 streams. {Fig. 28.) 
 
 TViP- rf^llc ar<^ cr. email branches and several cystocarps. X 25. ^, a procarpib 
 
 i HC CCllb die bU Sllldll branch showing carpogonium (/), and trichogyne (i'), 
 
 .1 , ., • 1 11 ,1 1-1 with an antherozoid (j) attached. X 255. C, a younger 
 
 tnat It is naraly WOrtn Wnlle branch showing carpogonlum and trichogyne. X 255. 
 
 ... , , D, branch with three antherozoids. X 255. 
 
 to attempt sectionmg them. 
 
 Very good preparations showing the nuclei may be obtained by 
 staining in Mayer's hsem-alum, or Haidenhain's iron alum-haema- 
 toxylin. After the material is in glycerine ready for mounting, 
 tease out a small portion, and still further dissociate the filaments 
 by tapping smartly on the cover. 
 
 Material stained in eosin shows the external structure well, 
 but may not bring out the nuclei. 
 
 Fig. 28, Batrachospermum moniliforme. 
 
 From a preparation stained in Mayer's haem-alum and 
 mounted in glycerine. A, portion of plant showing 
 
76 
 
 Methods in Plant Histology 
 
 Polysiphonia. — For preparations like those shown in fig. 2g 
 eosin is a very good stain. To get a brilliant coloring, stain 
 
 for about twenty-four hours, 
 so that the i per cent, acetic 
 acid may be allowed to act 
 for several minutes without 
 making the stain look weak. 
 Wash thoroughly in water. 
 Not the slightest trace of 
 color should be allowed to 
 come out in the glycerine. 
 
 Sections showing the cen- 
 tral and peripheral siphons 
 and other gross features are 
 easily cut in celloidin. It is 
 not very difficult to cut par- 
 affin sections, but the nuclei 
 are so small and so hard to 
 bring out that such prepa- 
 rations had better be left for the specialist. 
 
 Fig. 29. Polysiphonia fibrillosa. 
 
 From a preparation fixed in chromo- acetic acid, stained 
 in eosin. and mounted in glycerine. X 255. A, an 
 antheridium. B, a cystocarp with carpospores. C, 
 tetrasporic branch with tetraspores. 
 
CHAPTER XIL 
 
 THALLOPHYTES. FUNGI. 
 
 SCHIZOMYCETES. 
 Bacteria. — The methods of modern bacteriological technique 
 are so numerous and so specialized that we must refer to laboratory 
 manuals for instruction in this subject. The method given here 
 will merely enable the student to study 
 the form and size of those bacteria d't^^'^i 
 
 which are more easily demonstrated. ''^^x-' ■^■^^*^|^"' 
 
 Foul water at the outlets of sewers / >^ ^ •^ so- '^^. 
 
 and such places will usually afford an X. 4. "^ "^ ^)^K 
 
 abundance of coccus, bacillus, spiril- f^^ \ ® ^''' ^^ 
 
 • R "^JJ" C 
 
 lum, and beggiatoa forms. Place a 
 drop of the water on a slide, heat it 
 gently until the water evaporates, then 
 
 stain with fuchsin or methyl violet, f^^"^ ^^^k^ 
 
 dehydrate, clear in xylol, and mount 
 
 in balsam {fig. so)- C 3 ^"-J/ " C 
 
 Fine preparations may be obtained fig. 30. Bacteria. X535. 
 
 by inoculating a mouse with Anthrax ^.Badllusanthracis, from a paraffin 
 
 •' " section cut from the liver of a mouse. 
 
 or some other form, and then cutting ^^^'^^ZT^^^^^^'^ 
 paraffin sections of favorable organs. cTsTolin^str^uV^Ffor a^S-^ 
 Gentian violet with a faint bismark SumTp' 'Frfm"T;rS:ratio''n 
 
 1 r i_ 1 11 1 stained in fuchsin. 
 
 brown tor a background makes a good 
 
 combination. The following schedule gives good results with 
 
 Anthrax and many other bacteria : 
 
 1. Gentian violet, 5 minutes. 
 
 2. Rinse in water a few seconds. 
 
 3. Gram's solution (iodine 1 gram, potassium iodide 2 grams, water 300 cc.) 
 until the color is almost or quite black ; this will generally require i or 
 2 minutes. 
 
 4. Ninety-five per cent, alcohol until the color has nearly disappeared. 
 
 77 
 
78 Methods in Plant Histology 
 
 5. Rinse in water and examine. If the bacteria are well stained, a counter- 
 stain for the background may be added. 
 
 6. Erythrosin, 3 or 4 seconds ; or bismark brown 5 or 10 seconds. 
 
 7. Ninety-five and 100 per cent, alcohol, dehydrating as rapidly as possible. 
 Not more than 5 or 10 seconds can usually be allowed. 
 
 8. Xylol. 
 
 9. Balsam. 
 
 Leptothrix may often be obtained by scraping the inside of 
 the cheek. Beggiatoa, a form with oscillating movements like 
 Oscillaria, is often found in foul water. Its presence may be 
 indicated by whitish patches on the bottom. 
 
 It is doubtful whether the bacteria possess even a morpho- 
 logical forerunner of the nucleus of higher plants, consequently 
 there need be no disappointment if the larger forms, like some 
 of the Beggiatoas, fail to show a nucleus. 
 
 MYXOMYCETES. 
 
 With the exception of a few forms like Fuligo (often found 
 on oak stumps and on oak bark in tan yards), the Myxomycetes 
 are small, and are usually overlooked by collectors. ^Fig.ji.) A 
 
 careful examination of rotting logs 
 in moist woods will usually reveal 
 an abundance of these delicate 
 and beautiful organisms. They 
 should be pinned to the bottom 
 of a box for safe carrying. For 
 herbarium specimens they are 
 simply allowed to dry, and are 
 C then fastened with glue or paste 
 
 Myxomycetes. ^^ ^j^^ bottOm of a Small boX. 
 
 Growing on rotten wood, /i , Hemitrichia rubi- 
 
 formis. X 20. B, Stemonitis ferruginea. Nat- Plasmodia and voung SDoran- 
 
 ural size. C, Trichia varia. Xi%. . . . 
 
 gia may be fixed in chromo-acetic 
 acid or Flemming's fluid. Sections are easily cut in paraffin, 
 and should not be more than 5 ft in thickness, and should be 
 thinner, if possible. Acid fuchsin and iodine green is a good 
 stain. Delafield's haematoxylin used alone or with a little 
 orange G. is also to be recommended. , Excellent methods for 
 
Thallophytes. Fungi 
 
 79 
 
 living cultures were given in the January and February (i 
 numbers of 1\\q Jour?ial of Applied Microscopy. 
 
 PHYCOMYCETES. 
 
 Mucor. — This familiar mold appears with great regularity on 
 bread. The following is a sure and rapid method for obtaining 
 Mucor: Place a glass tumbler in a plate of water, put on the 
 tumbler a slice of bread which 
 has been exposed to the air for a 
 day, and cover with a glass jar. 
 
 To obtain such a series as is 
 shown in A-D of fig. J2, the ma- 
 terial should be studied before 
 the sporangia begin to turn black. 
 The phase in the life-history indi- 
 cated in F-// is rarely seen. The 
 writer would be glad to hear from 
 any who have met this phase, 
 especially if the information could 
 be accompanied by a few zygo- 
 spores. 
 
 Ci , . . / A-D, successive stages in the development 
 
 OrrOSlVe sublimate (^4 per of the sporangium. Drawn from living material. 
 
 , . . . . E, columella with a few spores adhering. F-H, 
 
 cent.] m 50 per cent, alcohol, stages in the formation of the zygospore. From 
 
 a preparation fixed in corrosive sublimate, 
 
 used hot, may be recommended stained in Delafleld's hsematoxylln, and mounted 
 
 •' in glycerine. 
 
 as a fixing agent. Haem-alum, or 
 
 Delafield's haematoxylin, is good for glycerine preparations. 
 Do not stain too deeply. A very satisfactory study may be 
 made from the living material. 
 
 Cystopus. — This fungus is quite common on Cruciferae, where 
 the white "blisters" or "white rust" form quite conspicuous 
 patches. 
 
 Affected portions of leaves and stems should be fixed in 
 chromo-acetic acid and cut in parafifin. Sections 5 /* or less in 
 thickness will be found most satisfactory. Safranin-gentian 
 violet-orange seems to be the best stain for differentiating the 
 nuclei. {Fig. 33.) 
 
 Fig. 32. Mucor stolonifer. X 255. 
 
8o Methods i?i Plant Histology 
 
 It is more difficult to get good sec- 
 tions of the plant in the oosporic condi- 
 tion. The oosporic phase of Cystopus bliti 
 is easily recognized on Antarantus, where 
 the oospores may be seen with the naked 
 eye by holding the leaf up to the light. 
 While better nuclear staining can be se- 
 cured with chromic or Flemming mate- 
 rial, it will be found somewhat easier to 
 cut material which has been fixed in pic- 
 ric acid (i per cent, solution in 70 per 
 cent, alcohol). Celloidin sections, stained 
 in Delafield's haematoxylin, can be rec- 
 "^^^oT^Tpf^^c ommended for showing the position of 
 
 Fig. 33. Cystopus candidus .. . , , i • i • 1,1 1 1 
 
 on Capseiia. oogonia and anthendia, although such sec- 
 
 leaT^Ti^s'Ftma'X-atlon' ^ions are too thick to givc satisfactory views 
 
 fixed in Flemming's fluid and f i.V-„ nnrlpi 
 stained in safranin-gentian vio- *-"■ '■**'- iiui-ici. 
 let-orange. 
 
 ASCOMYCETES. 
 
 This group, popularly known as the "sac fungi," contains an 
 immense number of saprophytic and parasitic forms. Yeast, 
 green mold on cheese and leather, leaf curl of peach, black 
 knot of cherry and plum, and the powdery mildews are familiar 
 to everyone. The few objects selected will enable the student 
 to experiment, but he must not be discouraged if success does 
 not crown the first attempt, for the group presents many diffi- 
 culties. 
 
 Saccharomyces. — Until somewhat recently it was considered 
 rather difficult to demonstrate the nucleus of the yeast cell. 
 With fresh growing yeast the following method by Wager 
 should be successful : Fix in a saturated aqueous solution of 
 corrosive sublimate for at least twelve hours. Wash success- 
 ively in water, 30 per cent, alcohol, 70 per cent, alcohol, and 
 methyl alcohol. Place a few drops of alcohol containing the 
 cells on a cover, and when nearly dry add a drop of water. 
 After the yeast cells settle, drain off the water and allow the 
 
Thallophytes. Fufigi 
 
 8i 
 
 cells to dry up completely. Place the cover, or slide, with its 
 layer of cells in water for a few seconds, and then stain with a 
 mixture of fuchsin and methyl green, or fuchsin and methylen 
 blue. Mount in glycerine or in balsam. 
 
 Eurotium. — For class use or for permanent preparations it is 
 best to select 
 rather y oung£^f^. 
 material which *'"'' 
 shows various 
 stages in devel- 
 opment, from the 
 swollenendof the 
 hypha to the ripe 
 spore [fig. 34) . 
 
 C D 
 
 F;g. 34. Eurotium. 
 
 From material growing on a hectograph pad. Fixed in chromo-acetic 
 , acid, stained in eosin, and mounted in glycerine, ^-^'j successive stages in 
 
 The nuclei are development, x 375- 
 
 exceedingly small, and can hardly be demonstrated with eosin. 
 
 Iron alum-haematoxylin would be better. 
 
 The following schedule will give good mounts for habit study: 
 
 1. Fix in a saturated solution of corrosive sublimate in 50 per cent, alcohol. 
 The addition of i cc. of glacial acetic acid to 100 cc. of this solution 
 improves it. Use it hot. 
 
 2. After it cools, transfer to 50 per cent, alcohol and add, a few drops at a 
 time, the iodine solution which is used in testing for starch. At first the 
 brownish color caused by the iodine will disappear, but after a certain 
 amount has been added the brown color will remain. The material is 
 then ready for the next step. 
 
 3. Thirty-five per cent, alcohol, 5 minutes. 
 
 4. Eosin (aqueous), at least 5 minutes ; a day will do no harm. 
 
 5. Put the material into i per cent, acetic acid for 15 seconds to 2 minutes. 
 The material should still have a vivid red color when taken from the acid. 
 
 6. Wash in water. Use a considerable volume of water or change the water 
 several times. If the acid is not all washed out, the preparations will fade. 
 
 7. Ten per cent, glycerine, and allow the glycerine to thicken. 
 
 8. Mount in glycerine or glycerine jelly, and seal. 
 
 Any of the filamentous fungi — like Mucor, Thamnidiutn, 
 Peronospora, Penicillium, Pythium, Saprolegnia, etc. — can be 
 mounted in this way. Saprolegnia, however, is much more satis- 
 factory if stained in iron alum-haematoxylin. 
 
82 
 
 Methods in Plant Histology 
 
 A very rapid method for the unicellular and filamentous 
 forms may be added : 
 
 1. One hundred per cent, alcohol, 2 minutes. 
 
 2. Eosin (aqueous), 2 minutes. 
 
 3. One per cent, acetic acid, 2 to 10 seconds. 
 
 4. Wash in water 5 minutes, changing frequently. 
 
 5. Mount directly in 50 per cent, glycerine, and seal. 
 
 If the material gets through the first four stages without 
 shrinking, but collapses at the fifth, put it into 10 per cent. 
 
 glycerine and allow it to 
 thicken as usual. 
 
 Uncinula necator. — 
 The mildews are found 
 throughout the summer 
 and autumn on the leaves 
 of various plants. The 
 lilacmildew( MicrosphcBra 
 alni^ and the mildew on 
 the Virginia creeper 
 {^Uncinula necator) are 
 particularly abundant. 
 For herbarium purposes 
 they may be preserved 
 by simply drying the 
 leaves under light pres- 
 sure. When mounted for 
 examination, the leaf 
 
 Fig. 35. yl, Uncinula necator on Ampelopsis quinquefolia. should bc SOakcd in WatCr 
 
 X 192. Four asci containing ascospores have been forced out - . • l Ct. 
 
 by pressing out the cover. Fixed in hot corrosive sublimate, lOr 3. iCW mmUtCS, alter 
 stained in fuchsin, and mounted in balsam. /?, a conidiospore ; , . , . 
 
 and C, an appendage of Microsphaera alni, drawn from living whlch the perithCCia may 
 material. X 192. '■ •' 
 
 be scraped off and 
 mounted in water. The asci may then be forced out by press- 
 ing smartly on the cover. {Fig. 35.) 
 
 For permanent mounts of entire perithecia with appendages, 
 fix in 3 per cent, formalin twenty-four hours, wash in water one 
 hour, stain in aqueous eosin twenty-four hours, treat with i per 
 
 JOUR APR MIC. 
 
Thallophytes. Fungi 
 
 83 
 
 cent, acetic acid one minute, wash thoroughly in water, and then 
 transfer to 10 per cent, glycerine, which should be allowed to 
 concentrate as usual. If chromic acid, corrosive sublimate, or 
 alcohol be used for fixing, the appendages become brittle, and 
 very easily break off. However, the chromo-acetic mixtures 
 are better if it is desired to make parafifin sections showing the 
 development of the perithecium with its asci and spores. For 
 this purpose the omnipresent 
 Erysiphe commune on Polygonum 
 aviculare is exceptionally favor- 
 able, because, after the material 
 is fixed and in alcohol, the whole 
 mycelium, with the developing 
 perithecia, may be stripped from 
 the leaf without the slightest 
 difficulty, thus avoiding the 
 necessity of cutting the leaf in 
 order to get the fungus. The 
 safranin- gentian violet -orange 
 combination seems to give the 
 best results, although cyanin 
 and erythrosin are quite satis- 
 factory when the 
 properly balanced. 
 
 ■V—l^-i- TV/r p i-1 several days in equal parts of gS per cent, alcohol 
 
 Aylaria. Many forms, like and glycerine, and then imbedded in celloidin. Not 
 
 Xylaria, Ustilina, Hypoxylon, and 
 
 Nummularia, in their mature condition, are woody and so 
 extremely brittle that it is almost impossible to cut them. As 
 good a plan as any seems to be to cut sections of the stroma 
 about one-eighth of an inch thick, soak them in equal parts of 
 glycerine and 95 per cent, alcohol, and then imbed them in 
 celloidin in the usual way. They might be cut without imbed- 
 ding, but most of the asci and spores would then be lost. 
 
 The younger stages, showing the development of perithecia 
 and asci, are more interesting, and can be cut in paraffin and 
 stained with ease {fig- 36). 
 
 A , transverse section of a young stroma showing 
 stains are PS"*hecia. X 8. Fixed in chromo-acetic acid, 
 stained in bulk in alum carmine, imbedded in celloi- 
 din, and mounted in balsam. B, two asci with 
 spores. X 245. The mature stroma was soaked for 
 
Methods i?i Plant Histology 
 
 Peziza. — The Pezizas and related forms are fleshy, and pre- 
 sent but little difficulty in fixing, cutting, or staining. They are 
 abundant in moist places, on decaying wood, 
 or on the ground. The apothecia have the 
 form of little cups, which are sometimes 
 black and sometimes flesh -colored, but 
 often orange, red, or green. For the devel- 
 opment of ascospores in the ascus, Flem- 
 ming's fluid (weaker solution), followed by 
 safranin-gentian violet-orange, has given the 
 best results with thin sections where the 
 mitotic figures are to be studied. Cyanin 
 and erythrosin is also to be recommended. 
 Such sections should not be- more than 
 5 /* in thickness. For a general morpho- 
 logical preparation, such as is shown in 
 the figure, it is better to stain in bulk in 
 alum carmine or in Delafield's haematoxy- 
 lin, and then tease out the asci in gly- 
 cerine or balsam. Sections thick enough 
 to show the entire ascus are not usually 
 as satisfactory as such teased preparations. 
 {Fig- 37-) 
 
 Fig. 37. Peziza odorata. 
 Three asci and many para- 
 physes. X 245. Fixed in cor- 
 rosive sublimate, stained in 
 bulk in alum carmine. Teased 
 out and mounted in balsam. 
 
 ^CIDIOMYCETES. 
 
 The .^cidiomycetes comprise the rusts [Uredine<^) and the 
 smuts [UstilaginecB). 
 
 Puccinia graminis. — The common rusts of wheat and oats 
 are familiar to everyone. The uredospores, or summer spores, 
 known as the red rust, and the teleutospores (last spores), or 
 winter spores, known as the black rust, are found in unfortunate 
 adundance, but the aecidium stage on the barberry is not neces- 
 sary for the vigorous development of rust in the United States, 
 and is seldom found. Most teachers are obliged to depend upon 
 botanical supply companies for this material. There are, how- 
 ever, various aecidia which are as good, or even better, for 
 
Thallophytes. Fungi 
 
 &5 
 
 morphological study. The aecidia growing on Euphorbia maculata 
 (spotted spurge), and on AriscEma triphy Hum (Jack-in-the-pulpit) 
 are much easier to cut, and seem easier to stain. Delafield's 
 haematoxylin, followed by a very light stain in erythrosin, is 
 
 tl/OURAPBMie.. 
 
 Fig. 38. Puccinia graminis. 
 
 _A, transverse section of barberry leaf showing secidia and spermagonia. X 7. B, longitudinal 
 section of a single aecidium. X 192. Fixed in Flemming's weaker solution and stained in Delafield's 
 haematoxylin. C, a single sperraagonium. X 192. Fixed and stained as in B. D, three uredospores 
 growing on oats. X 375. Fixed in 2 per cent, formalin, stained in bulk in alum carmine, and teased 
 out in glycerine. E, section of young teleutospores on oats. X 375. Fixed in picro-acetic acid and 
 stained in cyanin and erythrosin. G, F, //, three ripe teleutospores from a leaf of oats showing varia- 
 tion in form. X 375. /. germinating teleutospores. X 375. 
 
I I ^ I I 
 
 Fig. 39. 
 
 86 Methods in Plant Histology 
 
 good for both aecidia and spermagonia, especially after Flem- 
 ming's fluid. It is rather difficult to get good sections of uredo- 
 spores and teleutospores, because the leaves of wheat and oats 
 are refractory objects to cut. The cutting is easier after picro- 
 acetic acid than after corrosive sublimate or the chromic-acid 
 series. {Fig- jS.) 
 
 Every class which studies the rusts should attempt to germi- 
 nate the uredospores and teleutospores. For this purpose the 
 hanging drop culture may be employed. Cement a rubber or 
 zinc ring to the slide, or simply smear the lower surface of the 
 ring with vaseline and press it tightly against the slide ; smear 
 
 the upper surface of the ring 
 with vaseline, and over it invert 
 ID the cover-glass with a shallow 
 drop of water containing the 
 spores {fig- 39). The uredo- 
 spores germinate readily all summer, but it is said that the 
 teleutospores will germinate only in the spring following their 
 maturity. However, the teleutospores of many species, like 
 Puccinia xanthii on Xaftthium canaderise (cocklebur), will germi- 
 nate as soon as they ripen and will serve equally well for study. 
 If a particularly good specimen is secured, it may be preserved 
 by the method previously described for desmids, except that in 
 this case it might be worth while to attempt staining with Mayer's 
 haem-alum, or with eosin. 
 
 The smuts may be studied in the living material. The fol- 
 lowing method, recently described by Ellis, is worth remember- 
 ing : A supply of smutted barley may be obtained by sowing 
 soaked, skinned barley that has been plentifully covered by 
 Ustilago spores. In such material it is easy to trace stages in 
 the development of spores. Free-hand sections of ears about 
 three-eighths of an inch long show the mycelium and spore clus- 
 ters. If smutted ears be removed and kept floating on the 
 water, the spores continue to develop and often germinate. For 
 paraffin sections desirable stages should be fixed in Flemming's 
 fluid or picro-acetic acid. Delafield's haematoxylin, followed by 
 
Thallophytes. Fungi 87 
 
 a very light touch of erythrosin or acid fuchsin, will give a good 
 stain. 
 
 For a study of the germinating spores and conidia, cultures 
 may be made in beerwort on the slide or in watch crystals. 
 Harper's method of making preparations from such material is 
 ingenious and will undoubtedly prove valuable in making mounts 
 of various small plant and animal forms, A drop of the material 
 is taken up with a capillary tube and is then 
 gently blown out into a drop of Flemming's 
 weaker solution (fifteen minutes or an hour was 
 sufficient for the fungus spores). Cover a 
 slide with albumen fixative, as if for sections. 
 A drop of the material, without previous wash- 
 ing, is drawn up into the capillary tube and 
 touched lightly and quickly to the surface of 
 the albumen. A series of such drops, almost 
 as small as the stippled dots in a drawing, may 
 be applied to the slide. The fixing agent may 
 
 Transverse section of a 
 
 now be allowed to evaporate somewhat, but portion of one of the giiis 
 
 * snowing a part of the trama, 
 
 the preparation must not be allowed to dry. ''• and several basidia ,j, 
 
 r r J each with four sterigmata, 
 
 As the slide is passed rapidly through the t^^^^ly^TAf^Jl 
 alcohols, the albumen is coagulated, and the ™*''' ^"°* 
 preparation may be treated just as if one were dealing with 
 ribbons of sections. 
 
 BASIDIOMYCETES. 
 
 This is an immense group, of which the mushrooms, toad- 
 stools, puffballs, and bracket fungi are the most widely known 
 representatives. 
 
 Coprinus comatus. — This is the common shaggy-mane mush- 
 room. Cut from the cap pieces about one-fourth of an inch 
 square, and fix in chromo-acetic acid or in Flemming's fluid. Por- 
 tions in which the gills have just begun to turn brown will show the 
 spores still attached to the sterigmata {fig. 40) . If the gills have 
 become dark brown or black, the spores will wash off before the 
 sections can be mounted. Look in portions in which the gills 
 are still white or only slightly changing color for the develop- 
 
88 Methods in Pla?it Histology 
 
 ment of basidia and spores. The nuclei, although rather small, 
 are brought out nicely by safranin-gentian violet-orange. The 
 same procedure may be observed for other forms of similar con- 
 sistency, like many members of the genera Boletus, Hydnum, 
 Polyporus, Lycoperdon, etc. Leathery or woody forms like Stereum 
 and many species of Polyporus had better be fixed in picro-acetic 
 acid and imbedded in celloidin. Young stages of Cyathus or 
 Crucibulum (bird's-nest fungi) cut easily in paraffin, but the 
 older stages cut much better in celloidin. It is hard to get the 
 very soft, watery forms like Tremella into paraffin without shrink- 
 ing, but sections as thin as lo /u- may be cut in celloidin. While 
 this is too thick to give satisfactory views of such small nuclei, 
 it brings out very clearly the general morphological structures. 
 
 THE LICHENS. 
 
 The lichens are usually regarded as diflficult forms. In 
 younger stages they occasion no trouble, but an old apothecium 
 or a leathery thallus often fails to cut well. Difficulties may be 
 minimized by using prolonged periods. The following schedule 
 has proved satisfactory for the thalli and mature apothecia of 
 Physcia, Usnea, Sticta, Collema, Parmelia, and Peltigera: 
 
 1. Chromo-acetic acid (medium solution, p. 28), 2 to 4 days. 
 
 2. Wash in water, 6 to 24 hours. 
 
 3. Thirty-five, 50, 70, 85, and 95 per cent, alcohols, 6 to 24 hours each. 
 
 4. One hundred per cent, alcohol, 2 to 4 days, changing 2 or 3 times. 
 
 5. Mixtures of alcohol and xylol, i to 2 days. 
 
 6. Pure xylol, 6 to 24 hours. 
 
 7. Xylol and parafifin on the bath, i to 2 days. 
 
 8. Parafifin at 54° to 60", changing once or twice, 3 to 6 days. 
 
 9. Imbed in as thin cakes as possible. 
 
 Cyanin and erythrosin is a very good stain for lichens. The 
 algae stain blue and the filaments of the fungus take the red. 
 Where the association of the alga and the fungus is rather loose, 
 as in Dichonema, more satisfactory mounts can be made by stain- 
 ing in eosin, or haem-alum and eosin, and then teasing slightl}' 
 with needles and mounting in glycerine. 
 
CHAPTER XIII. 
 
 BRYOPHYTES. 
 
 The Bryophytes, comprising the two groups Liverworts 
 [Hepaticcs) and Mosses [Mtisci] , present a great diversity of 
 structure, some being so delicate that good preparations are very 
 uncertain, while others are so hard that it is difficult to get satis- 
 factory sections. Between these extremes, however, there are 
 many forms which readily yield beautiful and instructive prepa- 
 rations. 
 
 If but one fixing agent should be suggested for the entire 
 group, it would be chromo-acetic acid with ^ g. chromic acid 
 and ^ cc. acetic acid to lOO cc. of water. - It should be allowed 
 to act for at least twenty-four hours, and probably two or three 
 days would be better. Always make an effort to get the mate- 
 rial into paraffin, using celloidin only as a last resort for refrac- 
 tory structures which resist infiltration and for very delicate 
 structures which persist in collapsing. As one gains in experi- 
 ence and carefulness, the number of cases which seem to demand 
 celloidin will become fewer and fewer. 
 
 Instead of treating forms in a taxonomic sequence, we shall 
 consider first the gametophyte structures under the headings 
 thallus, antheridia, and archego?iia, and shall then turn our atten- 
 tion to the sporophyte. 
 
 HEPATIC-S:. 
 
 Some of the liverworts are floating aquatics, but most of 
 them grow on logs or rocks or upon damp ground. They are 
 found at their best in damp, shady places. Many of them may 
 be kept indefinitely in the greenhouse. Riccia, Ricciocarpus, 
 Marcha?itia, Cofiocephalus, Asterella, and many others vegetate 
 luxuriously, and often fruit if kept on moist soil in a shady part 
 of the greenhouse, and they do fairly well in the ordinary labo- 
 ratory if covered with glass and protected from too intense 
 
 89 
 
90 
 
 Methods in Plant Histology 
 
 Fig. 41 
 A, longitudinal. 
 
 Ptilidium ciliare. X 420. 
 8, transverse section of the apex of the 
 
 leafy gametophyte. Fixed in Flemming's weaker solution, 
 stained in a mixture of acid fuchsin and iodine green. Ten 
 
 light. The living plants are very desirable, since they not only 
 furnish the best possible material for habit work and the coarser 
 microscopic study, but they also enable one to secure complete 
 series in the development of the various organs. 
 
 The Thallus. — In many cases it will not be necessary to make 
 a special preparation for the study of the thallus, since prepara- 
 tions of antheridia, archegonia, or sporophytes may include good 
 
 sections of vegetative por- 
 tions. This is particularly 
 true of forms like Riccia, 
 where the various organs 
 are not raised above the 
 thallus. In forms like J/<2r- 
 chantia, where the antheri- 
 dia, archegonia, and sporo- 
 phytes are borne upon 
 stalked receptacles, it is 
 better to make separate preparations to show the structure of 
 the mature thallus. Sections intended to show the structure 
 of the mature thallus should be 15 ft to 25 /x in thickness, but 
 sections to show the growing point and development of the 
 thallus should not be thicker than 10 /u.. Material showing apical 
 cells and development of the thallus is easily gotten into paraf- 
 fin, even in forms like RicciocarpuSy which in their mature condi- 
 tion are in danger of collapsing. The apical region of the foliose 
 Jungermanniacecs {fig. 41) affords an excellent opportunity for 
 studying the development of the plant body from a single apical 
 cell. If mixtures containing osmic acid are used for fixing, 
 there may be difficulty in the staining, even after using per- 
 oxide of hydrogen. Corrosive sublimate-acetic, Carnoy's fluid, 
 or chromo-acetic acid are better for apical regions. A fairly 
 vigorous staiiiing with a mixture of acid fuchsin and iodine 
 green often brings the walls out very sharply. Chromo-acetic 
 acid, followed by Delafield's haematoxylin or bismark brown, is 
 good for the apical cells and developing regions, but a light 
 counter-stain with erythrosin improves preparations of the 
 
Bryophytes 
 
 mature thallus. After corrosive sublimate-acetic the material 
 maybe stained in bulk with alum cochineal or alum carmine, 
 thus giving fairly good preparations and saving considerable 
 labor. 
 
 Antheridia. — If you have the material, it is not difficult to 
 get good preparations showing the development of antheridia. 
 In forms like Asterella, Pellia, etc., cut out a small portion of the 
 thallus bearing the antheri- 
 dia. The piece should not 
 be more than a quarter of 
 an inch square, and if it can 
 be smaller, so much the 
 better. For early stages of 
 the antheridia of Marchan- 
 tia select young antheridio- 
 phores which still lie close 
 to the thallus. These 
 readily cut as thin as 5 /*, 
 and a single slide will usu- 
 ally show a more complete 
 series than is represented in the figure of Asterella {fi^. 42), but 
 after the stalk begins to lengthen, the younger stages become 
 infrequent, and it is not always easy to cut thin sections. Dela- 
 field's haematoxylin or bismark brown serves very well for such 
 stages as are shown in the figure. The protoplasm of the young 
 antheridia is so dense that the addition of a counter-stain is 
 almost sure to injure the preparation by obscuring the cell walls. 
 For stages older than that represented in D, showing the devel- 
 opment of the spermatozoid, the paraffin must be rather hard 
 (melting at 55° C. to 65° C.) , and the sections should not be 
 thicker than 5 /a, while 2 /it or 3 /* is best. For such stages use 
 the safranin- gentian violet -orange combination, Haidenhain's 
 iron alum-haematoxylin with or without a faint trace of eryth- 
 rosin or orange G, or use a mixture of acid fuchsin and methyl 
 green. Nothing but practice and patience will bring success in 
 such critical work. 
 
 Successive stages in the development of antheridia. 
 Fixed in chromo-acetic, stained with Delafield's haema- 
 toxylin. Section lo microns thick. 
 
92 
 
 Methods i?i Plant Histology 
 
 If antherozoids are found escaping, transfer them to a small 
 drop of water on a clean slide, invert the drop o\'er a i per cent, 
 solution of osmic acid for two or three minutes, allow the drop 
 to dry up, pass the slide through the flame two or three times, 
 as in mounting bacteria, and then stain sharply in acid fuchsin. 
 
 This should show the 
 general form of the an- 
 therozoid, and will usu- 
 ally bring out the cilia. 
 
 The Archegonia. — The 
 methods for archegonia 
 are practically the same 
 as for antheridia. Too 
 much stress cannot be 
 laid upon the importance 
 of carefully selecting the 
 material. Use very small 
 pieces, and, before pla- 
 cing them in the fixing 
 agent, trim them to such 
 a shape that the position 
 of the archegonia will be 
 accurately known even 
 after the pieces are imbedded in paraffin. For stages like^^. 4J, 
 A and B, Delafield's haematoxylin is a good stain, and lo /u is 
 about the right thickness. For stages like C, in such forms as 
 Marchantia, where the necks are long and often somewhat curved, 
 it is better for general purposes to use sections from 1 5 yit to 
 20 [i in thickness. If it is desired to obtain preparations show- 
 ing the cutting off of the ventral canal cell, the development of 
 the oosphere, and the process of fertilization, the sections should 
 be from 5 /i to 10 /* in thickness, and the same staining may be 
 used as for the development of antherozoids. For archegonia 
 containing young embryos, like that shown in D, Delafield's 
 haematoxylin without any counter-stain gives beautiful prepara- 
 tions when the staining is well done. It is easier for the beginner 
 
 Fig. 43. 
 
 Marchantia polymorpha. X 400. 
 
 A, three early stages in the development of the archegonia. 
 Delafield's haematoxylin. B, young archegonium showing two- 
 neck canal cells and the central cell before the cutting off of 
 the ventral canal cell. Fuchsin and methyl green. C, mature 
 archegonium just ready for fertilization. Safranin-gentian 
 violet-orange. D, young embryo. Delafield's haematoxylin. 
 
Bryophytes 
 
 93 
 
 to get good preparations with the safranin-gentian violet-orange 
 combination. 
 
 The Sporophyte. — Sporophytes in early stages of develop- 
 ment often yield good preparations without very much trouble, 
 but in later stages they are frequently difficult to cut on account 
 of the secondary thickening of the capsule 
 wall and the stubborn extine of the mature 
 spores. It is hard to get Ricciocarpus into 
 paraffin without shrinking, and the same 
 thing may be said of other forms which 
 have such loose tissue with large air cavi- 
 ties. For stages like that shown in fig. 44, 
 as well as for older sporophytes, it will be 
 found more uniformly satisfactory to use 
 celloidin, and cut the sections from 20 /x 
 to 30 /A thick. 
 
 For a study of archegonia, antheridia, 
 young sporophytes, and also for the devel- 
 opment of the thallus, it is better, even in 
 
 Ricciocarpus natans. 
 
 ig sporophyte inclosed in 
 the archegonium. Spore mother- 
 cell stage. All the cells of the 
 sporophyte except a single peri- 
 pheral layer (dotted in the figure) 
 produce spores. Fixed in picro- 
 acetic acid and stained in Dela- 
 field's haematoxylin. Celloidin 
 section 30 microns in thickness. 
 
 forms like Ricciocarpus, 
 to use paraffin. Pro- 
 longed fixing in chro- 
 mo-acetic acid (two to 
 six days), thorough 
 dehydrating, and 
 gradual transfer from 
 alcohol to xylol, and 
 from xylol to paraffin, 
 and also a moderate temperature in the bath (not more than 
 50" C), will often bring the material through in fine condi- 
 tion. 
 
 Fig. 45. Pellia epiphylla. 
 
 A, habit sketch of sporophyte. X 10. B, small portion of 
 sporophyte (at X of ^), showing the capsule wall, the spores, and 
 the elaters. Fixed in chromo-acetic acid and stained in cyanin 
 and erythrosin. Ten microns. 
 
94 
 
 Methods in Plant Histology 
 
 Forms like Pellia cut well in paraffin, especially in younger 
 stages, but even in case of mature sporophytes it is not neces- 
 sary to resort to celloidin. In Pellia and Conocephalus the spores 
 are very large and have a rather thin wall. Both these genera 
 show a peculiar, intrasporal development of the gametophyte, 
 /. e., the gametophyte develops to a considerable extent before 
 it ruptures the spore wall. ( Fig. ^5.) For the older sporophytes 
 of Marchantia it is better not to cut the whole receptacle, but 
 
 Jou(i./|pp,A|it 
 
 Fig. 46. Anthoceros laevis. 
 
 A, longitudinal section of lower portion of sporophyte imbedded in the gametophyte. X 45. B, 
 transverse section of lower portion of sporophyte. X 200. Delafield's hsematoxylin. Ten microns. 
 C, vegetative cell from lower portion of the sporophyte. X 560. Fixed in Flemming's weaker solution 
 and stained in a mixture of acid fuchsin and iodine green. Five microns. D, spore mother-cell showing 
 three of the four chloroplasts with numerous starch grains. The nucleus is in the metaphase of the first 
 division. X 560. Fixed in Flemming's weaker solution, stained in safranin-gentian violet-orange. 
 Five microns. 
 
 rather to remove the branches so that they may be cut separately. 
 For the very best preparations of mature sporophytes it will pay 
 to trim away the gametophyte structures, leaving only enough to 
 show the foot with a few of the surrounding cells. Sections 5 /a 
 to 10 /i thick can be made without much difficulty from material 
 prepared in this way. 
 
 Among the Bryophytes no form affords a better oppor- 
 tunity for studying the development of spores than Anthoceros, 
 
Bryophytes 95 
 
 since a single longitudinal section of the sporophyte may show- 
 all stages, from earliest archesporium to mature spores {^fig. 46) . 
 For studies like A and B, chromo-acetic material cut 10 /u, thick 
 and stained in Delafield's haematoxylin is very good. The 
 starch grains in the chloroplasts take a beautiful violet color 
 with the safranin-gentian violet-orange combination. It is very 
 difficult, however, to bring out the details of nucleus or chloro- 
 plast on account of the minute size of these structures. The 
 drawings from which C and D were reproduced were made with 
 a one-sixteenth oil immersion objective. The drawings, like all 
 the others illustrating the Bryophytes, were reduced one-half by 
 photography. 
 
CHAPTER XIV. 
 
 BRYOPHYTES. 
 
 MUSCI. 
 
 Material for a study of the mosses is much more abundant, 
 and a series of stages in the development of the various organs 
 is easily secured ; but it is much more difficult to obtain good 
 preparations, because so many of the 
 structures are hard to cut. Chromo- 
 acetic acid is to be recommended as 
 the most satisfactory fixing agent, but 
 where structures are refractory and very 
 likely to make trouble in cutting, it will 
 often be found more satisfactory to use 
 picro- acetic acid in the 70 per cent, 
 alcohol, since material fixed in this 
 reagent does not become as hard or as 
 brittle as that fixed in any of the chro- 
 mic-acid series. 
 
 Antheridia. — It is easy to find mate- 
 rial for a study of antheridia, because, 
 
 Fig. 47- 
 
 A, archegonia of Webbera candi- 
 ,1 ,1 •!• 1 1 . cans. X 104. Celloidin section. 
 
 m SO many cases, the anthendial plants Twenty microns. 5, young amheri- 
 can be detected at once without even a 
 
 Polytrichum 
 
 . X420. 
 
 pocket lens. Funaria, with its bunch of antheridia as large as a 
 pin-head, is extremely common everywhere. Spring is the best 
 time to collect it, but it is found fruiting in the autumn and 
 sometimes in summer ; besides, it is easily kept in the green- 
 house, where it may fruit at any time. Bryum proliferiim. has a 
 still larger cluster of antheridia, which may be seen at a distance 
 of several yards. Polytrichum also has a large cluster of anthe- 
 ridia surrounded by reddish leaves, so that the whole is some- 
 times called the moss "flower." In making preparations of 
 Polytrichum these colored leaves should be carefully removed 
 
 97 
 
98 
 
 Methods in Plant Histology 
 
 after the material has been gotten into 70 per cent, alcohol. A 
 single antheridial plant of Polytrichum often furnishes a fairly 
 complete series of stages in the development of antheridia. 
 {Fig. 47.) In all cases the stem should be cut off close up to 
 the antheridia, for many of the moss stems cut like wire. It is 
 not necessary to use celloidin for antheridia, nor is it desirable, 
 
 except where sec- 
 tions from 20 /A to 
 50 /A thick are want- 
 ed for habit work. 
 Delafield's haema- 
 toxylin is recom- 
 mended for staining. 
 Archegonia. — 
 Since the necks of 
 the archegonia are 
 usually long and 
 more or less curved, 
 it is necessary, for 
 habit work, to cut 
 sections as thick as 
 20 /i or 30 ft in or- 
 der to get a view of 
 Jm.flw.A1iQ> an archegonium in a 
 sinsfle section. Cel- 
 
 FiG. 48. Funaria hygrometrica. 
 
 endotheciu 
 )elafield 
 microns. B, C, and D, transverse sections 
 age as .^, taken at three different levels. X 255. Ten microns. 
 
 and amphithe- 
 
 A,3ipex of young sporophyte showing < 
 
 X 420. Chromo-acetic acid and Delafield's haematoxylin. Ten loidin mav be USCd 
 of a sporophyte of the same -^ 
 
 for such prepara 
 tions, but for the development of the archegonium, the oosphere, 
 the canal cells, and also for the process of fertilization, it is better 
 to use paraffin. For the thick celloidin sections the material 
 may be stained in bulk in alum cochineal, but thin paraffin sec- 
 tions should be stained on the slide with more critical stains. 
 
 {^^^- 47-) 
 
 The Sporophyte. — It is often difficult to get good mounts of 
 sporophytes. In the younger stages the calyptras are likely to 
 interfere with cutting, while in the older stages the peristome, or 
 
Bryophytes 
 
 99 
 
 hard wall of the capsule, occasions the trouble. If an attempt is 
 made to remove the calyptra in young stages, like A oi fig. 48, 
 the apex of the sporophyte usually comes with it. While 
 picro-acetic' acid material cuts more easily, chromo-acetic acid 
 followed by Delafield's haematoxylin gives so much sharper dif- 
 ferentiation in stages like those shown in fig. 48 that it is better 
 
 hm^.Hii. 
 
 Fig, 49. Funaria hygrometjica. X 420. 
 
 ^.longitudinal section of capsule. B, transverse section of capsule of about the same age as^. 
 The columella, archesporium, outer spore case, two layers of chlorophyll-bearing cells, and the beginning 
 of the air spaces can be distinguished at this stage. Delafield's haematoxylin and erythrosin. Ten 
 microns. 
 
 to use harder paraffin (55° to 60° C.) and make an effort to get 
 preparations from chromic material. 
 
 Stages like that shown in fig. 4g are cut with comparative 
 ease, for the calyptra is easily removed, and the capsule wall is 
 not yet hard enough to occasion any difficulty. The cell walls 
 are so easily stained in moss capsules that a light counter-stain 
 with erythrosin or acid fuchsin may be used to bring out the 
 
100 
 
 Methods in Platii Histology 
 
 cytoplasm and plastids without appreciably obscuring the cell 
 walls. Funaria and Bryum afford an excellent study in the 
 development of the capsule, since all the structures of a highly 
 differentiated moss sporophyte are present, and Bryum is par- 
 ticularly easy to cut in stages like those 
 shown in fig. §o. 
 
 Sporophytes, in their more mature 
 stages, are almost sure to present con- 
 siderable difficulty in cutting. For gen- 
 eral work fairly good preparations may 
 be gotten from celloidin material, but it 
 is worth while to try paraffin, for it is 
 sometimes successful, and when it does 
 succeed -it is far 
 Jow/j»;y,c superior. As soon 
 
 Fig. so. Bryum. X 200. aS the CcU Walls 
 
 Portion of a nearly mature capsule K^rrin fr> tViirl^^n 
 
 showing operculum, annulus, peris- DCgm tO iniCKCn, 
 tome, and three cells of the sporo- • .1^ rl<^ir/:»1r.r-i 
 
 genous tissue. Fixed in Flemming's ^S m tUC aeVCiOp- 
 weaker solution, stained in safranin j. £ tU 
 
 and Delafield's haematoxylin. Fif- mCUt Ot the pCri- 
 teen microns. . 
 
 stome, safranm is 
 an excellent stain, and this, followed by 
 Delafield's haematoxylin, will give an ele- 
 gant differentiation in the older stages of 
 the sporophyte. After capsules have be- 
 gun to turn brown it will be almost impos- 
 sible to infiltrate them unless they are 
 pricked with a needle. 
 
 The mature sporophytes of Sphagnum 
 '[fig- 51) are exceptionally hard to cut. It -^ -^^'n ry jo^mMtnm 
 
 will be worth while to prick the capsule '^" ^'' ^ ^gnum. 24. 
 
 ^ * Longitudinal section of mature 
 
 with a needle when the material is col- sporophyte, showing also the upper 
 
 lected. This wil 
 
 portion of the pseudopodium and 
 allnw thp fivincr pcrpnt 'he calyptra. Chromo-acetic 
 ailOW me nxmg agent ^^j^^ Delafield's hematoxylin. 
 
 to penetrate readily, and will also facilitate ^""*^"- Ten microns. 
 the infiltration of paraffin or celloidin. The puncture causes 
 only a slight damage, and need not reach the really valuable 
 portion which is to furnish the median longitudinal sections. 
 
Bryophytes lOi 
 
 Protonema and teased mounts of antheridia and archegonia 
 may be made directly in 50 per cent, glycerine without fixing or 
 staining. While this method is often recommended, we have 
 found it better to use 10 per cent, glycerine and allow it to con- 
 centrate. Mounts made in this way retain their green color for 
 a long time. 
 
CHAPTER XV. 
 
 PTERIDOPHYTES. 
 
 This group, including the Filicineae, Equisetineae, and Lyco- 
 podineae, or, more popularly, the ferns, horsetail rushes, and 
 club mosses, is familiar to everyone. Material is abundant, 
 and so easily recognized that anyone who pays a little attention 
 to collecting can, in a single season, get a fine supply for a study 
 of the group. Some desirable forms may not be present in all 
 localities, but these will be few and can be obtained at a reason- 
 able price from those who make a business of collecting. 
 
 FILICINEiE. 
 
 Without attempting to follow any taxonomic sequence, the 
 methods of preparing the various structures of the homosporous 
 forms will be presented, and then the peculiarities of the hetero- 
 sporous members will be considered. 
 
 The Prothallia. — Ripe spores of some fern or other can be 
 obtained at any greenhouse at any time in the year, and spores 
 of most of our native ferns germinate well and produce good 
 prothallia, even if the sowing is not made for several months 
 after the spores have been gathered. 
 
 Fine prothallia of Pteris aquilina have been grown two years 
 after the spores were gathered. Some, however, must be sown 
 at once, or they will not germinate at all. The spores of the 
 common Osmu?ida regalis, and probably of the other members of 
 the genus, must be sown as soon as ripe, or they fail to gefminate. 
 The prothallia of Osmu?ida regalis, if carefully covered with 
 glass, may be kept for a long time. Prothallia of this fern in 
 the writer's laboratory produced ribbon-like outgrowths three- 
 sixteenths of an inch wide, and often more than two inches in 
 length. These prothallia continued to produce archegonia, 
 antheridia, and the ribbon-like outgrowths for more than a year, 
 
 103 
 
104 Methods i?i Plafii Histology 
 
 when they suddenly "damped off." Pteris aquilina and many 
 other ferns often furnish a good supply of antheridia in three 
 weeks after sowing, and the archegonia appear soon after, but it 
 is well to make sowings six weeks before material is needed for 
 use. In Pteris aquilina and in many others, if the spores are 
 sown too thickly, only antheridial plants will be obtained. If 
 prothallia are to produce archegonia, they must have sufficient 
 room and nutrition. If there are no greenhouse facilities and 
 the prothallia must be grown in the laboratory, it is a good plan 
 to take a glass dish, ten or twelve inches in diameter and about 
 two inches deep, put a layer of broken pieces of flower pots on 
 the bottom, cover this with a layer of rich loam, and over this 
 sprinkle a layer of fine, clean sand, since sand is much more 
 easily washed away from the rhizoids than is the loam. The 
 whole should now be thoroughly wet, but not so as to have 
 water standing on the bottom. Sow the spores and cover with 
 a tightly fitting pane of ground glass. There should be no 
 need for moistening the culture again, for prothallia can be kept 
 fresh and vigorous for several months, or even for a year, without 
 any wetting. When it is desired to secure fertilized material, 
 sprinkle the prothallia with water, and the young sporophytes 
 will soon appear. If greenhouse facilities are available, any gar- 
 dener can grow prothallia in abundance without any directions 
 from those who want the material. 
 
 The peculiar tuberous prothallia of Botrychium are seldom 
 found except by the experienced collector. The older prothal- 
 lia, however, may be found by anyone who is able to recognize 
 Botrychium when he sees it. Dig up young plants not more 
 than three or four inches in height, and the prothallia, which 
 persist for years, will often be found still attached. They are 
 easy to cut and may be handled like other prothallia. 
 
 Fern prothallia of the usual type are excellent objects for 
 testing fixing agents, since the prothallia, while still in the fixing 
 agent, may be examined with the microscope, and fluids which 
 cause plasmolysis may be rejected. It will sometimes happen 
 that plasmolysis may be avoided by varying the proportions of 
 
Pteridophytes 
 
 105 
 
 the ingredients of a fixing agent. Chromo-acetic acid with 
 about 0.6 g. chromic acid and 0.4 cc. of acetic acid to 100 cc. of 
 water will seldom cause plasmolysis, and will usually insure good 
 fixing. It is a mistake to suppose that because prothallia are 
 suchdelicateobjects 
 the fixing will take 
 but a few minutes. 
 We should recom- 
 mend at least twen- 
 ty-four hours in 
 chromo-acetic o 
 Flemming's fluid 
 and two or three 
 days will do no harm 
 and may be better. 
 If hot corrosive sub- 
 limate-acetic acid or 
 hot picro-acetic be 
 used, the fixing re- 
 quires only two or 
 three minutes, but 
 results are not as 
 uniformly success- 
 ful as with members 
 of the chromic-acid 
 series. After any 
 
 Fig. 52. Pteris aquihua. 
 
 of the ChromiC-aCld ^, filamentous stage. 5, the apical cell has been established and 
 
 , several segments have been cut off. The figure shovifs the initial 
 
 series, two or three rhizold, and also three rhizoids coming from the main body of the 
 
 ^ . prothallium. C, an older prothallium covered with antheridia in vari- 
 
 hoUrS washing in ous stages of development. From a glycerine mount, fixed in chromo- 
 
 acetic acid and stained in Deiafield's hseraatoxylin. (Miss M. E. 
 
 water will be sufifi- tarrant.) 
 
 cient, if the water be changed as often as it becomes in the 
 
 least degree discolored. 
 
 If preparations are to be mounted whole, as shown in fig. j2, 
 they should be stained as soon as the washing is finished. Any 
 of the following methods gives good results : 
 
 a. Stain in Mayer's hsm-alum six hours or over night, 
 
io6 
 
 Methods in Plant Histology 
 
 wash in water one or two hours, and transfer to lo per cent, 
 glycerine. 
 
 b. Stain in Delafield's haematoxylin thirty minutes, wash in 
 water one or two hours, decolorize in water acidulated with 
 hydrochloric acid (3 drops of HCl to 100 cc. water) one to 
 
 Fig. S3, Pteris cretica, X 250. 
 
 A , early stage in the development of the archegonium. 5, later stage showing the oosijhere, ventral 
 canal cell, and three nuclei in the neck canal. C, still later stage almost ready for fertilization. The 
 ventral and neck canal cells are breaking down, and the oosphere is nearly mature. Cells surrounding the 
 oosphere have become richer in protoplasmic contents, and stain more deeply. D, first division of the 
 embryo. E, young embryo still showing the outlines of the four quadrants. The apical cell in the lower 
 left cjuadrant has cut off the first layer of the root cap. All drawn from material stained in bulk in alum 
 carmine, a method not to be recommended. 
 
 thirty minutes — the time can be determined only by experi- 
 ment — wash in water until the rich purple color of the haema- 
 toxylin replaces the red due to the acid, and then place in lo per 
 cent, glycerine. 
 
 c. Stain \Vith Delafield's haematoxylin as in b, and after the 
 last washing in water stain two to four minutes with an aqueous 
 
Pteridophytes 107 
 
 solution of eosin, wash thoroughly in water, and transfer to the 
 10 per cent, glycerine. Instead of using the acid alcohol after 
 the haematoxylin, the eosin may be allowed to act for several 
 hours, and then i per cent, acetic acid may be used for a few 
 minutes. Wash very thoroughly in water, and, if the stains 
 appear satisfactory, transfer to 10 per cent, glycerine as before. 
 
 d. Use a 2 per cent, solution of iron alum two hours, wash 
 in water five minutes, stain in )^ per cent, haematoxylin two to 
 six hours, wash in water five minutes, and then treat again with 
 iron alum until the stain is sat- 
 isfactory. Wash thoroughly in 
 water and transfer to 10 per 
 cent, glycerine, which, as usual, 
 will concentrate sufficiently for 
 mounting in three or four days. 
 
 For paraffin sections such A ' ^""'^ B 
 
 \ • /• J Fig. S4. Pteris cretica. 
 
 as are shown m Ap-5. 5? and 5</, . , , , .. , . 
 
 J a ~i^ ~r I ^, section of a nearly mature anthendium snowing 
 
 the material should be passed the antherozoids inside X 333. .S, a developing 
 
 r antherozoid showing the blepharoplast drawn out 
 
 tVirnno-h thf» alrnhnlQ allnwincr '°*° ^ deeply staining band. X 1900. Fixed in 
 
 tnrOUgn tne aiCOnOlS, ailOWmg chromo-acetic add and stained in Haidenhain's iron 
 
 about three or four hours for -'"-h— 'oxyiin. 
 
 each grade. The mixtures of xylol and absolute alcohol should 
 take about six hours, and as soon as the pure xylol has been 
 added, the piece of paraffin may be added at the same time. In 
 the bath two or three hours will give good results. It would be 
 worth while to determine the duration of the bath for such objects. 
 Some workers claim that ten or fifteen minutes is amply suffi- 
 cient, and that there is less danger from shrinking, while others 
 think that several hours is better, and that two or three days will 
 do no damage, if the fixing has been thorough and the tempera- 
 ture is not allowed to become higher than 50° C. If bergamot 
 oil be used instead of xylol, there is less danger of collapse pre- 
 vious to the bath, but the bath itself must usually be more pro- 
 longed, since the bergamot oil is not so easily gotten rid of as is 
 the xylol. 
 
 For morphological purposes sections 15 /i to 20 /^ thick are 
 better than thinner ones. Delafield's haematoxylin, with or without 
 
io8 
 
 Methods in Plant Histology 
 
 a counter-stain with erythrosin, is good for such sections as 
 are represented m fig. §j. If very thin sections are wanted for 
 cytological study, it is better to use the safranin-gentian violet- 
 orange combination. For such views of antheridia as are shown 
 in fig. 5^, Haidenhain's iron alum-haernatoxylin seems to bring 
 out the blepharoplast (centrosome) most sharply. 
 
 The Sporophyte. — Methods for young sporophytes like those 
 shown in fig. §j, D and E, are the same as for archegonia and 
 antheridia. 
 
 Roottips to show the prominent apical cell are easily imbedded 
 in parafifin. They should be cut i 5 ft to 20 /a thick. Delafield's 
 
 haematoxylin, without any con- 
 trast stain, is best for bringing 
 out the prominent apical cell and 
 its segments. 
 
 Sporangia should also be cut 
 m parafifin. Pteris is a good form 
 for sporangia, since the long mar- 
 gmal sorus makes it possible to 
 get an immense number of median 
 longitudinal sections of sporangia 
 in a single preparation. {^Fig. jj.) 
 Pteris ere tic a can always be 
 found in fruit in greenhouses. 
 Select a series of stages. The leaf should be cut with a razor — 
 not with scissors — into pieces about one-fourth of an inch long. 
 If sections of the whole leaf are not wanted, only the marginal 
 sorus need be cut off; in this way a much greater number of 
 sporangia may be gotten upon a slide. Aspidium and Cyrtomium 
 give beautiful views of the indusium covering the cluster of 
 sporangia. The most satisfactory preparations will be gotten 
 from material in which the sporangia have not yet begun to 
 turn brown. 
 
 Botrychium furnishes excellent and usually accessible material 
 for studying the development of sporangia of the eusporangiate 
 type. For the archesporium and early stages in the development 
 
 Fig. 55. 
 A , young ; 
 
 Pteris cretica X 560 
 
 gc in the development of the spo- 
 rangium. B, older stage showing the tapetum. 
 Fixed in chromo-acetic acid and stained in 
 Delafield's haematoxylin and erythrosin. 
 
Pteridophytes 1 09 
 
 of the sporangia the material should be collected in the latter 
 part of August or in September. When the sporangia for one 
 year are ready to shed their spores, the sporangia for the next 
 year will be found in early stages of development. When a 
 frond is found, dig the plant up very carefully and remove the 
 large frond. At its base will be found the frond for the next 
 season, the sterile portion bent over so that its tip is directed 
 downward and the fertile portion — one-fourth to three-fourths 
 of an inch in length — projecting from the ventral surface of the 
 sterile frond and directed upward. It is worth while to preserve 
 the whole bud and also portions of the stipe, rhizome, and roots. 
 These vegetative parts should be cut, with a razor or very sharp 
 knife, into pieces about one-fourth of an inch in length. The 
 larger roots are better for transverse sections. The root tips are 
 unusually favorable for preparations of the apical cell. The 
 upper part of the rhizome cuts easily, but all parts of this plant, 
 even the older portions of the rhizome, can be cut in paraffin. 
 Delafield's haematoxylin or iron alum-hsematoxylin gives fine 
 preparations of the young sporangia. The older sporangia, from 
 the mother-cell stage up to the shedding of the spores, stain 
 better in cyanin and erythrosin or in the safranin-gentian violet- 
 orange combination. The following schedule for paraffin sections 
 will give elegant mounts of the root, stipe, and rhizome : 
 
 1 . Stain in safranin, about 24 hours. 
 
 2. Fifty per cent, alcohol until little or no safranin is left in the cellulose 
 walls. Use a very weak acid alcohol, if necessary. 
 
 3. Delafield's haematoxylin, 5 to 10 minutes. 
 
 4. Water, 15 minutes. 
 
 5. Thirty-five and 50 per cent, alcohol, 10 seconds each. 
 
 6. Acid alcohol — the same as was used for the safranin — a few seconds. 
 
 7. Seventy per cent, alcohol until the purple color returns. The lignified 
 walls should now show a brilliant red and the cellulose walls a rich purple. 
 If either stain is too deep or too faint, do not proceed any farther before 
 making the necessary correction. If the safranin washes out, and the 
 haematoxylin is too intense, do not use any acid alcohol in the second 
 step and shorten the period in the haematoxylin. 
 
 8. Eighty-five and 95 per cent, alcohol, a few seconds each. 
 
 9. Hundred per cent, alcohol, 30 seconds to i minute. 
 
 10. Xylol until cleared. 
 
 11. Balsam. 
 
no 
 
 Methods m Plant Histology 
 
 The spore mother-cells of Osmunda are excellent for a study 
 of mitosis. The young sporangia of 0. cinnamomea and 0. Clayto- 
 niana show the mother-cell stage in the autumn, but the division 
 into spores does not occur until the following spring, in the 
 vicinity of Chicago the mitotic figures being found during the 
 
 latter part of April. 0. 
 P 9e^^Xi:iW^!/^^^^^i^'''i^i':'^^isJ^~ 'j. regalis does not reach the 
 
 mother-cell stage in the 
 autumn. Material for 
 mitosis should be col- 
 lected during the first 
 two weeks in May. The 
 material may be fixed in 
 the medium chromo- 
 acetic solution or in Flem- 
 ming's weaker solution. 
 Sections should not be 
 thicker than lO /l*, and 
 5 /* will be found more 
 satisfactory. 
 
 Preparations of the 
 woody structure of the 
 sporophyte are easily 
 made. The rhizome of 
 Pteris aquili?ia affords as 
 good material as any 
 
 Fig. 56. P.eris aquilina. {fig-S^^)' I" digging Up 
 
 A part of a transverse section of the vascular bundle of the rhizOmCS, do not merely 
 rhizome. X i66. ^, endodermis. /, pericycle. j/. sieve tube, _ ^ 
 
 /, scalariform tracheid. Drawn from a celloidin section from difif doWU Until thc rhi- 
 material fixed in picro-acetic acid and stained in safranin and ° 
 
 Deiafieid's hsematoxyiin. zomc Can be graspcd and 
 
 then pull it up, for such material is sure to show the pericycle of the 
 bundles torn away from the parenchyma. Dig carefully around 
 the rhizome and then cut off with a very sharp knife pieces about 
 two inches in length. Put the fresh rhizome into the hand micro- 
 tome and cut as thin sections as possible. Keep the knife wet 
 with 95 per cent, alcohol and put the sections into 95 per cent. 
 
Pteridophytes 
 
 alcohol as fast as they are cut. Fifteen or twenty minutes is 
 
 sufficient for fixing. Pass down through the grades of alcohol, 
 
 about two or three minutes in each grade. Stain in safranin 
 
 twenty-four hours, wash in water ten minutes, and then stain in 
 
 Delafield's haematoxylin ten to twenty minutes. Pass through 
 
 the alcohols, about one minute in each grade, at 70 per cent, 
 
 alcohol transferring to acid alcohol from one to five seconds ; 
 
 then pass on, clear in 
 
 xylol, and mount in 
 
 balsam. If the stain 
 
 is successful, the 
 
 xylem should show a 
 
 brilliant red, and the 
 
 cellulose walls a rich 
 
 purple. 
 
 Another method is 
 to stain for about twen- 
 ty minutes, or several 
 
 , .... Fig. 57. Marsilea quadrifolia. X 333. 
 
 hours, m lodme green ^ , • ^ i. • . • • i. • u 
 
 ' o A, apex of megaspore with archegonium containing the oosphere. 
 
 r>r mf»fVi-\7l crrp>f»n rincf Large starch grains are shown beneath the archegonium. ^,young 
 
 ui iiiCLH_yi giccii, iiiisc gn^bryo. Both fixed in picro- acetic acid and stained in Delafield's 
 
 in 70 per cent, alcohol ^hematoxylin. (M.ss M. E. Tarrant.) 
 
 until the green is prominent only in the xylem, then stain for 
 from one to three minutes in acid fuchsin (i per cent, solution 
 in 70 per cent, alcohol), dehydrate rapidly, clear in xylol, and 
 mount in balsam. The xylem should have a sharp bright green 
 color, and the cellulose a bright red. 
 
 The most beautiful preparations may be obtained by imbed- 
 ding in celloidin and staining in safranin and Delafield's haema- 
 toxylin. These stains allow a use of acid which extracts all color 
 from the celloidin and still leaves a sufficient amount in the 
 tissues. The large apical cells of rhizomes are easily cut in 
 either paraffin or celloidin. 
 
 The megaspores and microspores of Marsilea are easily 
 obtained {jigs. 57 and 58^. Cut away a portion of the hard 
 sporocarp and place the sporocarp in a dish of water. The 
 gelatinous ring with its sori will sometimes come out in a few 
 
 Fig. 57. Marsilea quadrifolia. 
 
12 
 
 Methods i?i Plant Histology 
 
 minutes. In less than twenty-four hours the microspores, start- 
 ing from the one-cell stage, will produce the mature antherozoids. 
 The development of the megaspore is equally rapid. Embryos 
 are abundant in two or three days. For morphological work, 
 picro-acetic acid, used hot, is very good, since the material does 
 not occasion so much difficulty in cutting. Chromo-acetic 
 material allows better staining, but the cutting is more uncertain. 
 
 It is best to prick the 
 megaspores with a 
 needle while they are 
 in the fixing fluid, in 
 order to facilitate the 
 infiltration of paraffin. 
 Better mounts of the 
 microspores can be 
 obtained if the trou- 
 blesome megaspores 
 be picked out from 
 the sorus while the 
 material is still in the fixing agent or the alcohols. The mega- 
 spores must be imbedded in rather hard paraffin, and one must 
 expect to hone the knife thoroughly before it can be used again, 
 for when the knife strikes a megaspore of one of the heterospor- 
 ous pteridophytes, it seems like striking a grain of sand. 
 
 The following schedule will usually give good sections of the 
 hard megaspores of the heterosporous pteridophytes, whether 
 they are to be cut separately or in their sori or strobili. The 
 essential features of the method are suggested by Miss F. M. 
 Lyon's work on Selaginella: 
 
 I. Chromo-acetic acid, medium solution, 2 to 6 days. 
 
 2.. Wash- in water, i day. 
 
 3. Thirty-five to 95 per cent, alcohol, i day each. 
 
 4. One hundred per cent, alcohol, 3 or 4 days, changing several times. 
 
 5. Mixtures of alcohol and xylol, 2 days. 
 •6, Pure xylol at 53° C, i or 2 days. 
 
 7. Add parafifin to the xylol and keep at 53° C. for 2 or 3 days. 
 
 Fig. 58. Marsilea quadrifolia. X 560. 
 
 A , microspore before germination. B, microspore with antheridia 
 nearly mature. Fixed in chromo-acetic acid and stained in safranin- 
 gentian violet-orange. 
 
Pteridophytes 113 
 
 8. Pure parafifin 53° C, 3 or 4 days, and then in harder paraffin at 60° C. to 
 70° C, 2 or 3 days. 
 
 9. Imbed in rather thin cakes. 
 
 While the method is tedious, it is worth the trouble, for even 
 old strobili of Selagiiiella yield smooth ribbons at 5/x. 
 
 ^s fig. ^y suggests, the mature archegonia, and especially the 
 young embryos, may be removed from the top of the megaspore 
 and cut with perfect ease. 
 
 The spermatozoid, which in Marsilea has an unusually large 
 number of turns in the spiral, may be mounted by methods 
 already described. 
 
CHAPTER XVI. 
 
 PTERIDOPHYTES. 
 EQUISITINEJE. 
 
 The prothalHa of Equisetum are easily grown by the method 
 already described for the Filicineae, but the spores must be 
 sown as soon as ripe, because they fail to germinate if kept more 
 than a few days. In the vicinity of Chicago the spores of 
 Equisetum arvense are shed during the latter part of April. The 
 methods for preparing the prothallia are the same as for the 
 Filicineae. The sporangia are harder to cut, but good prepara- 
 tions should be secured from paraffin material. E. arvense is 
 abundant everywhere, and is to be preferred on account of the . 
 comparative ease with which the sporangia and other portions of 
 the fertile shoot can be cut. Longitudinal sections of the 
 younger strobili show various stages in the development of the 
 spores, the more advanced stages being found at the base of the 
 strobilus. Tetrads may be found at the base of the strobilus, 
 while the spore mother-cells at the apex are still undivided. Of 
 course, it is impossible to stain a longitudinal section of such a 
 strobilus so that all stages will be satisfactory. For the beginner, 
 at least, this is not a serious objection, for he will be almost sure 
 to secure some stage beautifully stained. The experienced 
 worker, who is able to control his staining with more precision, 
 will prefer transverse sections. Material for sporangia should 
 be obtained as soon as the fertile shoot appears above ground, 
 and if it can be obtained earlier, so much the better. When the 
 spores are shed, the young sporangia which are to develop the next 
 year can already be detected. The safranin-gentian violet-orange 
 combination can be recommended for the development of the 
 mother-cell and the formation of tetrads, but Delafield's haema- 
 toxylin or Haidenhain's iron alum-haematoxylin will be more 
 satisfactory for earlier stages. 
 
 "5 
 
Ii6 Methods in Plant Histology 
 
 The roots are very small, but have large cells and easily yield 
 good preparations. In case of such small objects it is a good 
 plan to add a few drops of eosin to the alcohol during the pro- 
 cess of dehydrating, in order that the material may be seen more 
 easily. The slight staining does no damage, even if more 
 critical stains are to be used after the sections are cut. It is 
 easy to get longitudinal sections of the roots by cutting trans- 
 verse sections of the nodes. In E. arvense these roots at the nodes 
 are quite numerous. Sections of the stem of the fertile shoot of 
 E. arvense are easily cut in paraffin or celloidin, but sections of 
 the stem of E. hiemale or similar species do not cut in paraffin, 
 and results are rather uncertain even in celloidin. The growing 
 points of stems, however, may be cut with ease in paraffin. E. 
 arveme is particularly favorable on account of the numerous 
 apical cells which may be found in a single preparation. Dela- 
 field's haematoxylin, used alone, is good for the apical cells, but 
 for sections of older stems a slight counter-stain with erj'throsin 
 will improve the mount. 
 
CHAPTER XVII. 
 
 PTERIDOPHYTES. 
 LYCOPODINEiE. 
 
 It is not very difficult to get paraffin sections of young 
 sporangia of Selaginella, but the method just recommended for 
 Marsilea should be resorted to for the older strobili {fig. 5p). 
 
 The older megaspores had better be pricked with a needle 
 and cut one at a time. A slight puncture at the basal portion of 
 the megaspore 
 does no damage 
 and insures a thor- 
 ough infiltration. 
 If the megaspores 
 are imbedded sep- 
 arately, they will 
 usually orient 
 themselves so that 
 sections perpen- 
 dicular to the par- 
 affin cake will 
 show the most in- 
 structive views of 
 the gametophyte 
 structures. 
 
 Fig. 59. Selaginella Mertensii. X 93. 
 
 Ay microsporangium containing microspoi 
 ing the beginning of the prothallium. FixeU ni p 
 Delafield's hae- stained in Delafield's hsematoxylin. (Miss M. E. Tarrant.) 
 
 a megaspore show- 
 icro-acetic acid and 
 
 matoxylin, used alone, is a good stain. The safranin-gentian 
 violet-orange combination gives a brilliant differentiation in 
 stages likey?^. jp, B. The microspores of Isoetes offer the same 
 difficulties. Cyanin and erythrosin is a fine combination for the 
 reserve food-stuffs in Isoetes macrospores. 
 
 For young sporangia of Isoetes, the leaves should be cut off 
 about one-eighth of an inch above the sporangia, and the stem 
 
 117 
 
Ii8 Methods in Plaiit Histology 
 
 should be cut off below, leaving just enough to hold the 
 sporangia together. Cut longitudinal sections and stain in 
 Delafield's haematoxylin, with or without the addition of a light 
 touch of erythrosin. The older sporangia had better be removed 
 and cut separately. Transverse sections of the stem are very 
 interesting. Young stems and the apices of old ones cut well 
 in paraflfin, but older stems will give more satisfactory results in 
 celloidin. 
 
 It is much easier to get good preparations of the sporangia 
 of Lycopodium, since there are no megaspores with their hard 
 walls. Delafield's haematoxylin, without any contrast stain, will 
 bring out the developing sporangia, which are usually to be 
 found even after the sporangia in the lower part of the strobilus 
 are beginning to shed their spores. The directions for the 
 rhizome of the Filicineae will also serve for the stem of 
 Lycopodium. 
 
CHAPTER XVIII. 
 
 SPERMATOPHYTES. 
 
 In this immense group we cannot hope to give even approxi- 
 mately complete directions for making preparations, but must 
 be content to give a few hints which may prove helpful in col- 
 lecting material and in securing mounts of the more important 
 structures of the flowering plants. We shall consider the gym- 
 nosperms and the angiosperms separately, although . in many 
 respects the technique is the same for both. 
 
 GYMNOSPERMS. 
 
 Since Pinus is a characteristic type, we shall describe methods 
 for demonstrating various phases in the life-history of this genus, 
 hoping that the directions will enable the student to experi- 
 ment intelligently with other forms. 
 
 Spermatogenesis. — In October the clusters of staminate cones 
 which are to shed their pollen in the coming spring are already 
 quite conspicuous. The cones should be picked off separately, 
 and the scales should be carefully removed so as to expose the 
 delicate greenish cone within. At this time the archesporial 
 cells are easily distinguished. Material collected in January, or 
 at any time before growth is resumed in the spring, shows about 
 the same stage of development. If it is desired to secure a 
 series of stages with the least possible delay, a branch bearing 
 numerous clusters of cones may be brought into the laboratory 
 and placed in a jar of water. Growth is more satisfactory in 
 case of branches broken off in the winter than in those brought 
 in before there has been any period of rest. The material can 
 be examined from time to time, and a complete series is easily 
 secured. The karyokinetic figures in the pollen mother-cells 
 furnish exceptionally instructive preparations. Staminate cones 
 which will yield karyokinetic figures can be selected with con- 
 siderable certainty by examining the fresh material. Crush a 
 
 119 
 
120 Methods in Plant Histology 
 
 microsporangium from the top of the cone and one from the 
 bottom, add a small drop of water and a cover to each, and exam- 
 ine. If there are pollen tetrads at the bottom, but only undi- 
 vided spore mother-cells at the top, it is very probable that 
 longitudinal sections of the cone will yield the figures. If a drop 
 of methyl green be allowed to run under the cover, it will enable 
 one to see whether figures are present or not. When desirable 
 cones are found, they should be cut longitudinally into two 
 halves. The later stages, showing the germination of the micro- 
 spores, furnish better sections if the cones are cut transversely 
 into small pieces about three-sixteenths of an inch thick. It is 
 very easy to get excellent mounts of the pollen just at the time of 
 shedding. Shake a large number of cones over a piece of paper, 
 thus securing an abundance of material. Fix in chromo-acetic 
 acid, wash in water (a few minutes is sufficient, and the water need 
 not be changed), pass through the alcohols, allowing each to 
 act for about two hours, make the usual gradual transition from 
 alcohol to xylol, and from xylol to paraffin. It is best that the 
 material should be in a small bottle not more than one-fourth 
 of an inch in diameter ; at any rate, the pollen should be in such 
 a bottle during infiltration, which should not require more than 
 two or three hours, although a longer period does no harm if the 
 temperature does not rise above 52° or 53° C. Now put the lower 
 portion of the bottle into cold water, and thus harden the paraffin 
 as quickly as possible. Break the bottle carefully, cut off the lower 
 portion of the paraffin containing the pollen, mount it on a block 
 in the usual manner, and trim away some of the paraffin so that 
 two parallel surfaces will make the sections ribbon well. Material 
 in this stage shows a large tube nucleus, a somewhat lenticular 
 cell with a more deeply staining nucleus, and, lastly, two small 
 prothallial cells quite close to the spore wall. The prothallial 
 cells cannot always be detected at this stage, and there may be 
 some doubt as to whether two such cells are always present. 
 The division of the lenticular cell into "stalk cell" and "body 
 cell," and also the division of the body cell into the two male 
 cells, must be looked for in sections of the nucellus of the ovule. 
 
Spermatophytes 
 
 121 
 
 Oogenesis. — The entire ovulate cone at the time of pollina- 
 tion is easily cut in paraffin. Longitudinal sections of the cone 
 at this time give good views of the bract and ovuliferous scale 
 bearing the ovules. The integument is very well marked, and 
 in the nucellus one or more sporogenous cells can usually be 
 distinguished. As soon as the scales close up after pollination, 
 the cone will be too hard to cut, and it will be necessary to 
 remove the scales and cut them separately. For a study of the 
 ovule and the structures within it, better preparations will be 
 
 ^.<^- 
 
 Jovcc f\pp. Nt*t. 
 
 Fig. 6o. Pinus Laricio. X 104. 
 
 A, top of prothallium with an archegonium just before the cutting off of the ventral canal cell. 
 Fixed in Flemming's weaker solution and stained with Haidenhain's iron alum-haematoxylin. Collected 
 June 18, 1897. B, C, and D, early stages in the formation of the embryo. Fixed in chromo-acetic acid, 
 and stained in safranin -gentian violet-orange. Collected July 2, 1897. 
 
 obtained by carefully cutting off the pair of ovules from the 
 scale. For preparations like that represented in fig. 60, A, it is 
 a good plan to remove the endosperm with its archegonia from 
 the ovule. Fixing, infiltration, and cutting will then occasion 
 but little trouble, and the whole ribbon may be gotten upon a 
 single slide. However, at this stage the pollen tubes with their 
 contents are rapidly working their way through the nucellus 
 toward the archegonia, and consequently it is better to retain 
 
122 Methods i?i Plant Histology 
 
 enough of the tissues of the ovule to keep the nucellus in place. 
 In later stages, after fertilization has taken place, it is necessary 
 to remove the endosperm. In stages like fig. 60, B, C, D, and 
 later, the developing testa should be dissected away with great 
 care, for a very slight pressure is sufficient to injure the delicate 
 parts within. Mature embryos may be dissected out from the 
 endosperm before fixing, but it is hardly necessary, since they 
 cut quite well if left in place. The "pine nuts" or "pinon," to 
 be found upon the market, are good for a study of the mature 
 embryo. The testa, which is quite a hard shell, should be taken 
 off, and the endosperm should be allowed to soak in water for 
 about twenty-four hours, after which the embryo may be dis- 
 sected out and fixed. 
 
 The period at which the various stages may be found varies 
 with the species, the locality, and the season. In Pinus Laricio 
 (the common Austrian pine) at Chicago, in the season of 1897, 
 material collected May 27 did not yet show archegonia ; the 
 ventral canal cell was cut off about June 21 (see fig. 61), the 
 fusion of the pronuclei occurred about a week later, and stages 
 like fig. 60, B, C, and D, were common in material collected July 
 2. In the season of 1896 all the stages appeared about two 
 weeks earlier. In Pinus sylvestris the stages appeared a little 
 earlier than in Pinus Laricio. After the stage shown in fig. 60, A, 
 has appeared, it is necessary to collect at intervals of not more 
 than two days until the stage shown mfig. 60, D, is reached. If 
 collections are made at intervals of four or five days, the most 
 interesting stages, like the cutting off of the ventral canal cell, 
 fertilization, and the first divisions of the nucleus of the oospore, 
 may be missed altogether. It should be mentioned that all the 
 ovules of a cone will be in very nearly the same stage of devel- 
 opment. 
 
 A rather strong chromo-acetic acid (i g. chromic acid and 
 1/2 cc. glacial acetic acid to 100 cc. water) can be recom- 
 mended for the entire scries in spermatogenesis, oogenesis, fer- 
 tilization, and formation of the embryo. After repeated trials 
 the popular Flemming's solution does not seem to be at all 
 
Spennatophytes 
 
 123 
 
 superior and often fails to give as good results as the cheaper 
 fixing agent. 
 
 The following stains may be suggested : for studying the 
 pollen tubes in the nucellus, cyanin and erythrosin ; for the 
 development of the archegonium up to the stage shown in 
 fig. 60, A, 
 Delafield's 
 haematoxy- 
 lin; for the 
 stages shown 
 in fig. 60, A, 
 iron alum- 
 haematoxylin 
 or the safra- 
 nin - gentian 
 violet-orange 
 combination; 
 for the stage 
 shown in fig. 
 61, nothing 
 seems to 
 equal the 
 safranin-gen- 
 tian violet- 
 orange com- 
 bination ; for 
 stages like 
 fig. 60, B, C, 
 and D, and 
 also for later 
 stages in the 
 development 
 of the em- 
 bryo, Delafield's haematoxylin brings out the walls perfectly, but 
 since mitotic figures are very frequent in these stages, it is worth 
 while to use the safranin combination with some preparations, 
 
 
 1 
 
 • #« ^- 
 
 
 ^ ^- • 5 « 
 
 
 # "^ 
 
 
 # 
 
 1 
 
 ^ m ^ 
 
 '••S 
 
 f 
 
 
 
 
 
 
 ^ ■ .♦ *^' f ; 
 
 A 
 
 ^ -ir ■ v>,.i»_l^alMi 
 
 d 
 
 Fig. 61. Pinus Laricio. X 710. 
 
 The mitotic figure concerned in cutting off the ventral canal cell. The 
 nucleus at the lower end of the spindle is the nucleus of the oosphore. 
 Fixed in chromo-acetic acid, and stained in the safranin-gentian violet- 
 orange combination. Collected June 21, 1897. 
 
124 Methods in Plattt Histology 
 
 although it is much inferior to Delafield's hsematoxylin when 
 cellulose walls are to be emphasized. 
 
 All the stages which have been described can be cut in 
 paraffin with little difficulty. 
 
 The Leaves. — The leaves of our common gymnosperms cut 
 readily in paraffin while they are young and tender, but as they 
 approach maturity it is a fruitless task to attempt paraffin sec- 
 tions. Celloidin sections are far more satisfactory. Cut the 
 needles into pieces about one-fourth of an inch long, fix in a 
 picro- corrosive -acetic mixture {% g. picric acid, 2 g. cor- 
 rosive sublimate, i cc. glacial acetic acid, lOO cc. 50 per 
 cent, alcohol). If used hot, five minutes is sufficient, but if 
 used cold it should be allowed to act for two or three hours. 
 After the material has been imbedded in celloidin, the block 
 should be placed in equal parts of 95 per cent, alcohol and 
 glycerine for a few days, after which it should cut quite readily. 
 Stain with safranin and Delafield's haematoxylin, clear in Eycle- 
 shymer's clearing mixture, and mount in balsam. 
 
 Fairly good sections may be obtained in great quantities with 
 little trouble by the following method : Make a bunch of the 
 needles as large as one's little finger, wrap them firmly together 
 with a string, allowing about an eighth of an inch of the bunch 
 to project above the wrapping; then fasten the whole in a hand 
 microtome, and every stroke of the razor will give twenty or 
 thirty sections, some of which will surely be good. As the sec- 
 tions are cut, they maybe put directly into 95 per cent, alcohol, 
 and after a few minutes can be transferred to 50 per cent, alco- 
 hol and then to the stain. Dehydrate, clear in xylol, and mount 
 in balsam. 
 
 Stems and Roots. — With a sharp razor fairly good sections 
 of stems and roots may be made without imbedding, especially 
 if the hand microtome be used. Young buds may be cut in 
 paraffin. Stems and roots as large as half an inch in diameter 
 can be cut in celloidin. The material should be cut into pieces 
 not more than one-fourth of an inch long. The following treat- 
 ment should give good results : 
 
Spermatophytes 125 
 
 1. Picro-corrosive-acetic mixture, five minutes if used hot, or 
 wo to three hours if used cold. 
 
 2. Wash in 50 per cent, alcohol, to which a little iodine has 
 been added, two hours; 70 per cent., 85 per cent., 95 per cent., 
 
 our hours each ; absolute alcohol, ten hours ; then change to 
 fresh absolute alcohol, which should act for ten hours longer. 
 
 3. Ether alcohol, twenty-four hours. Some prefer to pre- 
 cede the ether alcohol by a mixture of equal parts of ether 
 alcohol and absolute alcohol. 
 
 4. Thin celloidin (about 2 per cent.), two or three days; 
 6 per cent, celloidin, two or three days; 10 per cent, celloidin, 
 two or three days. After the thin celloidin has acted for a few 
 days, the cork may be removed for a short time each day, thus 
 allowing the thin celloidin to become thick by the evaporation 
 of the ether alcohol. 
 
 5. Get some small blocks of wood (three-eighths inch cubes 
 of white pine are good), wet one of them in ether alcohol, dip it 
 into thin celloidin, place the object upon the block in convenient 
 position for cutting, pour over it a few drops of 10 per cent, cel- 
 loidin, and then plunge the whole into chloroform. Leave it in 
 the chloroform about twenty-four hours, and then transfer to a 
 mixture of isqual parts of 95 per cent, alcohol and glycerine, 
 where it should remain for several days. Material may be kept 
 here indefinitely. Even refractory stems may be cut after they 
 have been in this mixture for a couple of weeks. 
 
 6. Cut the sections, keeping the knife wet with the alcohol 
 and glycerine mixture. Transfer the sections to 70 per cent, 
 alcohol, then to 50 per cent., and then to the stain. 
 
 7. Stain in safranin, twenty-four hours; wash in 35 per cent, 
 alcohol, about a minute ; stain in Delafield's haematoxylin, five to 
 ten minutes; wash in water, two minutes ; 35 per cent, alcohol, two 
 to five minutes ; 50 per cent, alcohol, two to five minutes ; acid 
 alcohol, one to ten seconds; 70 per cent., 85 per cent., 95 per 
 cent., about two minutes each ; Eycleshymer's clearing mixture 
 until cleared, usually about one or two minutes. Mount in 
 balsam. 
 
:26 
 
 Methgds m Plant Histology 
 
 Xylem should show a brilliant red color and cellulose a 
 rich purple, if the stain is successful. If either stain is too weak 
 or too prominent, the duration of the stain, the length of time 
 in the alcohols, or the time in acid alcohol must be varied until 
 the desired result is secured. 
 
 People who make all their anatomical sections without imbed- 
 ding may regard this method as tedious and unnecessary, but 
 
 such preparations 
 
 TWUwdtr^itM °{ Chicag o. 
 
 i<x«9.mil,li>n3.ti,y. 
 <a. U/ THau^u. nio. 
 
 'M!mmmMmM>>Mmmm9* 
 
 
 
 
 /oixr. ;^pp. A\i5.. 
 
 
 
 ' 
 
 \ 
 
 lea ^aii-iM^ 
 
 
 i 1 
 
 i 
 
 g 
 
 s 
 
 
 
 
 ^^ 
 
 
 ',U/-A^A\.: 
 
 
 will show much 
 which is never seen 
 in mere free-hand 
 sections, for the 
 reason that free- 
 hand sections, if 
 thin enough to 
 show any detail, 
 will lose most of 
 their cell contents, 
 while in celloidin 
 sections every- 
 thing is held in 
 place. Even if the 
 celloidin sections 
 be passed through absolute alcohol and cleared with clove oil, a 
 process which dissolves away the celloidin, the contents of the 
 cells will still be retained in most cases, and stains which 
 cannot be extracted from celloidin may be used. However, 
 we prefer to use the safranin and Delafield's haematoxylin and 
 retain the celloidin, since this combination can be extracted 
 completely from the celloidin and still leave the object brilliantly 
 stained. 
 
 For preparations of the mature wood a piece of white pine 
 from a dry-goods box furnishes perfect material. It should not 
 be imbedded, but the cutting w^ll be facilitated if the piece be 
 soaked for a few hours in water or in the alcohol and glycerine 
 mixture. In all preparations designed to show the structure of 
 
 Fig, 62. 
 Slides showing labels and methods of arranging sections. 
 
Spermatophytes 127 
 
 wood there should be three sections — a transverse, a longitu- 
 dinal radial, and a longitudinal tangential. These may be 
 arranged as in the upper slide oi fig. 62, but if it is desired to 
 make a thorough study of the structure, it is a good plan to have 
 on each slide several sections of each kind, thus having an 
 opportunity to use a variety of stains. Fuchsin and iodine green 
 is a good combination. The safranin and Delafield's hasma- 
 toxylin is also excellent. 
 
 For a study of young stems or roots, preparations like that 
 shown in the lower slide oi fig. 62 will be found very convenient. 
 
CHAPTER XIX. 
 
 SPERMATOPHYTES. 
 ANGIOSPERMS. 
 
 Success depends largely upon judgment and care in selecting 
 and trimming material before it is put into the fixing agent in 
 the field. While the following directions cannot be applied to 
 all plants, they should, nevertheless, enable the student to make 
 such modifications as may be demanded by any particular form 
 
 Floral Development. — For a study of floral development very 
 young buds are necessary, and it is best to select those forms 
 which have rather dense clusters of flowers, in order that a com- 
 plete series may be obtained with as little trouble as possible. 
 
 The usual order of appearance of floral parts is (i) calyx, 
 (2) corolla, (3) stamens, and (4} carpels, but if any of these 
 organs are reduced or metamorphosed, their order of appearance 
 may be affected. 
 
 Floral development is easily studied in the common Capsella 
 bursa-pastoris. The best time to collect material is late in March 
 or early in April. Dig up the plant, carefully remove the leaves, 
 and in the center of the rosette a tiny white axis will be found. 
 A series of these axes from one-eighth of an inch to three- 
 eighths of an inch in length and from one-sixteenth of an inch 
 to three-sixteenths of an inch in diameter will give a very com- 
 plete series of stages in the development of the floral organs. 
 Preparations from the apex of the shoot taken after the inflores- 
 cence appears above ground are not to be compared with these 
 taken early in the season. Fix in chromo-acetic acid and stain 
 in Delafields haematoxylin. The sections should be longitudinal 
 and about 5 /x thick. 
 
 The common dandelion, Taraxacum officinale, affords an excel- 
 lent series with little labor. Examine vigorous plants which have, 
 as yet, no flowers or buds in sight. Dig up the plant and dissect 
 away the leaves. If there is a white cluster of flower buds, the 
 ' 129 
 
130 Methods in Plant. Histology 
 
 largest not more than three-sixteenths of an inch in diameter, 
 cut out the cluster, leaving only enough tissue at the base to 
 hold the buds in place. Larger heads should be cut separately. 
 Fix and stain as in Capsella. 
 
 Our most common thistle, Cnicus lanceolatus, shows the floral 
 development with unusual clearness, but the preparation of the 
 material is somewhat tedious. The involucre, which is too hard 
 to cut, must be carefully dissected away. Retain only enough of 
 the receptacle to hold the developing florets in place. A series 
 of sizes with discs varying from one-eighth of an inch to three- 
 eighths of an inch in diameter will show the development from 
 the undifferentiated papilla up to the appearance of the arche- 
 sporial cell in the nucellus of the ovule. The Canada thistle, 
 C?iicus arvensis, is equally good, but it is more diflficult to dissect 
 out the desirable parts. 
 
 In the willows, Salix, the bud scales must be removed and 
 the copious hairs should be trimmed off as much as possible 
 with scissors, after which the catkin should be cut in two longi- 
 tudinally and placed in the fixing agent. 
 
 Spermatogenesis. — The earlier stages in spermatogenesis will 
 be found in the preparations of floral development. For tracing 
 the nuclear changes involved in this process the lilies furnish 
 very good material, because the cells and nuclei are exceptionally 
 large. Several species of Lilium are common in greenhouses, 
 and these may be used where wild material is not available. In 
 early stages where the sporogenous cells have not yet begun to 
 round off into spore mother-cells it is sufificient to remove the 
 perianth, retaining just enough of the receptacle to hold the sta- 
 mens in place. Transverse sections show the six stamens and 
 also the young ovary. After the spore mother-cells have begun 
 »to round off, each stamen should be removed so as to be cut 
 separately. It is very desirable to secure stamens showing the 
 mitotic figures which occur during the division of the spore 
 mother-cell into the four microspores. Since the pollen mother- 
 cells are apt to be in approximately the same stage of develop- 
 ment throughout the anther, it is worth while to determine 
 
Spermatophytes 
 
 131 
 
 whether mitotic figures are present before putting the material 
 into the fixing agent. The procedure indicated for Pi?ius can be 
 followed here. It is not necessary to cut the stamens into 
 pieces before fixing, since they are easily penetrated and infil- 
 trated ; in later 
 stages the sta- 
 mens must not be 
 cut into pieces, 
 since the pollen 
 grains are easily 
 washed out. 
 
 Transverse 
 sections are bet- 
 ter for morpho- 
 logical purposes; 
 but where noth- 
 ing is desired ex- 
 cept thedevelop- 
 ment of pollen 
 grains from the 
 spore mother- 
 cells, much more 
 material can be 
 gotten under a 
 cover by using 
 longitudinal sec- 
 tions. 
 
 Fo r e a r 1 
 
 A^ Salix tristis. X 694. Transverse section of a portion of a young 
 
 StaffeS UD to that ^"'l^^'' showing large sporogenous cells, one layer of tapetal cells (more 
 o r deeply shaded in the figure) . and ' 
 
 from three to four layers of wall cells. 
 Salix petiolaris. X 594. Small portion of transverse section of a nearly 
 mature anther showing five pollen grains, two tapetal cells, and two layers 
 of wall cells. C, Lilium auratum. X,505. Section of a pollen grain show- 
 ing the large tube nucleus and two smaller generative nuclei. Fixed in 
 chromo-acetic acid and stained in safranin-gentian violet-orange. D, 
 Lilium tigrinum. X 505, Pollen grain showing tube nucleus in the mid- 
 dle and a lenticular cell with the generative nucleus at one end of the grain. 
 
 shown in fig. 6j 
 
 A; D,elafield's 
 
 haematoxylin is 
 
 very satisfactory; 
 
 in stages like B, C, and D of the same figure, cyanin and eryth- 
 
 rosin often give excellent differentiation, the tube nucleus taking 
 
 the erythrosin, and the generative nucleus the cyanin, while the 
 
132 
 
 Methods in Pla?it Histology 
 
 starch grains which are often abundant at this stage take a faint 
 pink from the erythrosin. However, the safranin-gentian violet- 
 orange combination shows nuclear details to better advantage, 
 and readily distinguishes the tube and generative nuclei, although 
 it does not give such a striking color contrast as the cyanin and 
 
 erythrosin. The starch grains 
 are brilliantly stained by the 
 gentian violet. 
 
 The mature pollen grain, 
 after shedding, may be pre- 
 pared by the method already 
 described for Pijius. 
 
 Oogenesis. — As in sper- 
 matogenesis, the early stages 
 will be found in preparations 
 of floral development. The 
 origin and development of the 
 macrospore are easily traced 
 in Lilium. In very young 
 stages, before the appearance 
 of the integument, the ovary 
 may be removed from the 
 flower and placed directly in 
 
 ;ad of Aster. 5, pod of Capsella. C, transverse the fixing agent, but in later 
 of ovary of Lilium. The dotted lines show how " ° 
 
 the material should be trimmed before fixing, StagCS, SUch aS are shown in 
 
 figs. 66-^1, strips should be cut off from the sides of the ovary 
 in order to secure more rapid fixing and more perfect infiltration 
 with paraffin. The dotted lines in fig. 64, C, show about how much 
 should be cut off. This is a much better plan than to secure 
 rapid fixing and infiltration by cutting the ovary into short 
 pieces, because the ovules will be in about the same st^ge of 
 development throughout the ovary, and when one finds desirable 
 stages like those from which these photomicrographs were taken, 
 it is gratifying to have these pieces as long as possible. 
 
 In lilies, and other forms with large cells, the entire series 
 shown in figs. 65-yo may be fixed in chromo-acetic acid and 
 
 A , head 
 section of ovary 
 
Spennatophyte 
 
 33 
 
 stained in safranin-gentian violet-orange, but stages earlier than 
 that shown m fig. 65 are more satisfactory if stained in Delafield's 
 haematoxylin. Stages like that shown in fig. 66 and also those 
 between ^^. 66 and fig. 6'j are the most difficult to stain, and 
 only the utmost care and patience will insure first-class prepara- 
 tions. The thread on the nucleus in fig. 66 shows a row of 
 chromatin granules and soon becomes segmented into twelve 
 chromosomes. If the stain is too dense, the thread will probably 
 appear smooth throughout all these stages, but if the staining is 
 successful, the granules are 
 sharply stained by the gen- 
 tian violet, while the linin 
 thread is stained lightly or 
 not at all. In successful 
 staining with cyanin and 
 erythrosin the granules 
 stain blue and the linin red. 
 Stages like figs. 6y and 68 
 are easily stained, and the 
 preparations are exception- 
 a 1 1 y beautiful. Stain for 
 twenty-four hours in safra- 
 nin (the solution in 50 per 
 cent, alcohol is very good), 
 and then rinse in 50 per cent, alcohol until the red color 
 disappears from the spindle, but remains bright in the chro- 
 mosomes and nucleoli ; stain in gentian violet four to eight 
 minutes; rinse in water about thirty seconds ; stain in aqueous 
 orange G fifteen to thirty seconds ; transfer to absolute alcohol, 
 and move the slide gently back and forth in order to dehydrate 
 as rapidly as possible (three to six seconds will usually be long 
 enough); the slide must be taken from the absolute alcohol 
 while the gentian violet is still coming out in streams ; treat with 
 clove oil ten to thirty seconds, and then drain off the clove oil 
 and add a few drops of cedar oil, since the gentian violet fades if 
 much clove oil is left in the preparation. If the cedar oil is 
 
 Fig. 65. Lilium philadelphicum. X 710. 
 
 Apex of nucellus contains a large archesporial cell with a 
 large nucleus. In Lilium this archesporial cell becomes 
 the embryo-sac directly without cutting off any tapetal 
 cell or dividing into potential megaspores. Fixed in 
 chromo-acetic acid and stained in Delafield's haematoxylin. 
 
34 
 
 Methods in Plant Histology 
 
 transparent and has a strong odor, do not use it, but remove as 
 much of the clove oil as possible. Mount directly in balsam. 
 Before transferring to absolute alcohol, a single dip in 95 per 
 cent, alcohol may do no damage and is a matter of economy, 
 since it avoids carrying so much water into the absolute alcohol. 
 
 When the 
 staining is 
 properly done, 
 the chromo- 
 somes will 
 show a bright 
 red color and 
 the spindle a 
 brilliantviolet. 
 If the action 
 of the gentian 
 violet be too 
 prolonged o r 
 poorly ex- 
 tracted, the 
 chromosomes 
 will appear 
 violet or red- 
 dish violet, 
 and such de- 
 tails as the re- 
 lation of spin- 
 d 1 e fibers t o 
 chromosomes 
 will be ob- 
 scured. Stages 
 shown in figs. 
 
 6g and 70 may be found in the same ribbons with stages like^^.y. 
 d/ and 68, and are well differentiated by the same treatment. 
 
 Later stages up to fertilization and the first divisions of the 
 embryo may be fixed and stained as already described, but 
 
 Fig. 66. Lilium philadelphicum. X 710. 
 
 Nucellus with megaspore. The chromatin thread in the nucleus is very 
 distinct. Fixed in chromo-acetic acid and stained in safranin-gentian violet- 
 orange. 
 
Spermatophytes 
 
 135 
 
 cyanin and erythrosin gives a particularly brilliant effect, 
 especially after Carney's fluid. At the stage shown in fig. yi, 
 the male nucleus takes the cyanin, and the oosphere nucleus the 
 erythrosin, although at a slightly later stage they stain alike. 
 
 Fig. 67. Lilium philadelphicum. X 710. Fig. 6S. 
 
 First division of the nucleus of the megaspore. _ (The same nucleus the earlier stages of which are 
 shown in figs. 65 and 66.) Fixed in chromo-acetic acid, and stained in safranin- gentian violet-orange. 
 Fifteen microns. 
 
 For the later stages in the development of the embryo, Delafield's 
 haematoxylin is a far better stain. 
 
 Many of the Compositae, like Aster, Taraxacum, Senecio, and 
 Silphmm, are excellent for a study of the mature sac and the 
 formation of the embryo. The whole head may be cut if trimmed 
 as indicated '\n fig. 64, but for the embryo-sac at the fertilization 
 period, and also for the development of the embryo, it is worth 
 while to resort to the tedious process of dissecting the ovules 
 
136 
 
 Methods in Plant Histology 
 
 out from the ovaries. Stages like those shown in figs. 6^-yo 
 are rather unsatisfactory in Composites. 
 
 Many of the Ranunculaceae are easily studied. Anemone 
 patens, var. Nuttalliana, has a very beautiful embryo-sac, the tgg, 
 
 Fig. 69. Lilium philadelphicum. X 710. 
 
 Megaspore (embryo-sac) containing two 
 daughter-nuclei resulting from the first division of 
 the nucleus of the megaspore. A portion of the 
 spindle still remains between the two nuclei. 
 Fixed in chromo-acetic acid, and stained in safra- 
 nin-gentian violet-orange. Fifteen microns thick. 
 
 Fig. 70. Li! 
 
 philadelphicum. X 710. 
 
 Later stage in the development of the embryo- 
 sac. Each of the two nuclei shown in fig. 69 has 
 divided. The nuclei are much smaller than those 
 in fig. 69. Fixed in chromo-acetic acid, and 
 stained in safranin-gentian violet-orange. Fifteen 
 microns thick. 
 
 synergids, endosperm nucleus, and antipodals being rather large 
 and sharply defined. Hepatica, Caltha, and some species of 
 Ra?iunculus are exceptionally good. 
 
 Development of the Embryo. — The common shepherd's purse 
 {Capsella btirsa-pastoris) is a favorable form for a study of the 
 development of a dicotyl embryo. The stages shown in fig. y2, 
 
Spermatophytes 
 
 137 
 
 A-F, will be found in pods which are about one-eighth of an 
 inch long. These may be put directly into the fixing agent, but 
 stages like G and //are found in pods about three-sixteenths of 
 an inch long, and such stages will be more readily fixed, infil- 
 trated, and cut if the pods are trimmed, as shown '\i\fig. 64, B, 
 before putting them into the fixing 
 agent. Cut sections parallel to the 
 flat face of the pod. Delafield's 
 hsematoxylin, without any contrast 
 stain, gives the best results which 
 we have secured. 
 
 For tracing the development of 
 a monocotyl embryo Sagittaria vari- 
 abilis can be recommended. Mate- 
 rial is abundant, sections are easily 
 cut, except in the latest stages, and 
 it is not difficult to get a complete 
 series. Alisma plaiitago, which is 
 commonly figured in text-books, is 
 extremely hard to cut, especially in 
 later stages. 
 
 Leaves. — Where only a rapid ex- 
 amination is to be made, free-hand 
 sections may be made in great num- 
 bers by using the method employed 
 for pine needles. It is easy to get 
 good sections of leaves which can 
 
 , . -_,..,.-_ Embryo-sac at time of fertilization. A, 
 
 be gotten mtO paraffin, but it is dlffl- the three amipodals. £•, protoplasm of the 
 
 sac. e, polar nuclei fusing to form the 
 
 cult to get tender, succulent leaves endosperm nucleus ; the male nucleus is 
 
 ® about to fuse with the nucleus of the 
 
 into paraffin without distortion. Such oo^phere. z the inner integument. // 
 
 J^ the pollen tube. Y ixed in Carnoy s tluid 
 
 leaves may be cut in celloidin. For fe" n SnVtVcr' (iTat^'f CyS": 
 
 a study of the StOmata, strip a piece Pedia of American Horticulture.) 
 
 of epidermis from the leaf, fix it, stain in Delafield's haematoxylin 
 and erythrosin, pass it gradually through the alcohols, clear in 
 xylol, and mount in balsam. Lily, tulip, hyacinth, and begonia 
 may be suggested as favorable forms. Epidermis from the leaf 
 
 Lilium philadelphicum. X 335. 
 
138 
 
 Methods in Plant Histology 
 
 of the common Sedum pupurasce?is will usually show stomata in 
 all stages of development. 
 
 Stems and Roots. — The earlier stages in the development of 
 vascular bundles in stems and roots are well shown in parafifin 
 
 Fig. 72. Capsella bursa-pastoris, X 400. 
 
 ^ , first division in the embryo cell. ^, quadrants. C, octants, i?, the dermatogen has been cut 
 off. There are eight cells in the suspensor, the lower cell being very large and vesicular. E. differ- 
 entiation in plerome and periblem. The plerome cells are shaded. F, the periblem of the root is com- 
 pleted at the expense of the upper cell of the suspensor. G, the mitotic figure in the suspensor cell indi- 
 cates that the upper suspensor cell by a second contribution is about to complete the dermatogen of the 
 root. H, plerome (shaded), periblem, dermatogen (shaded), and first layer of the root cap. Fixed in 
 chromo-acetic acid and stained in Delafield's hsematoxylin. Ten microns thick. 
 
 sections of young seedlings. The common bean is a favorable 
 form, and it is easy to get material. 
 
 Most herbaceous stems and roots, and also the younger 
 woody stems and roots, give the best results when cut in celloidin, 
 as already described for Pinus. Rumex crispus and Ranunculus 
 repens can be recommended for a study of the vascular bun- 
 dles. The cambium is very sharply brought out by Delafield's 
 
Spermatophytes 
 
 139 
 
 haematoxylin. Petioles or leaf blades of A^zz/Zz^r imbedded in cel- 
 loidin, and stained in safranin and Delafield's haematoxylin, yield 
 extremely beautiful preparations, the sclerotic cells taking a 
 brilliant red and the cellulose a rich purple. The sections should 
 be 20-30 /li thick. 
 
 Stems or petioles of the squash or pumpkin are to be pre- 
 ferred for demonstrating sieve tubes and companion cells. For 
 the more minute de- 
 tails of the sieve plate 
 it is best to cut out 
 small piecesaboutone- 
 fourth of an inch long 
 and one-eighth of an 
 inch square containing 
 the vascular bundle. 
 These pieces can be 
 imbedded in paraffin. 
 
 For demonstrating 
 the phellogen and the 
 tissue developed from 
 it, stems of Geranium 
 or Coleus about one- 
 fourth of an inch in 
 diameter or seedlings 
 of Xanthium canadense 
 about. three-sixteenths 
 of an inch in diameter 
 can be recommended. 
 They can be cut in paraffin, but satisfactory results are more 
 uniformly obtained from celloidin sections. 
 
 The stem of Indian corn, imbedded in celloidin and stained 
 in safranin and Delafield's haematoxylin, affords a good study of 
 monocotyl stem anatomy. 
 
 It must not be forgotten that root tips, besides showing ana- 
 tomical structure, furnish ever-ready material for the study of 
 karyokinesis. An onion thrown into a pan of water will soon 
 
 Fig. 73. Sparganium eurycarpum. X 53. 
 
 Tranverse section of a root. Delafield's haematoxyli 
 and acid fuchsin. Five microns. 
 
140 Methods in Plant Histology 
 
 send out numerous roots. About one-fourth of an inch should 
 be cut off from the tip of the root. Fix and stain as directed 
 for the mitotic figures in Lilium. While the onion is very avail- 
 able material, the figures are not as satisfactory as those to be 
 obtained in the root tips of Tradescantia virginica, Iris versicolor, 
 Podyphyllum. peltatum, Arismma triphyllum, Cypripedium pubescens, 
 and many others. Transverse sections of young roots often show 
 a remarkably regular arrangement of the cells, as can be seen in 
 
 fig' 73- 
 
 We are painfully aware that the directions which have been 
 given in this series of articles are very incomplete, but it is hoped 
 that they will enable the student to devise for himself such 
 methods as particular cases may demand. 
 
CHAPTER XX. 
 LABELING AND CATALOGUING PREPARATIONS. 
 
 THE LABEL. 
 
 The labels shown in fig. 62, on p. 126, show as much as 
 will generally be found desirable. The date of the collection 
 of the material is often needed in addition. The date of making 
 the preparation is of no value unless the student is testing the 
 permanence of stains or something of that sort. It is hardly 
 worth while to write upon the label the names of the stains used, 
 for the student will soon learn to recognize the principal stains. 
 We should say that the first tKmg to write upon a label is the 
 genus and species of the plant ; the next thing would be the 
 name of the organ or tissue, and then might be added the date 
 of collection : e. g., Marchantia polymorpha, young archegonia, 
 April 10, 1 90 1. A hasty sketch on the label will often indicate 
 any exceptionally interesting feature in the preparation. To 
 facilitate finding such a feature, it is a good plan to mark the 
 particular Section or sections with ink, the marking being 
 always on the underside of the slide so as not to cause any 
 inconvenience if an immersion lens should be used. 
 
 CATALOGUING PREPARATIONS. 
 
 As a collection grows, the student will need some device for 
 readily locating any particular preparation. Some have their 
 slides numbered and catalogued, but all devices of this sort are 
 too cumbrous and slow for the practical worker in the labora- 
 tory. After several years' experience with a collection which 
 now numbers about seven thousand preparations, the following 
 method can be confidently recommended : 
 
 Four wooden slide boxes of the usual type will do for a 
 beginning ; they should be labeled : Thallophyta, Bryophyta, 
 Pteridophyta, and Spermatophyta. As the collection grows 
 
 141 
 
142 
 
 Methods i?t Plant Histology 
 
 and new boxes are needed, the classification can be made more 
 definite ; e.g., there should be a box labeled Bryophytes HepaticcB 
 and one labeled Bryophytes Musci. As the liverwort collection 
 grows, three boxes will be necessary, and should be labeled 
 Bryophytes Hepaticce Marchantiales, Bryophytes HepaticcE Jun- 
 germanniales, and Bryophytes //"(f/^/zV^Anthocerotales. It will 
 readily be seen that the process can be continued almost indefi- 
 nitely, and that new slides may be at any time dropped into their 
 proper places. A rather complete label gradually built up in 
 this way is shown '\n fig. 7^/ 
 
 BRYOPHYTES 
 
 HEPATIC^ 
 
 Jungermanniales 
 
 Porella platyphyllum 
 
 Archegonia 
 
CHAPTER XXI. 
 
 A CLASS LIST OF PREPARATIONS. 
 
 Where a regular course in histology is conducted, it is a good 
 plan to give each student at the outset a complete list of the 
 preparations which he is expected to make. In a three-months' 
 course a fairly representative collection of preparations can be 
 made. The availability of material determines what a list shall 
 be. The following list was recently used by one of the writer's 
 classes : 
 
 LIST OF PREPARATIONS. 
 
 THALLOPHYTES. 
 
 ALG^. 
 
 CYANOPHYCE^. 
 
 1. Wasserbliithe. — The principal forms in this material are : 
 
 (a) Coelosphariuni Kutzingianum. — Colonies in the form of hollow 
 
 spheres. 
 (^) Anabcena gigantea. — Filaments straight. Preparations should show 
 
 vegetative cells, heterocysts, hormogonia, and spores. 
 (<r) Anabcena flos-aquce. — Filaments curved. Stain on the slide and 
 
 mount in balsam. 
 
 2. Nostoc. — Eosin or erythrosin. Glycerine. 
 
 3. Oscillaria. — Put living material into 10 per cent, glycerine and allow it 
 to concentrate. 
 
 4. Glo2otrichia. — Eosin, erythrosin, or iron alum-haematoxylin. Glycerine. 
 Should show heterocysts, spores, and vegetative filaments. . 
 
 5. Tolypothrix. — Treat like Oscillaria or Gloeotrichia. Should show hetero- 
 cysts, hormogonia, and false branching. 
 
 CHLOROPHYCEiE. 
 
 6. Volvox. — Stain some in eosin or erythrosin and some in very dilute 
 Delafield's haematoxylin. Mount in glycerine in a cell deep enough to 
 prevent crushing. Try to have antheridia, oogonia, and young colonies 
 in each mount. 
 
 7. Vaucheria. — Eosin, erythrosin, or iron alum-hsematoxylin. Glycerine. 
 Should show antheridia, oogonia, zoospores. 
 
 143 
 
144 Methods in Plant Histology 
 
 8. Hydrodictyon. — Iron alum-hsematoxylin or dilute Delafield's haematoxy- 
 lin. Glycerine. Look for young nets inside the older segments. 
 
 9. Spirogyra. — Iron alum-hsematoxylin. A counter-stain with eosin is a 
 great improvement, if successful. Glycerine. Should show chroma- 
 tophores, pyrenoids, and nucleus. Eosin, erythrosin, or dilute Delafield's 
 haematoxylin will be better for conjugating material. 
 
 I o. Zygnema. — Same treatment as for Spirogyra to bring out chromatophores, 
 pyrenoids, and nucleus. Use iron alum-haematoxylin for reproductive 
 stages. 
 
 1 1. Diatoms. — Living and fossil forms. Balsam. 
 
 12. Oedogonium. — Use iron alum-haematoxylin for oogonia, antheridia, and 
 zoospores. Eosin or erythrosin for caps and other vegetative structures. 
 Glycerine. 
 
 13. Chara. — Oogonia and antheridia may be mounted whole in a cell deep 
 enough to prevent crushing. Glycerine jelly. Paraffin sections of 
 oogonia, antheridia, and apical cell. 
 
 PHiEOPHYCEiE. 
 
 14. Ectocarpus. — Stain some in eosin and some in dilute Delafield's haema- 
 toxylin. Glycerine. Each mount should show both unilocular and mul- 
 tilocular sporangia. 
 
 15. Fucus vesiculosus. — Antheridial conceptacle with paraphyses and anthe- 
 ridia ; oogonial conceptacle with oogonia. Delafield's haematoxylin for 
 paraffin sections. Borax carmine for teased preparations. 
 
 RHODOPHYCEiE. 
 
 J 6. Batrachospermuvi (or Nemalion). — Mayer's hasm-alum. Glycerine. 
 
 Should show trichogyne, carpogonium, and cystocarp. 
 J/. Polysiphonia fibrillosa. — Iron alum-haematoxylin or eosin. Glycerine. 
 Vegetative structure, tetraspores, cystocarps with carpospores, antheridia. 
 FUNGI. 
 MYXOMYCETES. 
 
 J 8. Trichia varia. — Paraffin sections, 5/*. Safranin-gentian violet-orange. 
 
 SCHIZOMYCETES. 
 
 1 9. Bacteria. — Coccus, Bacillus, and Spirillum forms. Stain on cover-glass 
 or slide. 
 
 20. Bacillus anthracis. — In liver of mouse. Paraffin sections, 5 /*. Stain 
 in gentian violet, Gram's method. 
 
 PHYCOMYCaW"ES. 
 
 21. Mucor stolonifer. — Stain young sporangia in eosin or dilute Delafield's 
 haematoxylin. Zygosporic material may be mounted without staining or 
 after a very light staining in dilute Delafield's haematoxylin. Glycerine. 
 
Class List of Preparations 145 
 
 22. Cystopus candidus on Cakile americana. — Select white blisters which 
 have not yet broken open. Paraffin, 5 m- Safranin-gentian violet-orange. 
 
 23. Cystopus bliti on Amarantus retroflexus. — Cut out small portions of 
 leaves in which the oogonia can be seen in abundance. Paraffin, 5 ^x.. 
 
 ASCOMYCETES. 
 
 24. Eurotium. — Eosin or erythrosin. Glycerine. 
 
 25. Pencillium. — Eosin or erythrosin. Glycerine. 
 
 26. Erysiphe cotnmune on Polygonum aviculare. — Strip the fungus from the 
 leaf. Paraffin, 5 /*. Safranin-gentian violet-orange. 
 
 27. Uncinula necator on Ampelopsis quinquefolia. — Eosin or erythrosin. 
 Mount whole and break the perithecia under the cover. Glycerine or 
 balsam. 
 
 28. Xylaria. — Paraffin sections of younger stages. Delafield's haematoxj'lin 
 and erythrosin. Be sure that some section in each mount shows the 
 opening of a perithecium. 
 
 29. Peziza. — Paraffin sections of young apothecia, 5 m or less ; sections of 
 older apothecia, lO/t or 15 /it. Safranin-gentian violet-orange. 
 
 iECIDIOMYCETES. 
 
 30. Pticcifiia graminis. — ^cidium stage on barberry leaf. Paraffin. Cyanin 
 and erythrosin. Uredospore and teleutospore stage in celloidin or paraffin, 
 
 BASIDIOMYCETES. 
 
 3 1 . Coprittus comatus. — Paraffin. Transverse sections of gills showing trama, 
 paraphyses, basidia, and spores. To show the basidium with four spores, 
 the sections should be 15 m thick. For development of the spores, cut 
 5 M or less. Safranin-gentian violet-orange. Boletus, Hydnuni, and 
 Polyporus are treated in the same manner. 
 
 LICHENS. 
 
 32. Parmelia Borreri. — Paraffin, 5/^. Cyanin and erythrosin. 
 
 BRYOPHYTES. 
 HEPATIC^. 
 
 33. Ricciocarpus nutans. — Paraffin, iom or 15^; Delafield's haematoxylin. 
 Archegonia, antheridia, and sporophytes imbedded in the gametophyte. 
 
 34. Marchantia polyniorpha. — Paraffin, lo/i. Archegonia, antheridia, and 
 sporophytes. 
 
 35. Pellia epiphylla. — Paraffin, lo/x. Longitudinal sections of sporophyte 
 attached to gametophyte. Delafield's haematoxylin and erythrosin. 
 
 36. Porella platyphyllufu. — Paraffin, 10 /x. Delafield's haematoxylin. Arche- 
 gonia, antheridia, sporophyte, and apical cell. 
 
 37. Anthoceros Icevis. — Paraffin, 5 or 10 ix. Longitudinal and transverse 
 sections of sporophyte. Safranin-gentian violet-orange. 
 
146 Methods i?i Plant Histology 
 
 MUSCI. 
 
 38. Sphagnum. — Leaf buds. Safranin and Delafield's haematoxylin. Tease 
 and mount in balsam. 
 
 39. Sphagnum. — Capsule. Parafifin. Delafield's haematoxylin and erythro- 
 sin. 
 
 40. Bryum proliferum. — Parafifin. Antheridia, lO/tt ; archegonia, 15 to20ju; 
 capsule, 10 At. 
 
 41. Funaria hygrometrica. — Paraffin. Longitudinal and transverse sections 
 of young capsules. Delafield's haematoxylin. 
 
 42. Funaria hygrometrica or any favorable form. Protonema. Place the 
 well-cleaned material directly into 10 per cent, glycerine and allow it to 
 concentrate. 
 
 PTERIDOPHYTES. 
 
 FILICALES. 
 
 43. Botrychium virginianum. — Paraffin. Stain rhizome, stipe, and root in 
 safranin and Delafield's haematoxylin. Stain sporangia in Delafield's 
 haematoxylin. 
 
 44. Isoetes echinosPora. — Transverse section of stem. Celloidin. Safranin 
 and Delafield's haematoxylin. 
 
 45. Isoetes echinospora. — Paraffin. Longitudinal sections of microsporangia 
 and megasporangia. Safranin-gentian violet-orange. 
 
 46. Pteris aquilina. — Free-hand sections. Safranin and Delafield's haematox- 
 ylin, or iodine green and fuchsin. Balsam. 
 
 47. Pteris cretica. — Paraffin. Transverse sections of leaves with sporangia. 
 Delafield's haematoxylin and erythrosinor safranin-gentian violet-orange. 
 
 48. Adiantum cuneatum.— Prothallia mounted whole in glycerine. Paraffin 
 sections of antheridia and archegonia. 
 
 49. Marsilea quadrifolia. — Parafifin. Longitudinal and transverse sections 
 of sori. Safranin-gentian violet-orange. 
 
 50. Azolla carolinense. — Parafifin. Vertical sections of the whole plant 
 showing micro- and megasporangia. 
 
 EQUISETALES. 
 
 51. Equisetum arvense. — Prothallia in glycerine. Stem tips in parafifin. 
 Transverse section of stem in celloidin. 
 
 LYCOPODIALES. 
 
 52. Lycopodium lucidulum. — Transverse section of stem. Free-hand or in 
 celloidin. Safranin and Delafield's haematoxylin. 
 
 53. Lycopodium inundatum. — Paraffin. Longitudinal sections of strobilus. 
 
 54. Seiaginella. — Paraffin. Longitudinal sections of rather mature strobili. 
 Cyanin and erythrosin. 
 
Class List of Preparations 147 
 
 SPERMATOPHYTES. 
 
 GYMNOSPERMS. 
 
 55. Pinus Laricio. — Transverse section of needles and young stem. Free- 
 hand or cefloidin. Safranin and Delafield's hasmatoxylin. 
 
 56. Pinus strobus. — Free-hand sections of well-seasoned wood. Iodine 
 green and fuchsin, or safranin and Delafield's haematoxylin. 
 
 57. Pinus Laricio. — Paraffin. Ovule with archegonia. Safranin-gentian 
 violet-orange. 
 
 58. Pinus sylvestris or P. Laricio. — -Paraffin. Embryos. 
 
 59. Pinus Laricio. — Paraffin. Longitudinal section of mature staminate 
 strobilus. Safranin-gentian violet-orange. 
 
 ANGIOSPERMS. 
 MONOCOTYLS. 
 
 60. Lilium philadelphicum. — Paraffin, 10 fj.. Transverse sections of ovary; 
 transverse sections of anthers. 
 
 61. Tradescantia virgittica. — Paraffin, 5 and i o joc. Longitudinal sections of 
 root. Stain for mitosis. 
 
 62. Allium. — Paraffin. Transverse sections of older roots for vascular 
 system. 
 
 63. Liliufn longiflorum. — Strip epidermis from leaf and stain for stomata. 
 Cut sections of leaf in paraffin, 10 to 20 /x. 
 
 64. Zea mais. — Celloidin. Longitudinal and transverse sections of stem. 
 
 DICOTYLS. 
 
 65. Nuphar advena. — Celloidin. Section of leaf 1 5 to 40 yn thick. Safranin 
 and Delafield's haematoxylin. 
 
 66. Capsella bursa-pastoris. — Paraffin. Floral development, 5 /*. Embryos, 
 5 to lo/x. Stain both in Delafield's haematoxylin without any contrast 
 stain. 
 
 67. Taraxacum officinale. — Paraffin. Floral development, 5 /u. Embryo- 
 sac, 10 to 15 \t.. 
 
 68. Anemone patens. — Paraffin. Embryo-sac. 
 
 69. Smilax herbacea. — Celloidin. Transverse section of root. Safranin 
 and Delafield's haematoxylin. 
 
 70. Xanthium canadense. — Celloidin. Transverse section of stem of seed- 
 ling to show cambium and phellogen. 
 
 71. Tilia atnericana. — Celloidin or free-hand. Transverse sections of small 
 stems one-eighth to one-fourth of an inch in diameter. SafrAnin and 
 Delafield's haematoxylin. 
 
 In making the mounts the order indicated in the list should 
 not be followed. Begin with free-hand sections, then study the 
 
148 Methods in Plant Histology 
 
 glycerine method, and then devote a large. portion of the time to 
 the paraffin method. Let the celloidin method come last. 
 
 It is neither possible nor desirable that each student should 
 in every case go through all the processes from collecting mate- 
 rial to labeling. Some of the material may be in 70 per cent, 
 alcohol, some in formalin, some in glycerine, and some in paraffin. 
 One student may imbed in paraffin enough of the A?iemone for 
 the whole class, another may imbed the Lilium stamens, and by 
 such a division of labor a great variety of preparations may be 
 secured without a corresponding demand upon the time of the 
 individual. 
 
CHAPTER XXII. 
 
 FORMULA FOR REAGENTS. 
 FIXING AGENTS. 
 
 Carnoy's Fluid. — 
 
 Absolute alcohol, 6 parts. 
 Chloroform, 3 parts. 
 Glacial acetic acid, i part. 
 Strong Chromo-Acetic Solution. — 
 Chromic acid, i g. 
 Glacial acetic acid, i cc. 
 Water, 98 cc. 
 
 Weak Chromo-Acetic Solution (Schaffner's) . — 
 
 Chromic acid, 0.3 g. 
 Glacial acetic acid, 0.7 cc. 
 Water. 99 cc. 
 
 Medium Chromo-Acetic Solution. — 
 
 Chromic acid, 0.7 g. 
 Glacial acetic acid, 0.5 cc. 
 Water, 100 cc. 
 Flemming's Fluid (weaker solution). — 
 ( I per cent, chromic acid, 25 cc. 
 A ■< I per cent, acetic acid, 10 cc, 
 
 ( Water, 55 cc. 
 B. I per cent, osmic acid, 55 cc. 
 
 Keep the mixture A made up, and add B as the reagent is 
 needed for use, since it does not keep well. 
 
 Flemming's Fluid (stronger solution). — 
 
 1 per cent, chromic acid, 45 cc. 
 
 2 per cent, osmic acid, 12 cc. 
 Glacial acetic acid, 3 cc. 
 
 MerkePs Fluid. — 
 
 1.4 per cent, solution of chromic acid, 25 cc. 
 1.4 per cent, solution of platinic chloride, 25 cc. 
 
 Hermann's Fluid. — 
 
 1 per cent, platinic chloride, 15 parts. 
 Glacial acetic acid, i part. 
 
 2 per cent, osmic aid, 4 or 2 parts. 
 
 149 
 
150 Methods m Plant Histology 
 
 Picric Acid. — 
 
 Picric acid, i g. 
 
 Water, or 70 per cent, alcohol, 100 cc. 
 
 Corrosive Sublimate. — 
 
 Bichloride of mercury, 4 g. 
 
 Glacial acetic acid, a cc. 
 
 Water, or 70 per cent, alcohol, 100 cc. 
 
 Formalin (weaker solution). — 
 Formalin, 2 cc. 
 Water, 98 cc. 
 
 Formalin (stronger solution). — 
 Formalin, 4 cc. 
 Water, 100 cc. 
 
 Osmic Acid. — 
 
 Osmic acid, i cc. 
 Distilled water, 100 cc. 
 
 The bottle in which the solution is to be kept, and also the 
 glass tube in which the acid is sold, must be thoroughly cleaned. 
 Break off the end of the tube, and drop both tube and acid into 
 the distilled water. 
 
 STAINS. 
 
 Delafield's Haematoxylin. — "To 100 cc. of a saturated solution 
 of ammonia alum add, drop by drop, a solution of l g. of haema- 
 toxylin dissolved in 6 cc. of absolute alcohol. Expose to air 
 and light for one week. Filter. Add 25 cc. of glycerine and 
 25 cc. of methyl alcohol. Allow to stand until the color is suf- 
 ficiently dark. Filter and keep in a tightly stoppered bottle." 
 (Stirling and Lee.) 
 
 The solution should stand for at least two months before it 
 is ready for using. 
 
 Erlich's Haematoxylin. — 
 
 Distilled water, 50 cc. 
 Absolute alcohol, 50 cc. 
 Glycerine, 50 cc. 
 Glacial acetic acid, 5 cc. 
 Haematoxylin, i g. 
 Alum in excess. 
 Keep it in a dark place until the color becomes a deep red. 
 If well stoppered, it will keep indefinitely. 
 
FormulcB for Reagents 151 
 
 Boehmer's Haematoxylin. — 
 
 Hsematoxylin, i g. 
 Absolute alcohol, 12 cc. 
 Alum, I g. 
 Distilled water, 240 cc. 
 
 The solution A must ripen for two months. When wanted 
 for use, add about 10 drops of A to 10 cc. of B. Stain ten to 
 twenty minutes. Wash in water and proceed as usual. 
 
 Mayer's Haem-Alum. — Haematoxylin, i g., dissolved with 
 heat in 5a cc. of 95 per cent, alcohol and added to a solution of 
 50 g. of alum in a liter of distilled water. Allow the mixture to 
 cool and settle ; filter ; add a crystal of thymol to preserve from 
 mold. (Lee.) 
 
 It is ready for use as soon as made up. Unless attacked by 
 mold, it keeps indefinitely. 
 
 Haidenhain's Iron Alum-Haematoxylin. — This stain was intro- 
 duced by Haidenhain in 1892 and has gained a well-deserved 
 popularity with those engaged in cytological work. Two solu- 
 tions are used, and they are never mixed : 
 
 A. One and one-half to 4 per cent, aqueous solution of 
 ammonia sulphate of iron. (At present we use a 3 per cent, 
 solution.) 
 
 B. One-half per cent, aqueous solution of haematoxylin. 
 Greenacher's Borax Carmine. — 
 
 Carmine, 3 g. 
 
 Borax, 4 g. 
 
 Distilled water, xoo cc. 
 
 Dissolve the borax in water and add the carmine, which is 
 quickly dissolved with the aid of gentle heat. Add lOO cc. of 
 70 per cent, alcohol and filter. (Stirling.) 
 
 Alum Carmine. — A 4 per cent, aqueous solution of ammonia 
 alum is boiled twenty minutes with i per cent, of powdered car- 
 mine. Filter after it cools. (Lee.) 
 
 Alum Cochineal. — 
 
 Powdered cochineal, 50 g. 
 
 Alum, 5 g. 
 
 Distilled water, 500 cc. 
 
152 Methods i?i Pla?it Histology 
 
 Dissolve the alum in water, add the cochineal, and boil ; 
 evaporate down to two-thirds of the original volume, and filter. 
 Add a few drops of carbolic acid to prevent mold. (Stirling.) 
 
 Picro-Carmine. — 
 
 Picro-carmine (picro-carminate of ammonia), i g. 
 Water, 100 cc. 
 
 Eosin. — 
 
 Eosin, 1 g. 
 
 Water, or 70 per cent, alcohol, 100 cc. 
 
 General Formula for Anilins. — Make a 3 per cent, solution of 
 anilin oil in distilled water ; shake well and frequently for a day; 
 add enough alcohol to make the whole mixture about 20 per 
 cent, alcohol ; add i g. of cyanin, erythrosin, safranin, gentian 
 violet, etc., to each 100 cc. of this solution. 
 
 Iodine Green. — 
 
 Iodine green, i g. 
 
 70 per cent, alcohol, 100 cc. 
 
 Methyl Green. — 
 
 Methyl green, i g. 
 Glacial acetic acid, i cc. 
 Water, 100 cc. 
 
 If the preparation is to be mounted in balsam, a slight trace 
 of acetic acid and also a trace of methyl green should be added 
 to the absolute alcohol used for dehydrating. 
 
 Fuchsin. — 
 
 Fuchsin, i g. 
 
 95 per cent, alcohol, 100 cc. 
 
 Water, 100 cc. 
 
 ZiehPs Carbol Fuchsin. — 
 
 Fuchsin, i g. 
 
 Carbolic-acid crystals, 5 g. 
 
 95 per cent alcohol, 10 cc. 1 
 
 Water, 100 cc. 
 
 Fuchsin and Iodine Green Mixtures. — Two solutions are kept 
 separate, since they do not retain their efficiency long after they 
 
 are mixed. 
 
 0.1 g. fuchsin (acid). 
 50 cc. distilled water. 
 
Formulce for Reagents 153 
 
 j 0.1 g. iodine green, 
 \ 50 cc. distilled water. 
 
 iioo cc. absolute alcohol. 
 I cc. glacial acetic acid. 
 0.1 g. iodine. 
 
 Stain in equal parts of A and B, Transfer from the stain 
 directly to solution C and from C to xylol. 
 Another Formula. — 
 
 Acid fuchsin, 0.5 g. 
 Water, 100 cc. 
 Iodine green, 0.5 g. 
 
 ' Water, 100 cc. 
 
 Mix a pipette full of A with a pipette full of B ; stain two to 
 eight minutes ; dehydrate rapidly and mount in balsam. 
 Safranin. — 
 
 Safranin, i g. 
 
 95 per cent, alcohol, 50 cc. 
 
 Water, 50 cc. 
 
 Gentian Violet. — 
 
 Gentian violet, i g. 
 
 95 per cent, alcohol, 20 cc. 
 
 Water, 80 cc. 
 
 Anilin, 3 cc. 
 
 Orange G. — 
 
 Grange G, i g. 
 Water, 100 cc. 
 
 Bismark Brown. — 
 
 Bismark brown, 2 g. 
 
 70 per cent, alcohol, 100 cc. 
 
 Nigrosin. — 
 
 Nigrosin, i g. 
 Water, 100 cc. 
 
 Gram's Solution. — 
 
 Iodine, i g. 
 
 Iodide of potassium, 2 g. 
 
 Water, 300 cc. 
 
INDEX 
 
INDEX 
 
 The references are to pages, 
 
 Acid alcohol, formulae, 35, 43. 
 
 ^cidiomycetes, 84. 
 
 Air bubbles, 32, 
 
 Alcohol, for fixing, 25 ; formulae, 9, 
 
 Algae, 63. 
 
 Alisma, embryo, 137. 
 
 Alum carmine, 40. 
 
 Alum cochineal, 40. 
 
 Ammonia, 35. 
 
 Anabaena, 6j. 
 
 Angiosperms, embryo, 136, 1^8; fertiliza- 
 tion, 135, /j7,- floral development, 129 ; 
 oogenesis, 132; root, 139; spermato- 
 genesis, 130, 131 ; stem, 138. 
 
 Anilins, formulae, 41, 152. 
 
 Antheridia, Chara, 72, 73; ferns, 103, 
 107; liverworts, gi ; mosses, 11, gj. 
 
 Anthers, fixing, 31 ; staining, 131, 132. 
 
 Anthoceros, Q4. 
 
 Anthrax, jy. 
 
 Apparatus, i. 
 
 Archegonia, ferns, 106, 107 ; pines, 121, 
 122, i2s; liverworts, g2; mosses, 97, 98. 
 
 Arisaema, root tip, 140. 
 
 Ascomycetes, 80. 
 
 Aspidium, sporangia, 108. 
 
 Aster, 18, /j2. 
 
 Asterella, 89, gi. 
 
 Azolla, fixing, 31 ; infiltrating, 112. 
 
 Bacteria, yy. 
 
 Balsam, mounting in, 22. 
 
 Basidiomycetes, 87. 
 
 Batrachospermum, 75. 
 
 Beggiatoa, 78. 
 
 Bismark brown, 45. 
 
 Boletus, 88. 
 
 Borax carmine, 40. 
 
 Botrychium, anatomy, 109; prothallia, 
 
 104; sporangia, 108. 
 Bryophytes, 89. 
 Bryum, antheridia, 97 ; capsule, gg. 
 
 Italic figures indicate illustrations. 
 
 Capsella, floral development, 129; pod, 
 
 132; embryo, 138. 
 Carmines, 39. 
 Carnoy's fluid, formula, 26. 
 Cataloguing preparations, 141, 142. 
 Cedar oil, 43. 
 Celloidin, 15; method, 55. 
 Centrosomes, 39. 
 Chara, paraffin sections, 72, 73; glycerine 
 
 mounts, 58. 
 Chlorophyceae, 65. 
 Chorda, 75. 
 Chromosomes, 39. 
 Chromic acid group, 26. 
 Chromo-acetic acid, formulae, 28. 
 Cladophora, 65, by. 
 Clearing, 16, 22; clearing agents, 10. 
 Clove oil, 41. 
 
 Cnicus, floral development, 130. 
 Coelosphaerium, 63. 
 Coleus, stem anatomy, 139. 
 Collema, 88. 
 Conocephalus, 89, 94. 
 Coprinus, 87. 
 Corn, stem anatomy, 139. 
 Corrosive sublimate, 30. 
 Cover-glasses, cleaning, 22. 
 Crucibulum, 88. 
 
 Cutting paraffin, 19 ; celloidin, 56. 
 Cyanin, formula, 8. 
 Cyanin and erythrosin, staining in, 42. 
 Cyanophyceae, 63. 
 Cyathus, 88. 
 
 Cypripedium, root tip, 140. 
 Cyrtomium, sporangia, 108. 
 Cystopus, 79. 
 
 Diatoms, 69, yo. 
 Dehydrating, 14. 
 Desmids, 71, 72. 
 Dichonema, 88. 
 
 157 
 
58 
 
 Methods in Plant Histology 
 
 Ectocarpus, 7j. 
 
 Eosin, formula, 9. 
 
 Equisetum, 115, 116. 
 
 Erysiphe, 83. 
 
 Erythrosin, formula, 8 ; staining in, 42. 
 
 Eurotium, 81. 
 
 Filicinese, 103. 
 
 Fixative, Mayer's albumin, 20. 
 
 Fixing, general hints, 31. 
 
 Flemming's fluid, formula, 28. 
 
 Floral development, 129. 
 
 Formalin, 31. 
 
 Formulae for reagents, 149. 
 
 Fuchsin acid, formula, 8 ; staining in, 44. 
 
 Fucus, 74. 
 
 Fuligo, 78. 
 
 Funaria, antheridia, 97; capsule, g8, gg. 
 
 Fungi, 77. 
 
 Gentian violet, 8, 43. 
 Geranium, stem anatomy, 139. 
 Glceeotrichia, 65. 
 Glycerine jelly, 58. 
 Glycerine method, 58. 
 Gymnosperms, 119. 
 
 Hsem-alum, Mayer's, 8, 38. 
 
 Hsematoxylin, Boehmer's, 38 ; Dela- 
 field's, 7, 35 ; Erlich's, 37; Haiden- 
 hain's iron alum, 8, 38 ; Kleinenberg's, 
 37. 
 
 Hardening, 14. 
 
 Hemitrichia, y8. 
 
 Hepaticse, 89. 
 
 Hermann's fluid, formula, 29. 
 
 Hydnum, 88. 
 
 Hydrodictyon, 66, 67. 
 
 Hypoxylon, 83. 
 
 Imbedding in paraffin, 18; in celloidin, 
 
 55. 
 Iodine green, formula, 8 ; staining in, 44. 
 Iris, root tip, 140. 
 Isoetes, 117. 
 
 Karyokinesis, 39; in Pinus, 119, 122, 123; 
 in Lilium, /jj, 136; in Osmunda, no; 
 in root tips, 139. 
 
 Killing and fixing, 13, 25. 
 
 Labels, 126, 141. 
 
 Laminaria, 75. 
 
 Leptothrix, 78. 
 
 Lichens, 88. 
 
 Liliuin, archesporium, ijj; fertilization, 
 137 ; fixing anthers, 31; fixing ovaries, 
 132; germination of megaspore, /yj, 
 136; megaspore, 134. 
 
 List of preparations, 143. 
 
 Lycoperdon, 88. 
 
 Lycopodinese, 117. 
 
 Lycopodium, 118. 
 
 Marchantia, 89, 90, 91, 92, 94. 
 
 Marsilea, antheridium, 112; archegonium, 
 
 III; imbedding megaspores, 19, 11 2; 
 
 procuring material, in. 
 Merkel's fluid, formula, 29. 
 Mica slips, 65. 
 Microscope, i, 2. 
 Microsphsera, 82. 
 Microtome, hand, / ; sliding, 2. 
 Mildew^s, 82. 
 Mosses, 97. 
 Mucor, 79. 
 Myxomycetes, y8. 
 
 Nigrosin, 45. 
 Nostoc, 65. 
 Nummularia, 83. 
 Nuphar, sclerotic cells, 139. 
 
 Oedogonium, 72. 
 Orange G, formula, 8. 
 Oscillaria, 64. 
 Osmotic apparatus, 15. 
 Osmunda, prothallia, 103; sporangia, 
 no. 
 
 Paraffin bath, ./,- infiltration, 17; removal 
 of, 21; summary of method, 23. 
 
 Parmelia, 88. 
 
 Pellia, antheridia, 91 ; sporophyte, 9j,- 
 spores, 94. 
 
 Peltigera, 88. 
 
 Penicillium, 81. 
 
 Peronospora, 81. 
 
Index 
 
 159 
 
 Peziza, 84. 
 
 Phaeophycese, 73. 
 
 Phycomycetes, 79. 
 
 Physcia, 88. 
 
 Picric acid, 14, 30. 
 
 Pinus, embryo, 121 ; leaves, 124 ; oogen- 
 esis, 121; spermatogenesis, 119, 120; 
 stems and roots, 124, 125; ventral ca- 
 nal cell, 123. 
 
 Podophyllum, root tip, 140. 
 
 Pollen, of Lilium, 131 ; of Pinus, 120. 
 
 Polyporus, 88. 
 
 Polysiphonia, jb. 
 
 Polytrichum, antheridia, 97. 
 
 Prothallia, 11, 26, 103, /oj, ///, 112. 
 
 Protonema, li, loi. 
 
 Pteridophytes, 103. 
 
 Pteris, antheridia, 107; archegonia, 106; 
 prothallia, 103, lo^; rhizome, no; 
 sporangia, 108. 
 
 Ptilidium, qo. 
 
 Puccinia, 84, 83. 
 
 Pythium, 81. 
 
 Ranunculus, 138. 
 
 Razor, 4. 
 
 Reagents, 7. 
 
 Rhodophycese, 75. 
 
 Riccia, 89, 90J 
 
 Ricciocarpus, 89, 9j. 
 
 Rivularia, 64. 
 
 Roots, of seed plants, 138, I3g; of Equi- 
 
 setum, 116; tips for karyokinesis, 139, 
 
 140. 
 Rumex, 138. 
 
 Saccharomyces, 80. 
 
 Safranin, formulae, 8, 42; safranin-gentian 
 violet-orange method, 42. 
 
 Sagittaria, embryo 
 Salix, 130, 131. 
 Saprolegnia, 81. 
 Schizomycetes, 77. 
 Scytonema, 65. 
 Selaginella, 113, 7/7. 
 Silphium, ovules, 1 19 
 Spermatophytes, 119. 
 
 37- 
 
 Sphagnum, 100. 
 
 Spirillum, "jy. 
 
 Spirogyra, 13; fixing, 26, 67, 68; imbed- 
 ding, 69. 
 
 Sporangia, of ferns, 108. 
 
 Staining, 33 ; analytical value, 49 ; Fi- 
 scher's experiments, 50; general re- 
 mark, 47; practical hints, 53; Stras- 
 burger's theory, 48. 
 
 Stains, formulae, 7. 
 
 Staphylococcus, 77. 
 
 Starch, 11. 
 
 Stemonitis, 78. 
 
 Stem, of seed plants, 138. 
 
 Stender dish, 5. 
 
 Stereum, 88. 
 
 Sticta, 88. 
 
 Taraxacum, 129. 
 Teleutospores, 83, 86. 
 Temporary mounts, 11. 
 Thamnidium, 81. 
 Tolypothrix, b^. 
 Tradescantia, root tips, 140 
 Tremella, 88. 
 Trichia, 78. 
 Turn table, 5. 
 
 Uncinula, 82, 
 Uredospores, 83, 86. 
 Ustilago, 86. 
 Ustilina, 83. 
 Usnea, 88. 
 
 Vaucheria, 27, bb. 
 Volvox, 65. 
 
 Wasserbliithe, 6j. 
 
 Washing, 14. 
 
 Webbera, archegonia, q7. 
 
 Xanthium, stem anatomy, 139. 
 
 Xylaria, 83. 
 
 Xylol, removal of, 21. 
 
 Zea, stem anatomy, 139. 
 Zygnema, bg. 
 
UC SOUTHERN REGIONAL UBRARY FAaUTY 
 
 iillllllil 
 
 A 000 865 422 
 
 STAi£,^...aAL SCHOOL,