U.S. DEPARTMENT OF AGRICULTURE • FOREST SERVICE • FOREST PRODUCTS LABORATORY l MADlSON, WIS

U . S . F O R E S T S E R V I C E

R E S E A R C H N O T E

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A u g u s t 1 9 6 4

P R E P A R A T I O N O FD E C A Y E D W O O D F O RM I C R O S C O P I C A L E X A M I N A T I O N

Table of Contents

Page

Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

Preparation of Decayed Samples . . . . . . . . . . . . . . . . . . . . . . 2

Embedding Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2Celloidin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2Paraffin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

Polyethylene Glycol . . . . . . . . . . . . . . . . . . . . . . . . . . 6Freezing. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

Maceration. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

Sectioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

Preparation of Sections for Staining. . . . . . . . . . . . . . . . . . . . . 10Adhesives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10Removal of Embedding Matrix. . . . . . . . . . . . . . . . . . . . . 11

Staining. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11Differentiation of Hyphae and Wood . . . . . . . . . . . . . . . . . . 12

Picro aniline blue . . . . . . . . . . . . . . . . . . . . . . . . 12Pianeze IIIb . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

Differentiation of Wood Structures and Components . . . . . . . . . 13Safranin and fast green . . . . . . . . . . . . . . . . . . . . . 14AzureB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15Zinc-chlor-iodide and phloroglucinol . . . . . . . . . . . . . . 15Iodine-potassium iodide . . . . . . . . . . . . . . . . . . . . . 16

Mounting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17

Microscopical Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . 18Observational . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18Measurement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18

Literature Cited . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21

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PREPARATION OF DECAYED WOOD

FOR MICROSCOPICAL EXAMINATION

By

W. WAYNE WILCOX, Pathologist

Forest Products Laboratory,’ Forest ServiceU.S. Department of Agriculture

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Summary

This report describes some of the methods that were devised or found to beparticularly satisfactory for the microscopical observation of wood in variousstages of decay. These methods include the celloidin, paraffin, and polyethyleneglycol embedding methods, methods for macerating, sectioning, staining, andmounting, and a discussion of several optical systems which facilitate micro-scopical observation of decayed wood. A rapid method for the measurement ofchanges in the amount of cell wall substance visible in cross sections isdiscussed.

Introduction

Because of its generally soft or friable nature, decayed wood may be difficultto prepare satisfactorily for microscopical examination. For example, thinsectioning often requires that an embedding matrix be used to hold the woodstructure intact during the cutting process. Since samples of decayed wood maydiffer greatly in their hardness and strength, it is desirable to have availablea number of methods, each of which may be applicable to a specific set of con-ditions. This report is a survey of some methods used in a detailed study ofchanges caused by decay in the microstructure of wood, which was undertakenin the preparation of a Doctoral thesis.

1Maintained at Madison, Wis., in cooperation with the University of Wisconsin.

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Preparation of Decayed Samples

In this study, suitable decay samples were obtained by following the majorprocedures of the soil-block method outlined in ASTM Standard MethodD1413-61 (1).2 Blocks to be decayed were cut with a thickness of 1/8 inchalong the grain, so that specimens of satisfactory size for microscopical exam-ination could be prepared by simply splitting the blocks along the grain with asharp knife. Reducing the thickness of the blocks increased the uniformity ofdecay and made further cutting across the grain unnecessary. Cutting across thegrain, after the blocks had been decayed, could have produced considerable dis-tortion. The blocks were conditioned and weighed according to the specificationsof the soil-block method..

Following incubation the blocks were again conditioned and weighed to determinethe weight loss that had occurred. Small specimens were then split from theblocks and embedded.

Although air-drying of the blocks had no apparent adverse effects upon woodstructure, hyphae present in the wood collapsed and became distorted. Presumablythis difficulty could be avoided by submerging the blocks in a fixative (7 ,15) im-mediately after removal from culture, but such treatment would prevent accuratedetermination of the weight loss sustained by the blocks.

Embedding Methods

Celloidin

The celloidin method proved to be the most satisfactory procedure forembedding decayed wood. The results fully justified the required expenditure oftime--nearly 2 months for the preparation of fully embedded specimens. Even theslight amount of structure still present in white-rotted wood at weight losses ofover 70 percent was held intact by this method during the cutting of sections4 microns in thickness.

The embedding of specimens in celloidin required no special skills but entailedfollowing a number of steps with reasonable care. The specimens were first eitherair-dried or dehydrated by an ethyl alcohol series (15). The dry specimens wereplaced under vacuum in several changes of absolute ethyl alcohol until they

2Underlined numbers in parentheses refer to Literature Cited at the end of this report.

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absorbed enough alcohol and sank. They were then transferred to several changesof ethylene glycol monomethyl ether (also under vacuum), which proved to be anexcellent solvent for celloidin. Unlike ether-alcohol, it presents no explosionhazard and it evaporates slowly enough to allow convenient handling. The speci-mens in ethylene glycol monomethyl ether were placed in a stoppered bottle in anoven at 52° to 54° C. for 2 days or more. They were transferred successivelythrough 2, 4, 6, 8, and 10 percent3 solutions of celloidin in ethylene glycol mono-methyl ether maintained at 52° to 54° C., and were allowed to remain in eachgrade for a minimum of 2 days. With a film of celloidin around them at all times,the specimens were removed from 10 percent celloidin, transferred with the faceto be sectioned downward, to hardening chambers (fig. 1),4 and covered with a12 percent solution of celloidin in ethylene glycol monomethyl ether. The celloidinin the hardening chambers was allowed to concentrate by evaporation at roomtemperature. Additional celloidin was added periodically in order to keep thechambers full and to allow for further concentration. During this process, thesurface of the celloidin became dry and hard. The configuration of the surfacecrust was used as an indicator of the concentration of the celloidin to be added.Thus, when the crust of celloidin formed a ring around the edge of the chamber,a 20 percent solution of celloidin in ethylene glycol monomethyl ether was added,but when a solid plug of hard celloidin formed at the mouth of the chamber a40 percent solution was added. The crust was removed with a knife prior to eachaddition to prevent the trapping of air bubbles in the matrix. The 40 percentcelloidin was allowed to evaporate at room temperature until it became quitefirm, or until it, began to pull the bottom of the chamber inward. At this stagethere was little danger of the celloidin becoming too hard, provided the shrinkagedid not begin to distort the specimen, since it was found that very hard celloidinstill could be successfully sectioned merely by increasing the period of storagein glycerin-alcohol.

The embedded specimen was next removed from the hardening chamber. Witha knife, the chamber was cut away from the glass slide and loosened from itscontents. The embedded specimen was then removed from the chamber bypressing on one end and placed on a wooden mounting block of such size that itcould be firmly held in the microtome. The mounting block had been impregnatedpreviously with 15 to 25 percent celloidin in ethylene glycol monomethyl ether.The embedded specimen was then surrounded with 40 percent celloidin and sub-merged in chloroform overnight, or until completely hardened throughout.

3Throughout this report, the term “percent.” as applied to solutions of solid reagents, denotes the

number of grams of solute per 100 milliliters of solvent. The percentage concentration of liquidreagents is expressed on a volumetric basis.

4The idea from which these chambers were developed was originally that of Dr. Catherine Duncan

of the Forest Products Laboratory.

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Figure 1.-- Celloidin hardening chambers. These consisted of two 1/2-inch-long segmentsof plastic tubing 1/2 inch in outside diameter adhered to a 1- by 3-inch glass microscopeslide with a room-temperature-curing nitrile-phenolic resin in organic solvents. Theuse of a glass slide with one etched end allowed the contents of the chambers to belabeled with a pencil. Such chambers facilitated the handling of large numbers ofspecimens and allowed the specimens to be oriented for sectioning at a stage in theembedding process when they could be easily observed.

M 126 011

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After removal from the chloroform, the hardened material was trimmed down forsectioning and stored in a 50-50 mixture of glycerin and 95 percent ethanol. Thelonger the material was stored in glycerin-alcohol, the better were its cuttingproperties. Sectioning was performed on a sliding microtome.

Paraffin

Paraffin embedding was compared with celloidin for specimens in very advancedstages of decay, since with paraffin there would be less opportunity for dis-tortion or damage due to handling. Paraffin proved quite satisfactory for materialin advanced stages of decay but was inferior to celloidin embedding for sound woodor wood in early stages of decay. An advantage of paraffin embedding was thatserial sections could be obtained by sectioning on a rotary microtome.

The methods of Sass (15) for paraffin embedding, which employ tertiary butylalcohol (TBA), proved satisfactory for application to decayed wood. Only minormodifications suggested by Rogers (14) were utilized. The material to be em-bedded was gently split into small blocks approximately 1/8-inch square and1/8 inch to 3/16 inch in length along the grain. The specimens were dehydratedor moistened, depending upon the beginning moisture content, by means of anethanol series (15), to the stage of 60 percent (by volume) ethanol. They wereevacuated until they sank in the first liquid used in the series. The samples werethen transferred through the TBA series of Sass (15) (table 1).

Table 1.--Dehydration solutions using tertiary butyl alcohol (TEA)1

95 percent ethanol Absolute ethanol TBA Distilled water

Grade 1 50 ml. 10 ml. 40 ml.Grade 2 50 ml. 20 ml. 30 ml.Grade 3 50 ml. 35 ml. 15 ml.Grade 4 50 ml. 50 ml.Grade 5 25 ml. 75 ml.

1Reprinted by permission from John E. Sass, Botanical Microtechnique, 3rd ed., The Iowa State

University Press, Ames. 1958.

The samples were placed under vacuum briefly after each transfer and werestored in each grade for 4 hours or more. From grade 5 they were transferredto several changes of anhydrous TBA and then to a 50-50 mixture of anhydrousTBA and paraffin oil, using the same evacuation and storage schedule as withgrades 1 through 4. The bulk of the TBA-paraffin oil then was decanted into a

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waste container. The rest, along with the specimens, was gently poured into avial on top of molten embedding paraffin which had been cooled sufficiently sothat a film strong enough to support the specimens had formed across its surface.The paraffin was of a type with a high melting point, commercially supplied fortissue embedding. The vial containing the specimens was then stored in an ovenhot enough to keep the paraffin completely fluid. Upon the melting of the surfacefilm, the specimens dropped into the pure paraffin. Periodically the paraffin wasdecanted into a waste container and replaced with fresh, molten paraffin. Theparaffin was changed four or five times, or until the odor of TBA was no longerdetectable.

The specimens, along with the final change of paraffin, were poured into a paperboat placed upon a casting table--a heavy steel plate heated at one end with asmall flame (15). The boat was placed close enough to the flame to allow theparaffin to remain molten until the specimens had been arranged. By sliding theboat away from the flame, a thin film of solidifying paraffin was formed at thebottom of the boat, which aided in maintaining the specimens in the desiredpositions. As soon as the paraffin had solidified enough to hold the specimensfirmly, the boat was placed in a freezing compartment to quickly and thoroughlyharden the paraffin. When the paraffin had become hard, the paper boat was re-moved, and blocks of paraffin containing individual specimens were cut out with ahacksaw blade. Then the specimens were affixed to mounting blocks previouslyimpregnated with molten paraffin, by melting portions of the paraffin with a hotneedle or scalpel (15).

Polyethylene Glycol

An alternative embedding method for use with the rotary microtome, employingpolyethylene glycol (PEG), was tried on a limited number of specimens. Thismethod was of interest because of certain reported advantages: (1) green or moistsamples may be used directly, reducing the problem of distortion due todehydration, and (2) the method is rapid and requires a minimum of specimenhandling. The limited utilization of PEG embedding in the thesis study was insuf-ficient for a thorough evaluation of the method, but the evidence obtained showeddistinct promise for application to relatively soft or decayed wood. Also, itappeared that it might be especially useful for studying fungal hyphae in woodsince, in the absence of severe dehydration, it should minimize distortion andcollapse of the hyphae.

The method utilized was taken without modification from Gjovik.5 He foundcommercial PEG with a molecular weight of 1450 most suitable for histologicalapplication. However, the ability to form good ribbons varied considerably from

5Gjovik, L. R. The application of fluorescence microscopy to the study of the permeability of

aspen. (Populus tremuloides Michx.) M.S. Thesis. University of Minnesota. 1961.

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batch to batch, and it was suggested by Gjovik that samples be tested for thisproperty before a batch is selected for histological use.

Prior to embedding, the specimens were thoroughly water saturated undervacuum. They were then placed in a 50-50 mixture of PEG-1450 and distilledwater at 50° C. for 2 hours. Next the 50-50 mixture was decanted, replaced withmelted, pure PEG-1450, and stored at 50° C. for 2 hours. The old PEG-1450 wasthen exchanged for fresh liquid and stored at 50° C. for 4 to 6 hours. Althoughadditional evacuation while in pure PEG was recommended by Gjovik, this pro-cedure probably was necessary only if air had been incompletely removedinitially. Individual specimens were mounted by placing them in the cavity formedby wrapping a wooden mounting block with masking tape so that it extended beyondthe end of the block However, a number of specimens may be prepared at onetime by the use of paper boats as in the paraffin technique. The PEG was thenhardened by placing the mounted specimens, or the paper boats, in a refrigeratorfor 30 to 40 minutes until the PEG exhibited a grayish-white translucence. Theembedded and mounted specimens were then trimmed for sectioning and storedin a desiccator containing calcium chloride. Gjovik found that special techniques,such as cooling of the microtome knife or softening or cooling of the embeddedspecimen, were unnecessary for satisfactory sectioning.5 He also found thatacceptable sections could be obtained in less than 1 hour by using an abbreviatedschedule consisting of the same steps as outlined above but allowing only 15 to20 minutes of storage at each stage.

Gjovik5

suggested a potassium dichromate-gelatin adhesive, instead of Haupt’sadhesive, for use with sections of PEG-embedded material. This avoided the useof formalin which dissolved the embedding matrix (discussed in the section“Preparation of Sections for Staining”).

Freezing

Probably the simplest but least effective “embedding” method utilized in thisstudy was the process of freezing a water-soaked specimen prior to cutting (7 ,15).This method offered support to weak materials during cutting, but the support waslost when the ice melted soon after the section had been cut. Freezing of thespecimens impregnated and surrounded with water can be accomplished inseveral ways (15). In trials on decayed wood a carbon dioxide freezing apparatuswas used. Some additional support was obtained during cutting by deflecting thejet of carbon dioxide so that it cooled the knife in addition to the specimen. Aspecial attachment is commercially available for this procedure.

The specimen to be sectioned was placed under vacuum in water until itsank. The specimen was placed upon the freezing attachment and frozen;water was dripped on it so that it became thoroughly surrounded by ice.

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Periodic reapplication of carbon dioxide was necessary during sectioning tomaintain the specimen in the frozen state. The specimens were sectioned on asliding microtome, and the sections were handled with a moist brush in the samemanner as sections from nonembedded specimens. It has been reported that addi-tional support may be obtained if, instead of pure water, an aqueous solution of asubstance such as gelatin or agar is utilized (7,15).

Maceration

It was found in the study that not all microscopical features of decayed woodwere clearly visible in sections. For example, bore holes were best observed byexamining the entire cell walls of isolated wood elements; the various woodelements were separated for such observation by the process of maceration.

Several methods for the maceration of wood are available (5,19), but Jeffrey’smethod (7,15,19) is simple and proved entirely satisfactory in this study. The bestresults were obtained when the wood to be macerated was gently split into sliversabout 1/32-inch thick. The slivers were evacuated in the macerating fluid until‘they sank and were placed in an oven at 40° to 50° C. overnight. Jeffrey’s macer-ating fluid consists of equal amounts of 10 percent (by volume) aqueous nitricacid and 10 percent aqueous chromic acid. Small glass beads were added to thevials containing the macerating fluid and the wood samples, and the vials wereshaken to aid separation of the elements. If the wood did not subdivide easily atthis stage, it was allowed to settle, and the darkened macerating fluid was drawnoff and replaced with fresh fluid. The samples in fresh fluid were stored again inthe oven overnight.

This procedure was repeated until satisfactory maceration was achieved. Thesamples were then washed with water using a low-speed centrifuge to settle thembetween washings. Each macerated sample was stored in 50 to 70 percent ethanol.For microscopical examination, the sample was observed under a sealed coverglass, using the storage fluid as a wet mount (discussed in the section on“Mounting”).

Sectioning

The use of the rotary microtome on woody tissues may cause some degree ofcrushing and distortion; therefore, the sliding microtome was utilized except for

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the limited observations on specimens embedded in paraffin or polyethyleneglycol. Such specimens were cut dry, and a ribbon of serial sections was formed.The knife had to be sharp and free of nicks to minimize tearing and crushing ofthe specimen. The use of a razor blade in place of the microtome knife was foundto be very convenient and highly satisfactory, since the blade could be discardedwhen dull or nicked. Further details of rotary microtomy are discussed bySass (15).

The sliding microtome was used for nonembedded and for celloidin-embeddedspecimens. A conventional, wedge-shaped microtome knife was used with thisinstrument. The slice angle, the angle the knife makes with its direction oftravel, was kept as small as possible, making fullest use of the slicing action ofthe knife. The clearance angle, the angle between the bottom surface of the sharp-ened bevel of the knife and the surface of the specimen, was also kept small. Thebottom surface of the bevel was not allowed to be completely parallel to thespecimen’s surface and the back of the bevel was never placed below the cuttingedge. However, too steep a clearance angle resulted in curling of the section.Detailed discussion of the above concepts is presented by Richards (12).

The condition and position of the microtome knife were especially critical whencutting sections of the thinness required in this study. Difficulty in sectioning withthe sliding microtome usually could be attributed to some of the following factors.The most common cause of substandard sections was a dull or imperfect knifeedge. Dullness of the knife edge was indicated by disintegration of the sectionsduring cutting or variations in thickness from one section to the next. Coarsenicks in the blade edge usually resulted in the division of the section into severalpieces. Fine nicks were hard to detect but often were the cause of weak sectionswhich fragmented when touched by the brush. Such fine nicks usually caused smallscratches in the specimen, detectable only when the specimen surface was wipeddry. These difficulties were overcome by restropping or in severe cases rehoningthe knife edge. However, if defects appeared only in localized areas of the knife,other regions could be used for sectioning. Occasionally tearing of the sectionsoccurred when the knife was in perfect condition. It was found that such tearingcould be overcome by diluting the glycerin-alcohol, used for lubrication withadditional 95 percent ethanol, by applying less pressure to the brush while guidingthe sections, or by decreasing the slice angle of the knife.

Nonembedded specimens were saturated with water or 50 to 70 percent alcoholby evacuation prior to sectioning. Very hard specimens in the water-soaked con-dition were cut with ease by allowing a small jet of steam to flow over thespecimen surface during cutting. Care should be taken with this technique,however, since the sliding surfaces of some microtomes rust when exposed to

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steam. In order to avoid tearing and distorting the section, it was touched with asoft-bristled brush during the cutting and held lightly in the approximate positionit occupied while a part of the specimen. A brush with bristles mounted in plasticrather than metal was used in order to avoid accidental damage to the knife edge.The blade surface (especially its cutting edge), the specimen, and the brush allwere lubricated by frequently filling the brush with the liquid so that the sectionliterally floated off of the specimen during cutting. The liquid used was water ordilute alcohol for water-soaked specimens, or glycerin-alcohol for celloidin-embedded specimens.

Difficulty was experienced in obtaining intact sections of very hard specimens,such as sound southern pine summerwood, even when they were embedded incelloidin. An attempt was made to alleviate this problem by using the Cellulose-Tape Method of Bonga (2). This method consists of allowing a loop of cellulosetape to hang down over the specimen mounted in the sliding microtome. The tapeis firmly affixed to the thoroughly dried surface of the specimen, and the sectionis cut with a slight tension applied through the tape to its leading edge, to keep thetape from sticking to the knife. Sections made by this method were of high qualityas long as they were left on the tape and viewed directly through it. However, itwas not possible to satisfactorily remove the tape and adhesive from the sections.Soaking in acetone was helpful, but this process resulted in curled, brittle sectionswhich were difficult to handle.

In general, thin sections produced sharper microscopical images than did thicksections; therefore, in this study, celloidin-embedded specimens were sectionedat a thickness of 4 microns. Thinner sections were easily obtained with suchmaterial but proved extremely difficult to handle because they were not easy tosee. Since it was difficult to obtain sections as thin as 4 microns from specimenswhich were not embedded in celloidin, paraffin- and polethylene glycol-embeddedspecimens were sectioned at thicknesses of 8 to 12 microns and nonembeddedspecimens at thicknesses of 10 to 20 microns. Such thick sections were necessaryfor critical observation of irregular structures such as hyphae. The thick sectionsalso stained more densely with many of the stains used than did the thin sections.

Preparation of Sections for Staining

Adhesives

For thin sections, and especially for sections of wood in more advanced stagesof decay, it was found most convenient to adhere the sections to the slide prior to

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staining. For attaching the sections, Haupt’s adhesive proved satisfactory (7).This adhesive was prepared by dissolving 1 gram of purified, finely dividedgelatin in 100 milliliters distilled water at a temperature not exceeding 30° C.When dissolved, 2 grams of phenol and 15 milliliters of glycerin were added. Theuse of purified components made the difficult process of filtration unnecessary.To use this adhesive, a small portion, less than a drop, was placed upon a cleanglass slide and spread with the finger, leaving a barely perceptible layer. Severaldrops of 4 percent (by volume) aqueous formaldehyde were placed on this surfaceand the sections, which previously had been washed in water to remove most ofthe glycerin or alcohol, were added and arranged. The slide was then placed upona warming table or in an oven at 40° to 50° C.

Gjovik5 suggested the use of a potassium dichromate-gelatin adhesive withpolyethylene glycol-embedded specimens. This adhesive consisted of an aqueoussolution of 0.5 percent potassium dichromate and 0.5 percent gelatin, with theexcess potassium dichromate removed by dialysis. For use, a light film of thepotassium dichromate-gelatin adhesive was spread on the slide. The sections werecarefully placed upon this moist surface, and the slide was dried upon a warmingtable for 6 to 8 hours at 50° C.

Removal of Embedding Matrix

If hyphae or ‘other such structures present in the cell lumina were to beobserved, it was found desirable to hold these in place by allowing some of theembedding matrix to remain in the sections, provided that the stains used pene-trated the matrix sufficiently. If the wood structure was of primary interest,however, most of the embedding matrix was removed, prior to staining, toimprove image clarity.

Celloidin was removed by soaking for 24 to 48 hours in ethylene glycolmonomethyl ether at room temperature, or for shorter periods at 50° to 54° C.Paraffin was removed by briefly soaking the slides in xylene. Polyethyleneglycol was quickly removed with water.

Staining

Good reviews of histological stains and staining are given by Sass (15), Connet al. (4), and Johansen (7), and histochemical methods by Jensen (6). Certainmethods found to be suitable for the study of decayed wood and adaptations of

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more general methods are discussed here. The Color Index numbers of thestains used are in accordance with the second edition of “Staining Procedures” (4 ) .

Differentiation of Hyphae and Wood

Picro aniline blue.--This method provided the best results in the differentiationof hyphae; it was also used successfully to reveal bacteria in wood.6 Originallydeveloped by Cartwright (3), it is presented in this report in somewhat modifiedform. With this procedure, wood cell walls appear pink, and hyphae or bacterialcells appear blue.

Stock Solutions

safranin: 1 percent aqueous safranin 0 (C.I. 50240).picro aniline blue: 25 milliliters saturated, aqueous

aniline blue (C.I. 42755), plus100 milliliters saturated, aqueous picric acid.

Staining Schedule

1. Stain in safranin--2 minutes.Place sections in 10 milliliters of distilled water andadd 3 drops of the safranin stock solution;

2. Wash in water.

3. Stain in picro aniline blue.Place sections in 10 milliliters of distilled water and add5 drops of the picro aniline blue stock solution. Heat on amedium-warm hot plate until the stain begins to steamslightly.

4. Wash in water.

5. Mount in water, 50 percent glycerin, etc.

The sections may also be dehydrated by means of an alcohol series to producepermanent mounts (discussed in the section on “Mounting”).

6Knuth, D. T. Bacteria associated with wood products and their effects on certain chemical and

physical properties of wood. Ph. D. Thesis. University of Wisconsin. 1964.

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Pianeze IIIb.--This is an alternative method of differentially stainingfungal hyphae in wood and has the advantage of requiring only a single stainapplication. However, the differentiation proved to be less striking than that ofthe picro aniline blue method in some material. The following procedure is anadaptation of Vaughan’s method (17), used extensively for routine examinations atthe Forest Products Laboratory.

Stock Solution

Pianeze IIIb:malachite green (C.I. 42000)acid fuchsin (C.I. 42685)martius yellow (C.I. 10315)distilled water95 percent ethanol

1.0 g.0.5 g.0.05 g.

150.0 ml.50.0 ml.

Staining Schedule

1. Wash in water.

2. Stain in stock solution for 10 to 45 minutes, dependingupon color desired.

3. Wash in water.

4. Decolorize in acid alcohol (95 percent ethanol, to whicha few drops of concentrated hydrocholoric acid have beenadded).

5. Wash with 95 percent ethanol.

6. Clear with carbol-turpentine (400 milliliters melted phenol,600 milliliters oil of turpentine).

7. Wash with xylene.

8. Mount.

Differentiation of Wood Structures and Components

Various histological and histochemical methods have been devised for theexamination of plant materials, including wood. Several of these common methods

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were also found in this study to greatly aid in the detection of changes in woodstructure and composition resulting from decay.

Safranin and fast preen. --This is one of the commonest staining proceduresapplied to wood and other plant material. It was found to be very useful because itwas effective and nearly foolproof in application. The only critical length of timein the schedule was the period in which the sections were in contact with fastgreen, By this method heavily lignified portions of the cell wall stain red andless lignified portions (the secondary walls of many woods) green. Even in ad-vanced stages of decay, this differentiation was maintained. Although thesestains are not specific for lignin or cellulose (6), the staining reactions obtainedwith them correlated well with data from more specific methods and with theexpected chemical composition of the decayed wood. Numerous modifications ofthis staining procedure have appeared, but the schedule described belowprovided the author with fairly uniform results on a variety of materials.

Stock Solutions

safranin: 1 percent solution of safranin 0 (C.I. 50240) in50 percent ethanol; filtered.

fast green: 1 percent solution of fast green (C.I. 42053) inclove oil-alcohol (1 part clove oil, 9 parts95 percent ethanol); filtered.

Staining Schedule

1. Stain in safranin--1 hour (or more).

2. Wash in 50 percent ethanol.

3. Wash in 95 percent ethanol.

4. Stain in fast green--1 minute (with intermittent agitation).

5. Wash in 95 percent ethanol.

6. Wash in absolute ethanol.

7. Clear by covering the sections with several drops ofclove oil for a minimum of 5 minutes.

8. Flush with xylene.

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9. Mount.

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Azure B. --According to Jensen (6) this method is fairly specific for ligninand for nucleic acids. All tissues which were stained red by the safranin and fastgreen technique were found to stain blue-green with azure B, and as with safraninthe staining properties were retained into advanced stages of decay. A full dis-cussion of this staining method is offered by Jensen (6), while only the stainingschedule is outlined below.

Stock Solutions

azure B: 0.25 milligrams azure B (C.I. 52010) per milliliterof citrate buffer at pH 4.0.

citrate buffer (pH 4.0): 2.7 grams citric acid and 2.1 gramssodium citrate dissolved in100 milliliters distilled water.

Staining Schedule

1. Hydrate sections in water.

2. Stain in azure B stock solution for 2 hours at 50° C.

3. Wash in water.

4. Place in anhydrous tertiary butyl alcohol for 30 minutes.

5. Wash in xylene.

6. Mount.

Zinc-chlor-iodide and phloroglucinol. --Zinc-chlor-iodide has been suggestedfor determining the presence of cellulose and phloroglucinol for the presence oflignin. Both methods are discussed by Jensen (6). Apparently neither technique isspecific for the component it is designed to indicate. Furthermore, it was foundin this study that the residual lignin in heavily decayed wood failed to react withphloroglucinol. This information may be of importance as an indication of subtlechanges occurring in the lignin molecule as a result of decay, but it also rendersthe technique less useful as a stain for general application to decayed wood. Dueto swelling of the cellulose, zinc-chlor-iodide had the disadvantage of distortingthe section.

In general, secondary walls became bluish, and the compound middle lamellabecame red, except in advanced stages of decay. In very thin sections, however,

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the coloration was found to be too faint to be observed. In this study a method,described below, was devised for the application of these two methods incombination.

Stock Solutions

zinc-chlor-iodide: according to Venning(18), dissolve 15 gramszinc chloride in 10 milliliters of distilledwater, then add 0.15 grams potassium iodideand 0.25 grams iodine. The iodine is in excessto insure a concentrated solution.

phloroglucinol: according to Jensen (6), saturated aqueous solution.combination stain: 1 part 20 percent (by volume) hydrochloric

acid, 2 parts phloroglucinol stock solution,3 parts zinc-chlor-iodide stock solution.

Staining Schedule

1. Cover sections with zinc-chlor-iodide stock solution.

2. Allow to stand for about 30 minutes, until the sectionsappear black.

3. Remove the zinc-chlor-iodide with a blotter and replacewith the combination stain stock solution.

4. Cover and observe as a wet mount.

Iodine-potassium iodide.--This method proved useful for observing the locationof starch in both sound and decayed wood, but it also is not specific for thiscomponent (6). Starch is colored dark blue to black by this reagent. The formu-lation reported here is that of Jensen (6).

Stock Solution

iodine-potassium iodide: dissolve 2 grams potassium iodide in100 milliliters distilled water. Thendissolve 0.2 grams iodine in thepotassium iodide solution.

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Staining Schedule

1. Mount sections in the stock solution and observe directlyas a wet mount.

Mounting

Both permanent and temporary types of mounting procedures were used in thestudy. Permanent mounting provided a record for later reference, and it had theadded advantage that the subject was dehydrated and cleared, thereby increasingits resolution. For some purposes, however, wet mounts were required.

By sealing the edges of the cover glass to prevent evaporation, the length ofservice of wet mounts was considerably extended. A commercial product, amixture of lanolin and a resin, was found satisfactory for such application. Thissubstance was melted and applied by means of a heated, bent metal rod.

For permanent mounting, a synthetic resin mounting fluid was used. This resinwas applied in the same manner as Canada balsam but had certain advantages.The mounting fluid has no color which was an advantage for color photography,required no cleaning of the slides after application since a relatively Smallamount was needed, and it hardened in about 30 minutes at room temperature. Inpreparation for permanent mounting, the sections were dehydrated by means ofan alcohol series, cleared for 5 minutes or more in clove oil, and washed withxylene. Excess xylene was blotted away, but enough was left so that the sectionswere thoroughly covered. A clean cover glass was passed quickly through a flame,and a large drop of resin was placed at one end of it. The cover glass was loweredto the slide so that the end containing the drop of resin contacted the xylene.Any bubbles occurring at this stage were removed by raising the other end ofthe cover glass and waiting momentarily. The raised end of the cover glass wasthen slowly lowered to the slide. Excess xylene and resin were removed byblotting the edges of the cover glass. Troublesome bubbling could be avoided byusing more xylene on the sections or by thinning the mounting medium. If bubblesformed, they usually disappeared during the process of removing the excessxylene. A small brass or lead weight was placed on top of the cover glass forabout 30 minutes to 1 hour at room temperature, after which the slide was readyfor use.

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Microscopical Methods

Observational

In addition to chemical means of differentiating wood structures and componentsin various stages of decay, several optical-microscopical methods also gave gooddifferentiation (11,16).

The use of a phase-contrast optical system with unstained sections aided inlocating voids and other features of the cell wall, which were denoted byvariations in contrast resulting from differences in refractive index. Cracks,bore holes, and other voids in the cell wall appeared dark and in sharp contrastto the light cell-wall substance.

Polarized illumination aided the detection of alterations in the quantity andlocation of the birefringent, crystalline cellulose. By this method areas of thecell wall containing crystalline cellulose appeared bright, while noncrystallineareas of the wall were dark when viewed between crossed polarizing filters.

The application of polarized light, in conjunction with a first-order redcompensator, greatly facilitated the observation of macerated, decayed wood forchanges in quantity and location of crystalline cellulose, and for the occurrenceand distribution of bore holes and similar features. This method, as applied tosound wood, was described by Ritter (13). It gave crystalline areas of the cellwall a yellow or blue color against a magenta background, while voids and non-crystalline areas of the wall also were magenta.

Absorption of specific wavelengths of ultraviolet radiation was utilized tofollow lignin distribution in sections of sound and decayed wood. When irradiatedwith wavelengths of about 2800Å lignin appeared dark, due to intense absorptionof the radiation, while the carbohydrates appeared light. This method is discussedby Lange (9,10).

Measurement

Numerous dimensional changes are known to occur in wood structure as aresult of decay; therefore, a method was sought for the measurement of suchchanges. The magnitude of dimensional changes in individual cells was found tobe highly variable. This variability necessitated the measurement of largenumbers of cells in order to arrive at satisfactory average values. The common

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methods for determining cross-sectional area, utilizing ocular micrometry,planimetry, or photographic weighing, were considered too time consuming toprovide the necessary replication. Instead, a method developed by Ladell (8) formeasuring the area of features in cross sections of wood was employed (fig. 2).This method proved to be quite rapid, making it possible for a large number ofcells to be measured.

The method consisted of projecting the image of a section from the microscopeonto a white card containing 100 pinholes punched on grid coordinates determinedfrom a table of random numbers. The card containing the randomly spaced pin-holes was illuminated from below, so that the holes appeared as spots of lighton the projected image of the specimen. In the area occupied by the 100 spots,the number of spots falling upon a given structure provided a measure of thepercentage of the area occupied by the structure.

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Figure 2. --Adaptation of alight microscope for measurement of cross-sectional area on microtomesections. (A) prism; (B) mirror: (C) opaque sampling card with sampling field (outlined byblack line) containing 100 randomly spaced pinholes; (D) light-tight illuminating box containtnga 15-watt bulb and aluminum-foil reflector; (E) light shield; (F) hand counter. This apparatuswas adapted from a method developed by Ladell.

M 126 765

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Literature Cited

1. American Society for Testing and Materials.1961. Standard method of testing wood preservatives by laboratory soil-

block cultures. ASTM Designation D 1413-61. Book of ASTMStandards, Part 6, pp. 1013-1026. Philadelphia.

2. Bonga, J. M.1961. A method for sectioning plant material using cellulose tape.

Canad. Jour. Bot. 39:729.

3. Cartwright, K. St. G.1929. A satisfactory method of staining fungal mycelium in wood sections.

Ann. Bot., London 43:412-413.

4. Conn, H. J., Darrow, Mary A., and Emmel, V. M.1960. Staining Procedures Used by the Biological Stain Commission.

2nd ed. Williams and Wilkins Co., Baltimore.

5. Forest Products Laboratory.1960. Annotated list of references on the preparation of wood for

microscopic study. U.S. Forest Prod. Lab. Rep. No. 1939.

6. Jensen, W. A.1962. Botanical Histochemistry. W. H. Freeman and Co., San Francisco.

7. Johansen, D. A.1940. Plant Microtechnique. McGraw-Hill Book Co., Inc., New York.

8. Ladell, J. L.1959. A method of measuring the amount and distribution of cell wall

material in transverse microscope sections of wood. Jour. Inst.Wood Sci. No. 3:43-46.

9. Lange, P. W.1950. Optical methods for micro analysis of the plant cell wall. Svensk

Papperstidning 53:749-766.

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10. Lange, P. W.1958. The distribution of the chemical constituents throughout the cell

wall. pp. 147-185. In Fundamentals of Papermaking Fibres,F. Bolam, (ed.), Transactions of the Symposium held atCambridge, Sept. 1957. British Paper and Board Makers’Association.

11. Needham, G. H.1958. The Practical Use of the Microscope. Charles C. Thomas,

Springfield, Illinois.

12. Richards, O. W.1959. The Effective Use and Proper Care of the Microtome. American

Optical Co., Buffalo, New York.

13. Ritter, G. J.1951. Microscopic polarized light method for studying cellulose fibers.

Paper Indus. 33: 926-931.

14. Rogers, J. D., and Berbee, J. G.1964. Developmental morphology of Hypoxylon pruinatum in bark of

quaking aspen. Phytopathology 54:154-162.

15. Sass, J. E.1958. Botanical Microtechnique. 3rd ed. The Iowa State College Press,

Ames.

16. Shillaber, C. P.1944. Photomicrography in Theory and Practice. John Wiley and Sons,

Inc., New York.

17. Vaughan, R. E.1914. A method for the differential staining of fungous and host cells.

Ann. MO. Bot. Gard. 1:241-242.

18. Venning, F. D.1954. Manual of Advanced Plant Microtechnique. Wm. C. Brown CO.,

Dubuque, Iowa.

19. Wilson, J. W.1954. Fibre Technology--II. A critical survey of laboratory wood

maceration techniques. Pulp and Paper Mag. of Canada55(7):127-129.

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