PREPARATION of DECAYED WOOD for MICROSCOPICAL EXAMINATION Table of Contents

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

U. S. FOREST SERVICE RESEARCH NOTE FPL-056 August 1964

PREPARATION OF DECAYED WOOD FOR MICROSCOPICAL EXAMINATION Table of Contents

Page

Summary ...... 1

Introduction ...... 1

Preparation of Decayed Samples ...... 2

Embedding Methods ...... 2 Celloidin ...... 2 Paraffin ...... 5 Polyethylene Glycol ...... 6 Freezing...... 7

Maceration...... 8

Sectioning ...... 8

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

Staining...... 11 Differentiation of Hyphae and Wood ...... 12 Picro aniline blue ...... 12 Pianeze IIIb ...... 13 Differentiation of Wood Structures and Components ...... 13 Safranin and fast green ...... 14 AzureB ...... 15 Zinc-chlor-iodide and phloroglucinol ...... 15 Iodine-potassium iodide ...... 16

Mounting...... 17

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

Literature Cited ...... 21

FPL-056 PREPARATION OF DECAYED WOOD

FOR MICROSCOPICAL EXAMINATION

W. WAYNE WILCOX, Pathologist

Forest Products Laboratory,’ Forest Service U.S. Department of Agriculture

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Summary

This report describes some of the methods that were devised or found to be particularly satisfactory for the microscopical observation of wood in various stages of decay. These methods include the celloidin, paraffin, and polyethylene glycol embedding methods, methods for macerating, sectioning, staining, and mounting, and a discussion of several optical systems which facilitate microscopical observation of decayed wood. A rapid method for the measurement of changes in the amount of cell wall substance visible in cross sections is discussed.

Introduction

Because of its generally soft or friable nature, decayed wood may be difficult to prepare satisfactorily for microscopical examination. For example, thin sectioning often requires that an embedding matrix be used to hold the wood structure intact during the cutting process. Since samples of decayed wood may differ greatly in their hardness and strength, it is desirable to have available a 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 of changes caused by decay in the microstructure of wood, which was undertaken in the preparation of a Doctoral thesis.

1 Maintained at Madison, Wis., in cooperation with the University of Wisconsin. FPL-056 Preparation of Decayed Samples

In this study, suitable decay samples were obtained by following the major procedures of the soil-block method outlined in ASTM Standard Method D1413-61 (1).2 Blocks to be decayed were cut with a thickness of 1/8 inch along the grain, so that specimens of satisfactory size for microscopical examination could be prepared by simply splitting the blocks along the grain with a sharp knife. Reducing the thickness of the blocks increased the uniformity of decay and made further cutting across the grain unnecessary. Cutting across the grain, after the blocks had been decayed, could have produced considerable distortion. The blocks were conditioned and weighed according to the specifications of the soil-block method..

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

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

Embedding Methods

Celloidin

The celloidin method proved to be the most satisfactory procedure for embedding decayed wood. The results fully justified the required expenditure of time--nearly 2 months for the preparation of fully embedded specimens. Even the slight amount of structure still present in white-rotted wood at weight losses of over 70 percent was held intact by this method during the cutting of sections 4 microns in thickness.

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

2 Underlined numbers in parentheses refer to Literature Cited at the end of this report. FPL-056 -2- absorbed enough alcohol and sank. They were then transferred to several changes of ethylene glycol monomethyl ether (also under vacuum), which proved to be an excellent solvent for celloidin. Unlike ether-alcohol, it presents no explosion hazard and it evaporates slowly enough to allow convenient handling. The specimens in ethylene glycol monomethyl ether were placed in a stoppered bottle in an oven at 52° to 54° C. for 2 days or more. They were transferred successively through 2, 4, 6, 8, and 10 percent3 solutions of celloidin in ethylene glycol monomethyl ether maintained at 52° to 54° C., and were allowed to remain in each grade 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 face to be sectioned downward, to hardening chambers (fig. 1),4 and covered with a 12 percent solution of celloidin in ethylene glycol monomethyl ether. The celloidin in the hardening chambers was allowed to concentrate by evaporation at room temperature. Additional celloidin was added periodically in order to keep the chambers full and to allow for further concentration. During this process, the surface of the celloidin became dry and hard. The configuration of the surface crust 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 a 40 percent solution was added. The crust was removed with a knife prior to each addition to prevent the trapping of air bubbles in the matrix. The 40 percent celloidin was allowed to evaporate at room temperature until it became quite firm, or until it, began to pull the bottom of the chamber inward. At this stage there was little danger of the celloidin becoming too hard, provided the shrinkage did not begin to distort the specimen, since it was found that very hard celloidin still could be successfully sectioned merely by increasing the period of storage in glycerin-alcohol.

The embedded specimen was next removed from the hardening chamber. With a knife, the chamber was cut away from the glass slide and loosened from its contents. The embedded specimen was then removed from the chamber by pressing on one end and placed on a wooden mounting block of such size that it could be firmly held in the microtome. The mounting block had been impregnated previously 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.

3 Throughout 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 liquid reagents is expressed on a volumetric basis. 4 The idea from which these chambers were developed was originally that of Dr. Catherine Duncan of the Forest Products Laboratory.

FPL-056 -3- Figure 1.--Celloidin hardening chambers. These consisted of two 1/2-inch-long segments of plastic tubing 1/2 inch in outside diameter adhered to a 1- by 3-inch glass microscope slide with a room-temperature-curing nitrile-phenolic resin in organic solvents. The use of a glass slide with one etched end allowed the contents of the chambers to be labeled with a pencil. Such chambers facilitated the handling of large numbers of specimens and allowed the specimens to be oriented for sectioning at a stage in the embedding process when they could be easily observed.

M 126 011

FPL-056 -4- After removal from the chloroform, the hardened material was trimmed down for sectioning and stored in a 50-50 mixture of glycerin and 95 percent ethanol. The longer the material was stored in glycerin-alcohol, the better were its cutting properties. Sectioning was performed on a sliding microtome.

Paraffin

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

The methods of Sass (15) for paraffin embedding, which employ tertiary butyl alcohol (TBA), proved satisfactory for application to decayed wood. Only minor modifications suggested by Rogers (14) were utilized. The material to be embedded was gently split into small blocks approximately 1/8-inch square and 1/8 inch to 3/16 inch in length along the grain. The specimens were dehydrated or moistened, depending upon the beginning moisture content, by means of an ethanol series (15), to the stage of 60 percent (by volume) ethanol. They were evacuated until they sank in the first liquid used in the series. The samples were then transferred through the TBA series of Sass (15) (table 1).

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

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.

1 Reprinted 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 were stored in each grade for 4 hours or more. From grade 5 they were transferred to several changes of anhydrous TBA and then to a 50-50 mixture of anhydrous TBA and paraffin oil, using the same evacuation and storage schedule as with grades 1 through 4. The bulk of the TBA-paraffin oil then was decanted into a

FPL-056 -5- waste container. The rest, along with the specimens, was gently poured into a vial on top of molten embedding paraffin which had been cooled sufficiently so that 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 for tissue embedding. The vial containing the specimens was then stored in an oven hot enough to keep the paraffin completely fluid. Upon the melting of the surface film, the specimens dropped into the pure paraffin. Periodically the paraffin was decanted into a waste container and replaced with fresh, molten paraffin. The paraffin was changed four or five times, or until the odor of TBA was no longer detectable. The specimens, along with the final change of paraffin, were poured into a paper boat placed upon a casting table-- a heavy steel plate heated at one end with a small flame (15). The boat was placed close enough to the flame to allow the paraffin to remain molten until the specimens had been arranged. By sliding the boat away from the flame, a thin film of solidifying paraffin was formed at the bottom of the boat, which aided in maintaining the specimens in the desired positions. As soon as the paraffin had solidified enough to hold the specimens firmly, the boat was placed in a freezing compartment to quickly and thoroughly harden the paraffin. When the paraffin had become hard, the paper boat was removed, and blocks of paraffin containing individual specimens were cut out with a hacksaw blade. Then the specimens were affixed to mounting blocks previously impregnated with molten paraffin, by melting portions of the paraffin with a hot needle or scalpel (15).

Polyethylene Glycol An alternative embedding method for use with the rotary microtome, employing polyethylene glycol (PEG), was tried on a limited number of specimens. This method was of interest because of certain reported advantages: (1) green or moist samples may be used directly, reducing the problem of distortion due to dehydration, and (2) the method is rapid and requires a minimum of specimen handling. The limited utilization of PEG embedding in the thesis study was insuf- ficient for a thorough evaluation of the method, but the evidence obtained showed distinct promise for application to relatively soft or decayed wood. Also, it appeared that it might be especially useful for studying fungal hyphae in wood since, in the absence of severe dehydration, it should minimize distortion and collapse of the hyphae. The method utilized was taken without modification from Gjovik.5 He found commercial PEG with a molecular weight of 1450 most suitable for histological application. However, the ability to form good ribbons varied considerably from 5 Gjovik, 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. FPL-056 -6- batch to batch, and it was suggested by Gjovik that samples be tested for this property before a batch is selected for histological use. Prior to embedding, the specimens were thoroughly water saturated under vacuum. They were then placed in a 50-50 mixture of PEG-1450 and distilled water at 50° C. for 2 hours. Next the 50-50 mixture was decanted, replaced with melted, pure PEG-1450, and stored at 50° C. for 2 hours. The old PEG-1450 was then exchanged for fresh liquid and stored at 50° C. for 4 to 6 hours. Although additional evacuation while in pure PEG was recommended by Gjovik, this procedure probably was necessary only if air had been incompletely removed initially. Individual specimens were mounted by placing them in the cavity formed by wrapping a wooden mounting block with masking tape so that it extended beyond the end of the block However, a number of specimens may be prepared at one time by the use of paper boats as in the paraffin technique. The PEG was then hardened by placing the mounted specimens, or the paper boats, in a refrigerator for 30 to 40 minutes until the PEG exhibited a grayish-white translucence. The embedded and mounted specimens were then trimmed for sectioning and stored in a desiccator containing calcium chloride. Gjovik found that special techniques, such as cooling of the microtome knife or softening or cooling of the embedded specimen, were unnecessary for satisfactory sectioning.5 He also found that acceptable sections could be obtained in less than 1 hour by using an abbreviated schedule consisting of the same steps as outlined above but allowing only 15 to 20 minutes of storage at each stage. 5 Gjovik suggested a potassium dichromate-gelatin adhesive, instead of Haupt’s adhesive, for use with sections of PEG-embedded material. This avoided the use of 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 this study 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 was lost when the ice melted soon after the section had been cut. Freezing of the specimens impregnated and surrounded with water can be accomplished in several ways (15). In trials on decayed wood a carbon dioxide freezing apparatus was used. Some additional support was obtained during cutting by deflecting the jet of carbon dioxide so that it cooled the knife in addition to the specimen. A special attachment is commercially available for this procedure. The specimen to be sectioned was placed under vacuum in water until it sank. The specimen was placed upon the freezing attachment and frozen; water was dripped on it so that it became thoroughly surrounded by ice.

FPL-056 -7- Periodic reapplication of carbon dioxide was necessary during sectioning to maintain the specimen in the frozen state. The specimens were sectioned on a sliding microtome, and the sections were handled with a moist brush in the same manner as sections from nonembedded specimens. It has been reported that additional support may be obtained if, instead of pure water, an aqueous solution of a substance such as gelatin or agar is utilized (7,15).

Maceration

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

Several methods for the maceration of wood are available (5,19), but Jeffrey’s method (7,15,19) is simple and proved entirely satisfactory in this study. The best results were obtained when the wood to be macerated was gently split into slivers about 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 macerating fluid consists of equal amounts of 10 percent (by volume) aqueous nitric acid and 10 percent aqueous chromic acid. Small glass beads were added to the vials containing the macerating fluid and the wood samples, and the vials were shaken to aid separation of the elements. If the wood did not subdivide easily at this stage, it was allowed to settle, and the darkened macerating fluid was drawn off and replaced with fresh fluid. The samples in fresh fluid were stored again in the oven overnight.

This procedure was repeated until satisfactory maceration was achieved. The samples were then washed with water using a low-speed centrifuge to settle them between washings. Each macerated sample was stored in 50 to 70 percent ethanol. For microscopical examination, the sample was observed under a sealed cover glass, 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 of crushing and distortion; therefore, the sliding microtome was utilized except for

FPL-056 -8- the limited observations on specimens embedded in paraffin or polyethylene glycol. 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 of the specimen. The use of a razor blade in place of the microtome knife was found to be very convenient and highly satisfactory, since the blade could be discarded when dull or nicked. Further details of rotary microtomy are discussed by Sass (15).

The sliding microtome was used for nonembedded and for celloidin-embedded specimens. A conventional, wedge-shaped microtome knife was used with this instrument. The slice angle, the angle the knife makes with its direction of travel, was kept as small as possible, making fullest use of the slicing action of the 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. The bottom surface of the bevel was not allowed to be completely parallel to the specimen’s surface and the back of the bevel was never placed below the cutting edge. 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 when cutting sections of the thinness required in this study. Difficulty in sectioning with the sliding microtome usually could be attributed to some of the following factors. The most common cause of substandard sections was a dull or imperfect knife edge. Dullness of the knife edge was indicated by disintegration of the sections during cutting or variations in thickness from one section to the next. Coarse nicks in the blade edge usually resulted in the division of the section into several pieces. Fine nicks were hard to detect but often were the cause of weak sections which fragmented when touched by the brush. Such fine nicks usually caused small scratches in the specimen, detectable only when the specimen surface was wiped dry. These difficulties were overcome by restropping or in severe cases rehoning the knife edge. However, if defects appeared only in localized areas of the knife, other regions could be used for sectioning. Occasionally tearing of the sections occurred when the knife was in perfect condition. It was found that such tearing could be overcome by diluting the glycerin-alcohol, used for lubrication with additional 95 percent ethanol, by applying less pressure to the brush while guiding the sections, or by decreasing the slice angle of the knife.

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

FPL-056 -9- steam. In order to avoid tearing and distorting the section, it was touched with a soft-bristled brush during the cutting and held lightly in the approximate position it occupied while a part of the specimen. A brush with bristles mounted in plastic rather 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 all were lubricated by frequently filling the brush with the liquid so that the section literally floated off of the specimen during cutting. The liquid used was water or dilute 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 in celloidin. 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 cellulose tape to hang down over the specimen mounted in the sliding microtome. The tape is firmly affixed to the thoroughly dried surface of the specimen, and the section is cut with a slight tension applied through the tape to its leading edge, to keep the tape from sticking to the knife. Sections made by this method were of high quality as long as they were left on the tape and viewed directly through it. However, it was not possible to satisfactorily remove the tape and adhesive from the sections. Soaking in acetone was helpful, but this process resulted in curled, brittle sections which were difficult to handle.

In general, thin sections produced sharper microscopical images than did thick sections; therefore, in this study, celloidin-embedded specimens were sectioned at a thickness of 4 microns. Thinner sections were easily obtained with such material but proved extremely difficult to handle because they were not easy to see. Since it was difficult to obtain sections as thin as 4 microns from specimens which were not embedded in celloidin, paraffin- and polethylene glycol-embedded specimens were sectioned at thicknesses of 8 to 12 microns and nonembedded specimens at thicknesses of 10 to 20 microns. Such thick sections were necessary for critical observation of irregular structures such as hyphae. The thick sections also 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 stages of decay, it was found most convenient to adhere the sections to the slide prior to

FPL-056 -10- staining. For attaching the sections, Haupt’s adhesive proved satisfactory (7). This adhesive was prepared by dissolving 1 gram of purified, finely divided gelatin 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. The use 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 clean glass slide and spread with the finger, leaving a barely perceptible layer. Several drops of 4 percent (by volume) aqueous formaldehyde were placed on this surface and the sections, which previously had been washed in water to remove most of the glycerin or alcohol, were added and arranged. The slide was then placed upon a warming table or in an oven at 40° to 50° C.

Gjovik 5 suggested the use of a potassium dichromate-gelatin adhesive with polyethylene glycol-embedded specimens. This adhesive consisted of an aqueous solution of 0.5 percent potassium dichromate and 0.5 percent gelatin, with the excess potassium dichromate removed by dialysis. For use, a light film of the potassium dichromate-gelatin adhesive was spread on the slide. The sections were carefully placed upon this moist surface, and the slide was dried upon a warming table 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 be observed, it was found desirable to hold these in place by allowing some of the embedding 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, to improve image clarity.

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

Staining

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

FPL-056 -ll- more general methods are discussed here. The Color Index numbers of the stains 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 differentiation of hyphae; it was also used successfully to reveal bacteria in wood.6 Originally developed by Cartwright (3), it is presented in this report in somewhat modified form. With this procedure, wood cell walls appear pink, and hyphae or bacterial cells 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), plus 100 milliliters saturated, aqueous picric acid.

Staining Schedule

1. Stain in safranin--2 minutes. Place sections in 10 milliliters of distilled water and add 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 add 5 drops of the picro aniline blue stock solution. Heat on a medium-warm hot plate until the stain begins to steam slightly.

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 produce permanent mounts (discussed in the section on “Mounting”).

6 Knuth, 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.

FPL-056 -12- Pianeze IIIb.--This is an alternative method of differentially staining fungal hyphae in wood and has the advantage of requiring only a single stain application. However, the differentiation proved to be less striking than that of the picro aniline blue method in some material. The following procedure is an adaptation of Vaughan’s method (17), used extensively for routine examinations at the Forest Products Laboratory.

Stock Solution

Pianeze IIIb: malachite green (C.I. 42000) 1.0 g. acid fuchsin (C.I. 42685) 0.5 g. martius yellow (C.I. 10315) 0.05 g. distilled water 150.0 ml. 95 percent ethanol 50.0 ml.

Staining Schedule

1. Wash in water.

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

3. Wash in water.

4. Decolorize in acid alcohol (95 percent ethanol, to which a few drops of concentrated hydrocholoric acid have been added).

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 the examination of plant materials, including wood. Several of these common methods

FPL-056 -13- were also found in this study to greatly aid in the detection of changes in wood structure and composition resulting from decay.

Safranin and fast preen. --This is one of the commonest staining procedures applied to wood and other plant material. It was found to be very useful because it was effective and nearly foolproof in application. The only critical length of time in the schedule was the period in which the sections were in contact with fast green, By this method heavily lignified portions of the cell wall stain red and less lignified portions (the secondary walls of many woods) green. Even in advanced stages of decay, this differentiation was maintained. Although these stains are not specific for lignin or cellulose (6), the staining reactions obtained with them correlated well with data from more specific methods and with the expected chemical composition of the decayed wood. Numerous modifications of this staining procedure have appeared, but the schedule described below provided the author with fairly uniform results on a variety of materials. Stock Solutions safranin: 1 percent solution of safranin 0 (C.I. 50240) in 50 percent ethanol; filtered. fast green: 1 percent solution of fast green (C.I. 42053) in clove oil-alcohol (1 part clove oil, 9 parts 95 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 of clove oil for a minimum of 5 minutes.

8. Flush with xylene.

9. Mount.

FPL-056 -14- Azure B. --According to Jensen (6) this method is fairly specific for lignin and for nucleic acids. All tissues which were stained red by the safranin and fast green technique were found to stain blue-green with azure B, and as with safranin the staining properties were retained into advanced stages of decay. A full discussion of this staining method is offered by Jensen (6), while only the staining schedule is outlined below.

Stock Solutions

azure B: 0.25 milligrams azure B (C.I. 52010) per milliliter of citrate buffer at pH 4.0. citrate buffer (pH 4.0): 2.7 grams citric acid and 2.1 grams sodium citrate dissolved in 100 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 suggested for determining the presence of cellulose and phloroglucinol for the presence of lignin. Both methods are discussed by Jensen (6). Apparently neither technique is specific for the component it is designed to indicate. Furthermore, it was found in this study that the residual lignin in heavily decayed wood failed to react with phloroglucinol. This information may be of importance as an indication of subtle changes occurring in the lignin molecule as a result of decay, but it also renders the technique less useful as a stain for general application to decayed wood. Due to swelling of the cellulose, zinc-chlor-iodide had the disadvantage of distorting the section.

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

FPL-056 -15- 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 in combination.

Stock Solutions

zinc-chlor-iodide: according to Venning(18), dissolve 15 grams zinc chloride in 10 milliliters of distilled water, then add 0.15 grams potassium iodide and 0.25 grams iodine. The iodine is in excess to 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 sections appear black.

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

4. Cover and observe as a wet mount.

Iodine-potassium iodide.--This method proved useful for observing the location of starch in both sound and decayed wood, but it also is not specific for this component (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 in 100 milliliters distilled water. Then dissolve 0.2 grams iodine in the potassium iodide solution.

FPL-056 -16- Staining Schedule

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

Mounting

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

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

For permanent mounting, a synthetic resin mounting fluid was used. This resin was 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 Small amount was needed, and it hardened in about 30 minutes at room temperature. In preparation for permanent mounting, the sections were dehydrated by means of an alcohol series, cleared for 5 minutes or more in clove oil, and washed with xylene. Excess xylene was blotted away, but enough was left so that the sections were 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 lowered to 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 of the cover glass and waiting momentarily. The raised end of the cover glass was then slowly lowered to the slide. Excess xylene and resin were removed by blotting the edges of the cover glass. Troublesome bubbling could be avoided by using more xylene on the sections or by thinning the mounting medium. If bubbles formed, they usually disappeared during the process of removing the excess xylene. A small brass or lead weight was placed on top of the cover glass for about 30 minutes to 1 hour at room temperature, after which the slide was ready for use.

FPL-056 -17- Microscopical Methods

Observational

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

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

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

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

Absorption of specific wavelengths of ultraviolet radiation was utilized to follow lignin distribution in sections of sound and decayed wood. When irradiated with wavelengths of about 2800Å lignin appeared dark, due to intense absorption of the radiation, while the carbohydrates appeared light. This method is discussed by Lange (9,10).

Measurement

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

FPL-056 -18- methods for determining cross-sectional area, utilizing ocular micrometry, planimetry, or photographic weighing, were considered too time consuming to provide the necessary replication. Instead, a method developed by Ladell (8) for measuring 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 of cells to be measured.

The method consisted of projecting the image of a section from the microscope onto a white card containing 100 pinholes punched on grid coordinates determined from a table of random numbers. The card containing the randomly spaced pinholes was illuminated from below, so that the holes appeared as spots of light on 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 the percentage of the area occupied by the structure.

FPL-056 -19- Figure 2. --Adaptation of alight microscope for measurement of cross-sectional area on microtome sections. (A) prism; (B) mirror: (C) opaque sampling card with sampling field (outlined by black line) containing 100 randomly spaced pinholes; (D) light-tight illuminating box containtng a 15-watt bulb and aluminum-foil reflector; (E) light shield; (F) hand counter. This apparatus was adapted from a method developed by Ladell.

M 126 765

FPL-056 -20- 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 ASTM Standards, 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.

FPL-056 -21- 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 at Cambridge, 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 Canada 55(7):127-129.

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