Microscopic Preparation Techniques for Plant Stem Analysis

Abstract

You can order the book by sending an email to the printing house.
(Price: 22.- Euro plus shipping) This booklet results from the experiences of teaching “Wood Anatomy of
Tree Rings” courses, during which we realized that there is a deficit regarding
available information on the practical aspect of wood anatomical work.
We aim to present basic preparation techniques in wood anatomy, in an understandable
way and supported by many photographs and illustrations. This
booklet is intended for experts and beginners in wood anatomy that come
from various fields of research such as dendrochronology, ecology, archeology,
geography, forestry and botany.
We provide instructions on how to produce permanent, high-quality micro
sections, using techniques which are simple to apply. The methods proposed
are suitable for analyzing living and dead wood as well as bark, from herbs,
shrubs and trees, growing in wet to dry environments. Special attention is
given to the use of collecting, polishing and sectioning tools for observations
on sample surfaces and micro slides. Various ecologically significant and wellestablished
staining methods are demonstrated to highlight cell contents and
cell walls from very soft to very dense and even artificial material such as plywood.
Furthermore, bleaching and staining processes are described to allow
analysis of fibers and fungal hyphae in decomposed wood.
By presenting many practical tips and tricks (e.g. using cheap disposal blades,
unrolling sections, avoiding cell-wall destruction, sectioning very soft material
and charcoal, and preparation errors), we intend to support scientists and
reduce frustration that may occur during laboratory work.
This booklet does not intend to present all existing microscopic preparation
techniques, but instead intends to provide advice on the most common techniques.
We encourage readers to apply the techniques presented herein, and
also to further develop the techniques based on their experiences and results.

Order the book:
You can order the book from the publisher Kessel Publishing House
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by sending an email to: nkessel(at)web.de indicating your full postal address

Full text

Holger Gärtner
Fritz H. Schweingruber
Design & layout Anne Börner
Microscopic
Preparaon Techniques
for Plant Stem Analysis

Dr. Holger Gärtner
Prof. Fritz H. Schweingruber
Swiss Federal Research Institute WSL
Landscape Dynamics / Dendroecology
Zürcherstrasse 111
8903 Birmensdorf
Switzerland
E-Mail: holger.gaertner@wsl.ch
Design & layout: Annett Börner, Adelaide
www.dn.com.au/annett-boerner.html
Originalausgabe
© 2013 Verlag Dr. Kessel
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ISBN: 978-3-941300-76-7

1
Contents
Introduction ............................................................................................................... 3
1. Sampling material and sampling strategies ...................................................... 4
1.1 Sampling in various biomes ........................................................................... 4
1.2 Sampling di erent life- and growth forms .................................................... 6
1.3 Sampling di erent parts of plants ................................................................. 8
1.4 Sampling dead wood .....................................................................................11
2. Collection and preservation ............................................................................... 13
2.1 Tools ...............................................................................................................13
2.1.1 The importance of sharpening increment borers ...............................15
2.2 Preserving samples for structural analysis ...................................................16
2.3 Labeling collected samples ........................................................................... 17
3. Sectioning and maceration ................................................................................18
3.1 Stabilization of the material..........................................................................18
3.2 Preparing stem disks for sectioning: sawing, splitting, and boiling ...........19
3.3 Sectioning by hand ........................................................................................ 21
3.4 Splitting and sectioning charcoal or mineralized wood ..............................22
3.5 Preparation of surfaces for macroscopic observation ................................25
3.5.1 Making annual rings visible by cutting and sanding ........................... 26
3.5.2 Observation surfaces by enhancing contrast ..................................... 28
3.6 Sectioning with microtomes ........................................................................ 30
3.6.1 Microtome knifes .................................................................................. 31
3.6.2 Microtome types ...................................................................................32
3.6.2.1 WSL-Core-microtome ..................................................................34
3.6.2.2 Microtome type GLS1 ..................................................................35
3.6.2.3 WSL-Lab-microtome (modied Reichert-type) ..........................37
3.6.3 Sectioning with microtomes and observation without staining ....... 39
3.6.3.1 Material of normal stiness ....................................................... 39
3.6.3.2 Sectioning conifers .....................................................................41
3.6.3.3 Sectioning of very soft material ................................................42
3.6.3.4 Stabilization and sectioning of waterlogged archeological wet
wood ...........................................................................................42
3.6.3.5 Observation of soft herb stems ................................................. 44
3.7 Storing glycerol-preserved sections ........................................................... 45
3.8 Maceration and measuring axial cell dimensions ......................................46
3.9 Preparation of textile fiber ......................................................................... 48
3.10 Preparation of technically composed wood .............................................49

2
4. Fixation, bleaching, staining, embedding, cleaning, labeling and storing .....52
4.1 Effec of staining, and safety instructions ....................................................52
4.2 Fixation of cell contents ............................................................................... 54
4.3 Bleaching: Destruction of cell contents and dark-stained cell wall
components .................................................................................................. 55
4.4 Staining cell walls ......................................................................................... 56
4.5 Staining cell contents ................................................................................... 58
4.5.1 Nuclei and protoplasts ......................................................................... 58
4.5.2 Starch grains ........................................................................................ 59
4.6 Staining cell walls and cell contents ............................................................60
4.7 Staining hyphae, bacteria and decomposed cell walls ................................61
4.8 Dehydration .................................................................................................. 62
4.9 Embedding micro sections ........................................................................... 63
4.10 Drying and stabilizing ................................................................................. 65
4.11 Removing from plastic strips and cleaning slides .....................................66
4.12 Labeling ....................................................................................................... 67
4.13 Operating a line for staining, dehydrating and embedding ...................... 67
4.14 Repairing cracked (embedded) slides .......................................................68
4.15 Final storage ................................................................................................68
4.16 Digital classificatio of slides...................................................................... 70
4.17 Processing time ........................................................................................... 70
5. Preparation errors .............................................................................................. 71
6. Microscopic observation ................................................................................... 74
6.1 Polarized light ............................................................................................... 74
6.2 Measuring cell dimensions ........................................................................... 76
6.3 Photography ................................................................................................. 76
References ................................................................................................................77
Acknowledgements ................................................................................................. 78

3
Introducon
If we had written this booklet 20 years ago, the title would have been different
“Microscopic preparation techniques for wood analysis”. However, in the last years
it became obvious that there is a need in dendrochronology, as well as in the sci-
ence of plant anatomy (Schweingruber et al. 2011, 2012), to study anatomical stem
structures in more detail. Many textbooks describe preparation techniques useful
in all field of biology and botany, e.g. Kremer (2002), Mulisch & Welsch (2010), Pur-
vis et al. (1964), Rudzin (1999); or special techniques for wood anatomy e.g. Chaffe
(2002), Ives (2001), Hoadley (1990). However, not a single text concentrates speci -
cally on xylem and phloem of plant stems, branches and roots.
Based on our experience in our laboratories (Swiss Federal Research Institute
WSL, Birmensdorf, Switzerland), and during more than 20 international anatomical
courses with participants from all over the world, we have concluded that a “cook
book” for simple microscopic plant stem analysis is useful for beginners. The main
goal of this booklet is to provide instructions for producing high quality micro sec-
tions using simple techniques in an efficie way. Therefore we focus on processes
that do not demand embedding plant tissues before sectioning. The main mes-
sage of this booklet is how to use traditional and new microtomes with disposable
cutter blades and simultaneous multiple staining techniques. Having worked with
students and experienced scientists with dozens of differen mother tongues and
sometimes with limited knowledge of the English language, we have come to the
conclusion that pictures can tell better technical stories, rather than long-written
instructions.
In the last few years we have learned that new sliding microtomes, new cutting
techniques and better staining methods allowed also the preparation of bark
and very small soft stems. Since we recognized that secondary stem thickening
occurs in all conifers and most dicotyledonous plants and that lignificatio occurs
in most terrestrial plants we overcome the boundaries between herb and tree, and
between woody and herbaceous plants. Therefore, we analyze the xylem, phloem,
cortex and phellem of plant stems in general.

4
1. Sampling material
and sampling strategies
Wood and bark in stems, branches and roots of trees, shrubs and herbs contain
information about their genetic origin, the formation time and environmental con-
ditions.
For all tree-ring related studies, the sampling strategy is the most important part
when initiating a project. ‘Tree’-ring related and wood anatomical studies are not
limited to trees, and not even to areas where trees are able to grow. Beyond the
limits of tree growth other woody plants such as shrubs, dwarf shrubs or even
herbs can be found, also forming annual rings. Therefore, these organisms can
potentially be used as proxies for reconstructing past environmental conditions.
Furthermore, research strategies and thus sampling strategies are not restricted to
the stem of the woody plants. Whether one is working with trees, shrubs, dwarf
shrubs or herbs, all parts of these plants (roots, stems and branches, or even bark)
potentially carry environmental information, which can be analyzed in detail. This
fact has to be respected before thinking about possible sampling strategies, inde-
pendent of whether studying macro- or microscopically the annual rings of woody
plants.
1.1 Sampling in various biomes
All terrestrial and lacustrine biomes contain important plant material, which can
be analyzed in relation to taxonomy, morphology and environmental conditions
(Walter & Breckle 1989) (Figs. 1.1–1.6).

5
Fig. 1.1. High Arctic: mainly dwarf shrubs and long
living perennial herbs (hemicryptophytes) grow-
ing under dry and moist conditions as well on
bolder fields on permafrost. Mosses, sedges and
grasses dominate wet plains and lakeshores.
Fig. 1.2. Hot desert: shrubs, dwarf shrubs and
hemicryptophytes are scattered on sand dunes
and rock fields. Annual plants germinate and
exist only during rainy seasons.
Fig. 1.3. Boreal forest: conifers, dwarf shrubs and
mosses dominate circumpolar boreal forests on
all sites.
Fig. 1.4. Forests in temperate zones: trees, shrubs,
and hemicryptophytes dominate most sites.
Fig. 1.5. Tropical forests: trees and lianas domi-
nate tropical rain forests.
Fig. 1.6. Wet sites: mainly sedges, grasses and
hydrophytes grow in, and on, the border of seas,
lakes and ponds.

6
1.2 Sampling dierent life- and growth forms
All terrestrial and lacustrine plant associations contain important plant material,
which can be analyzed to reconstruct environmental conditions. Consequently,
trees, shrubs, dwarf shrubs, palms, lianas, succulents, annual and perennial herbs,
grasses, mosses, lichens and algae can be of interest for dendroecological and/or
wood anatomical studies (Figs. 1.7–1.20).
Fig. 1.12. Dwarf shrub, prostrate
Fig. 1.7. Tree, conifer
Fig. 1.10. LianaFig. 1.9. Palm
Fig. 1.8. Trees, dierent species and growth forms
Fig. 1.11. Succulent

7
Fig. 1.13. Dwarf shrub, parasite
on tree Fig. 1.15. Monocotyledonous per-
ennial plant (hemicryptophyte)
Fig. 1.14. Dicotyledonous annual
plant (therophyte)
Fig. 1.20. Algae
Fig. 1.19. Lichen
Fig. 1.18. MossFig. 1.17. FernFig. 1.16. Water plant (hydro-
phyte)

8
1.3 Sampling dierent parts of plants
For anatomical analyses, the xylem and phloem of stems, branches, roots (root
collar), rhizomes of dicots and monocots, needles, leaves and below-ground stems
can be used (Figs. 1.21–1.39).
Fig. 1.21. Cross section of a deciduous tree: oak
stem (Quercus petraea) with heartwood (1) and
sapwood (2).
Fig. 1.22. Cross section of a conifer: spruce (Picea
abies), eccentric stem with compression wood
(1).
Fig. 1.25. Bark: coniferous tree, Mountain pine
(Pinus montana).
Fig. 1.26. Bark: deciduous tree, birch (Betula pen-
dula).
Fig. 1.23. Cross section of a liana (stem): alpine vir-
gin’s bower (Clematis alpina), assimilating xylem
with very large dilated rays (1).
Fig. 1.24. Cross section of a succulent (stem):
Giant cereus (Cereus giganteus), with small but
not continuous xylem.
1
2
1
1

9
Fig. 1.27. Exposed roots of a deciduous tree (Fraxi-
nus excelsior).
Fig. 1.28. Exposed roots of a conifer (Picea abies).
Fig. 1.29. Stem of a dicotyledonous tree (Convol-
vulus arborea) with successive cambia.
Fig. 1.30. Stem of a palm with single vascular
bundles.
Fig. 1.31. Stem of a tree-fern (Cyathea arborea)
with elongated closed vascular bundles. Fig. 1.32. Stem base of a herbaceous fern (Stru-
thiopteris onoclea), central stem and peripheral
leaf-bases.

10
Fig. 1.39. Rhizome of a monocotyledonous plant
(Festuca sp.).
Fig. 1.33. Long shoots of a coni-
fer (Pinus montana).
Fig. 1.34. Long shoot of a decid-
uous tree (Betula pendula).
Fig. 1.35. Short shoots of a coni-
fer (Larix decidua).
Fig. 1.36. Seedlings of a deciduous tree (Castanea
sativa).
Fig. 1.37. Polar root of an annual plant (Plantago
maritime).
Fig. 1.38. Rhizome of a dicotyledonous plant
(Viola riviniana).

11
1.4 Sampling dead wood
The xylem of dead material can contain useful information when analyzed micro-
scopically, i.e., well preserved wooden constructions and artifacts, charcoal,
archeological wet wood, petrifie wood, stems infected by fungi, stems with scars
(Figs. 1.40–1.48).
Fig. 1.40. Posts in the Lagoon of Venice Fig. 1.41. Artifact, violin
Fig. 1.42. Wood decomposed by insects Fig. 1.43. Wood decomposed by fungi
Fig. 1.44. Driftwood

12
Fig. 1.45. Charcoal
Fig. 1.47. Petried wood Fig. 1.48. Mineralized wood on a sword
Fig. 1.46. Degraded wet wood from a Neolithic
lake dwelling in Switzerland

13
Fig. 2.1. Garden pick
Fig. 2.3. Lopping shear and handsaw
Fig. 2.2. Pocket knife and paper knife (NT)
Fig. 2.4. Chain saw
2. Collecon and preservaon
2.1 Tools
Herbs can be excavated using common garden tools and parts of plants can be cut
with knives or lopping shears. Twigs and small stems can be cut with scissors (Figs.
2.1–2.3). Dead wood (logs, driftwood, etc.) can be cut with handsaws and/or chain
saws (Figs. 2.3 and 2.4). For stems of living trees collect samples with well-sharp-
ened punchers (Fig. 2.6) and/or increment corers with diameters of 5 mm or 10 mm
(Figs. 2.5 and 2.7). Important Note: Only very sharp borers and punchers yield useful
samples for microscopic slides (Figs. 2.8 and 2.9). Samples taken with dull borers
are mechanically stressed, frequently irregularly split and therefore difficu to use
for preparing micro sections (Fig. 2.10).

14
Fig. 2.6. Puncher for micro-cores (Type: Trephor;
Rossi et al. 2006)
Fig. 2.7. Borer heads (threaded auger) of incre-
ment borers. The front part (cutting edge) needs
to be sharpened frequently.
Fig. 2.9. Sharpening borers by hand. The inner
part of the cutting edge needs to be sharpened
using a conical sandstone.
Fig. 2.8. Sharpening borers by hand. The cutting
edge is sharpened using a quadrangular oil-sand-
stone.
Fig. 2.5. Increment borer with handles and extractors (Pressler Bohrer) with di erent diameters (left
one with 10 mm, right with 5 mm and dierent length).

15
Fig. 2.10. Tangential cracks on a sectioned sample due to dull borer tip.
The need to sharpen increment borers is frequently discussed (Bauck & Brown
1955, Jozsa 1988, Grissino-Mayer 2003). Nevertheless, most borers used in the fiel
are not properly sharpened and rarely anyone takes sharpening tools out to the
fiel to sharpen the borer as soon as the extracted cores are no longer absolutely
smooth.
The use of a dull borer results in cores showing uneven surfaces due to compres-
sion forces exerted to the core while turning the bit. This is caused by a blunt cut-
ting edge not really cutting into the wood, but being pressed in while turning. For
common ring-width measurements, these compressed cores can be prepared and
analyzed without problems. In some cases the cores need to be broken because
they are twisted. If cores are twisted, this causes problems for density measure-
ments, where the angle of the tracheids needs to be absolutely upright.
In wood anatomy, increment cores had been of no interest for a long time. This
changed as soon as the research topics turned in the direction of an ecologically-
based wood anatomy and related time-series analyses. In recent years, we devel-
oped special holders for increment cores (see section 3.6 – Sectioning with micro-
tomes) enabling to cut micro sections from core pieces up to a length of 6 cm.
For these special purposes, compressed cores cannot be used!
If the cutting edge of the corer is not sharp, the cutting edge is pressed into the
wood and the core is at least partly squeezed and twisted every time the borer is
turned. These compression forces cause microscopic cracks within the structure of
the annual rings, most frequently along the weakest area, the ring boundaries and/
or the rays. As a result, the micro sections fall apart (Fig. 2.10) and it is impossible to
prepare a continuous section.
2.1.1 The importance of sharpening increment borers
500 µm

16
2.2 Preserving samples for structural analysis
When sampling herbs or bark for microscopic preparation, only fresh material is
suitable for high quality slides. After sampling, put the plant, or parts of it, in a re-
sealable plastic bag and add a few drops of 40% ethanol. Use very soft pencils for
labeling the bags because ethanol dissolves pen ink and ink of permanent markers
(Fig. 2.11). Plastic is permeable to ethanol, therefore store bags in closable plas-
tic boxes (e.g., Tupperware: Figs. 2.12 and 2.13). If the boxes are perfectly closed,
the material will not dry out for at least one year. Important Note: if you need to
analyze starch grains, the material should only be preserved in water; ethanol will
decompose starch grains.
Fig. 2.11. Re-sealable plastic bag labeled with pencil, containing samples and some drops of ethanol.
Fig. 2.12. Plastic box with bagged plant material. Fig. 2.13. Closed plastic box (airtight) ready for
storage of transportation containing ethanol
soaked plant material.

17
2.3 Labeling collected samples
All scientifi results depend on careful labeling. Ecological and dendroclimatologi-
cal analysis demand more than just a species name. The site characteristics pre-
sented in the example below, are of importance. Additional information might be
useful, but details are always depending on the aim of your study. If you intend to
collect much material, print them on self-adhesive labels (Fig. 2.14)
Fig. 2.14. Two di erent types of labels. The upper one is computer generated (advantage: data addi-
tionally stored in the computer), the lower one is lled in by hand.
Example label
Species: Fagus sylvatica L.
Life form: Tree
Collected part of plant: increment core of the stem at breast height down slope
Plant height: 25 m (height includes flower stalks for herbs)
Short site description: e.g., exposition, hydrological conditions, influence of wind, light
conditions (shadow, competition, browsing)
Geographic location: Birmensdorf, Kanton Zürich, Switzerland
Altitude above sea level: 460 m
Coordinates: At least in degrees. Navigation tools (GPS) indicates them
much more precise.
Collection date: Day/Month/Year; season or however you think date should be
recorded
Name of collector: …
Notice about photographs: …

18
3. Seconing and maceraon
3.1 Stabilizaon of the material
Fixing the sample tightly in the microtome is a basic requirement for cutting micro
sections. The sample has to be “squeezed” between the clamps of the holder. This
can be a problem when working with soft material and hollow stems. Soft material
cannot be fixe in the holder without being compressed and deformed, so it needs
stabilization. The most suitable material for stabilizing soft tissue in the microtome
sample holder is homogeneous cork. By using, for example, a cutter knife you can
cut the cork to any form and size needed (Figs. 3.1–3.4). Also recommended, but
normally less suited, are carrots, the pith of elder stems or common plastic foam
(Styrofoam) (Fig. 3.5).
Fig. 3.1. Cutting cork with a cut-
ter knife.
Fig. 3.5. Examples of materials that can be used
to fix soft material: carrot, cork, elder pith (often
too soft for hollow stems), plastic foam of vari-
ous consistencies.
Fig. 3.4. Cork with opposite grooves with an
inserted plant stem, ready for placing in a micro-
tome clamp.
Fig. 3.3. Cork with a flower stalk
in the groove.
Fig. 3.2. Cutting a groove accord-
ing to the diameter of the object.

19
3.2 Preparing stem disks for seconing:
sawing, spling, and boiling
Parts of wooden stems have to be modifie into small blocks, which are suitable
to be clamped in microtome holders. It is important that the sidewalls are parallel
to each other, if your sample is wedge-shaped it will not be stable enough in the
clamp. Important Note: If your sample is not very stable in the microtome holder,
your cuts will not be of good quality because the sample is somehow displaced
while cutting, resulting in unevenly shaped thin sections. In extreme cases, the
sample will even become stuck in the blade.
When cutting micro sections, the orientation of the sample, and therefore the cut-
ting direction (Fig. 3.6) is of importance. Especially when focusing on the growth
development of the plant, a transversal (= cross) section is needed. This direction is
the one most frequently used in tree-ring related studies.
Sections across the stems have to be sawed (Fig. 3.7). Make sure that the cut is
perpendicular to the fibers Fibers in twisted stems are not parallel to the stem
axis. Sections parallel to the fib rs (radial and tangential) must be split (Fig. 3.8). In
doing so, the fibe (or tracheid) direction becomes visible and the sample can be
correctly oriented in the sample holder of a microtome (Fig. 3.9). The microtome
blade must precisely follow the split plane on the wooden block in case of cutting
radial or tangential sections. If the sample is split in such a way that it is not pre-
cisely parallel to the rays, then they appear short and do not show all of the details.
See section 5 on p. 67 for more information.
Fig. 3.6. Visualization of the three cutting directions (transversal, tangential and radial) used in wood
anatomy (modied after: Schweingruber 1990, p. 13).
tangential
radial
transverse

20
Dry hardwoods, especially tropical woods, must be softened before cutting. This
can be achieved either within several minutes or several hours depending on the
density of the material (Fig. 3.10). Extremely dense wood (e.g., ebony wood) can
be softened in a vapor pressure pan (Fig. 3.11). Dense wood can also be softened
by soaking small samples in a mixture of 96% ethanol, glycerol, and water (1:1:1) for
several weeks. Before soaking or boiling, label the samples with a soft pencil, or
wrap the sample in a labeled, heat resistant textile.
Fig. 3.9. Splitting a block with parallel sides. Only
one side must be perfectly radial.
Fig. 3.7. Sectioning of a stem in a bench by a ne-toothed saw.
Fig. 3.8. Splitting a sample parallel to the bers.
Fig. 3.10. Boiling hard woods in an Erlenmeyer
ask on a hot plate.
Fig. 3.11. Very dense wood can be softened by
boiling it in a pressure cooker.

21
3.3 Seconing by hand
Preparing sections by hand is unlikely to result in high quality slides. For a quick
look, parts of plants can be clamped between holders of elder pith or carrot, and
then cut (Figs. 3.12–3.15).
Handmade sections of sub-fossil or waterlogged wood are well suited for species
identification Important Note: Only the thinnest section (e.g., 10–20 microns) is
suitable for microscopic inspection.
New razor blades have to be pulled slowly in a very steep angle across the object.
For example, cut the object like you would cut a salami, not like you would cut but-
ter. Never push blades across the objects.
Fig. 3.12. Correct guidance of the blade for a non-
permanent slide of living material.
Fig. 3.15. Cross-cut of a waterlogged, soft sub-
fossil oak sample.
Fig. 3.14. Correct guidance of the blade for a per-
manent slide of sub-fossil material.
Fig. 3.13. Incorrect guidance of the blade for a
non-permanent slide of living material.


22
3.4 Spling and seconing charcoal
or mineralized wood
When a small sample of charcoal has to be identifie it is best to simply break it
(Fig. 3.16). The structure of the cross sectional plane is perfect for episcopic obser-
vation. For radial and tangential sections, the sample can be split using a knife
(Fig. 3.17).
This same procedure is also useful for mineralized wood, which often occurs along
metallic archeological artifacts, e.g., scabbards. Anatomical details in the trans-
verse and longitudinal planes become distinct if all dust particles are blown out
with compressed air (Figs. 3.18 and 3.19)
Fig. 3.16. Breaking charcoal across the fiber
between ngers.
Fig. 3.18. A broken piece of mineralized wood,
transverse surface of Alnus sp. Photo: Willy Tegel.
Fig. 3.19. A split piece of mineralized wood, radial
surface of Alnus sp. with scalariform perfora-
tions. Photo: Willy Tegel.
Fig. 3.17. Splitting charcoal parallel to the fiber
with a knife.

23
To prepare microscopic cross sections which are useful for photographic presenta-
tion, apply a drop of a mixture of a two-component adhesive, e.g., Araldit, on the
transverse side of a broken charcoal surface (Fig. 3.20). The liquid fill all cavities
and polymerizes within 30 minutes, after which the surface can be planed with a
normal microtome (Fig. 3.21).
In the next step, a small piece of transparent self-adhesive tape (e.g., Scotch tape)
is attached to the top surface (Fig. 3.22). The microtome knife is pulled through
the charcoal at a sharp angle and a thickness setting of 20–30 microns. The sec-
tion adheres to the tape and can be mounted onto a slide (Fig. 3.22). At this point,
the section is ready for microscopic observations (Fig. 3.23). By using a mounting
medium, e.g. Canada balsam, the slide remains permanent, but the adhesive tape
has to be kept intact to preserve the integrity of the charcoal section. This pro-
cedure stabilizes the fragile carbonized structures. This procedure does not work
for longitudinal sections because the fragile carbonized structures split within the
adhesive (Fig. 3.24) (Schweingruber 2012).
Fig. 3.20. Two-component adhesive.
Fig. 3.23. Microscopic cross-sections adhering to
adhesive tape on a slide.
Fig. 3.22. Charcoal stabilized with a two-compo-
nent adhesive, covered with an adhesive tape in
a microtome holder. The knife cuts the charcoal
and the section remains intact by adhering to the
transparent tape.
Fig. 3.21. Flat transverse cut through a piece of
charcoal, which is stabilized with a two-compo-
nent adhesive.

24
Fig. 3.24. Microscopic cross section of a piece of charcoal of Betula sp.
Fig. 3.25. Cross-section of a Neolithic, carbonized piece of Abies alba with an insect gallery.
500 µm
1 mm

25
3.5 Preparaon of surfaces for macroscopic observaon
Fresh samples (stems, root, branches) are cut with sharp paper knives or razor
blades. Stick the 1 cm-long pieces of your sample into grafting wax or modeling clay
and let them dry for a few hours. The dry samples and charcoals can be stabilized
on glass slides or in petri dishes (Fig. 3.26).
Samples can be observed under the binocular microscope (Fig. 3.27). Objects in
petri dishes can be closed, labeled and stored after analysis (Fig. 3.28). Charcoals
with more or less flat broken surfaces can also be stabilized in grafting wax
(Fig. 3.29).
Fig. 3.28. Samples with labels, ready for storage.
Sample details can be written on the cover of the
Petri dish.
Fig. 3.29. Charcoal stabilized in grafting wax.
Fig. 3.27. Herb samples below a binocular micro-
scope for counting rings.
Fig. 3.26. Stems in grafting wax in a Petri dish.

26
3.5.1 Making annual rings visible by cung and sanding
For identifying rings on various kinds of samples (i.e., cores, discs or other pieces
of trees, or even entire stems, branches or roots of shrubs or herbs) it is, in most
cases, sufficie to cut a plane surface using a sharp paper knife or a razor blade in
a correct manner (Figs. 3.30–3.34; see section 3.3).
Fig. 3.32. Perfectly sectioned surface of a small
dwarf shrub (Acinos alpinus).
Fig. 3.33. Perfectly sectioned surface of a rhizome
of a shrub (Rubus idaeus).
Fig. 3.31. Incorrect sectioning: Never use a knife
like a planer.
Fig. 3.30. Correct sectioning: Always cut at an
angle to the sample, not across it.
  



27
By polishing surfaces with belt sanders (Figs. 3.34 and 3.35), contrasts between
the cell lumen and cell wall become more distinct. Sanded surfaces are of diffe -
ent quality (Figs. 3.36–3.38). All of them look good at small magnifications but at
higher magnification we recognize the dislocated dust particles in all cell lumina.
Only very fine-g ained surfaces (400–600 grit sandpaper) yield useful pictures. The
contrast of the sample can be enhanced by blowing the dust out of the sample with
pressurized air, and fillin the lumina with chalk.
Important Note: Sanded cores cannot be used for isotopic analysis because dust
particles are dislocated.
Fig. 3.36. Optimal planed sur-
face of a pine, all lumina are
filled with dust and cell walls are
clearly visible.
Fig. 3.37. Badly polished surface of a fi , cell walls
cannot be observed. Fig. 3.38. Badly planed surface of an ash, not all
lumina are perfectly filled with dust and scratches
left by the sandpaper obscure the structure.
Fig. 3.35. Small belt sander for
polishing small surfaces, i.e.,
increment cores.
Fig. 3.34. Big belt sander for
treating large surfaces.

28
3.5.2 Observaon surfaces by enhancing contrast
If the wood you are working with is light-colored, with little contrast between
annual rings (e.g., diffuse-porou species), the vessels and annual rings can be
made more distinct by fillin the cell lumina of cut surfaces with chalk. Important
Note: This is only possible if the surface is cut; it is not possible to apply chalk on
sanded surfaces! If your surface is cut, take a piece of white chalk, cover the surface
of the section with the chalk and then use your finge to rub the surface, thus press-
ing the chalk into the cells (Figs. 3.39–3.43).
If the wood is very light-colored, a two-step procedure will further enhance the
contrast, increasing the visibility of the ring structures. First, on the cut surface of
the sample apply a stain by using a dark (green or black) felt marker (e.g., a thick
Edding or other brands). Simply paint the cut surface and wait until the stain is dry.
Second, cover the surface with white chalk and rub it into the cells as described
above. It is important to wait until the stain is dry, otherwise the chalk absorbs the
stain and the result is just a blurred surface with more or less no contrast.
There is another possibility to make rings of extremely diffuse- orous species vis-
ible. After staining the cut surface with a dark felt marker, wait a few minutes to let
it dry and then cut approximately 0.2–0.5 mm from the dark stained surface. Due
to their differen capillarity, earlywood and latewood become distinct, because the
stain penetrates less into the latewood than into the earlywood (Fig. 3.44) (Iseli &
Schweingruber 1989).

29
Fig. 3.39. Applying chalk on a cut surface. Fig. 3.40. Rubbing chalk into the cell lumina.
Fig. 3.41. Enhanced contrast by rubbing chalk
dust in cell lumina of a species with small vessels
(Rubus idaeus).
Fig. 3.43. Enhanced contrast by painting the sur-
face with a marker and rubbing chalk dust in cell
lumina (Rubus idaeus).
Fig. 3.42. Enhanced contrast by rubbing chalk
dust in cell lumina of a species with large vessels
(Pueraria lobata).
Fig. 3.44. Enhanced contrast between earlywood
and latewood by painting the surface with a
marker and cutting the surface until ring struc-
tures appear (Vaccinium myrtillus).

30
3.6 Seconing with microtomes
For a more detailed analysis of wood anatomical features, samples have to be pre-
pared for microscopy and successive image analysis (Schweingruber et al. 2006).
As transmission and scanning electron microscopy is mostly used to analyze spe-
cifi features as cell wall structures, perforation plates or pittings in high magni -
cation (< 1000x) (Carlquist 2001), the most common method for wood anatomical
analysis in plant sciences is transmitted light microscopy. For this, micro sections
have to be prepared using microtomes, which can also be used for charcoal analysis
(see section 3.4).
It is important to note that microtome sections of fresh as well as of degraded
plant material can yield high-quality permanent slides. This is especially true when
using one of the three types of microtomes presented in this book, which were
most recently developed by us and which are commercially available (Fig. 3.45).
Fig. 3.45. Three new types of microtomes, developed for wood anatomical purposes: microtome type
GLS1, the WSL-Lab-microtome (modified Reichert-type) and the WSL-Core-microtome. Design and
Production: Sandro Lucchinetti, Schenkung Dapples, Zürich.
WSL-Core-microtome
WSL-Lab-microtome
Microtome GSL1

31
3.6.1 Microtome knifes
While further developing our microtomes, we found an efficie replacement for
the commonly used microtome knifes, as well as for the rather expensive remov-
able microtome blades.
For all our microtomes we developed a special removable blade holder, enabling
to use common NT-Cutter blades (A-type, 0.38) known as paper knifes (Figs. 3.46–
3.48). These blades are cheap, available in any office-supp shop, and they provide
a comparable quality for cutting micro sections than the really expensive special
removable blades for microtomes (Fujii 2003). The stability of the blade while cut-
ting is guaranteed by the fixatio plate slightly overlapping the main holder sup-
porting the blade from the top against the forces occurring during the cutting pro-
cedure (Fig. 3.46).
Commercial microtome knives and corresponding sharpening machines are also
available, but are only used in special cases (Figs. 3.49 and 3.50).
Fig. 3.47. Disposal blade (NT-cutter).
Fig. 3.46. Schematic view of the holder for removable blades (Gärtner & Nievergelt 2010, p. 88).
Fig. 3.48. Removable blade holder.Fig. 3.49. Commercial microtome knife with two
plane surfaces. Such types are useful for section-
ing very dense wood.

32
3.6.2 Microtome types
Until recently, when using common microtomes (Fig. 3.51), the main problem was
that these sections are actually restricted to rather small specimen in the range of
about 0.5 x 2 cm. For an expanded time series analysis, especially when working on
increment cores, much larger sections would be required to reduce the amount of
sample preparation to an affordabl minimum.
Our microtomes are all based on a fixed-sledg guidance with no internal play. This
detail guarantees a high stability of the microtome while cutting, and therefore
enables cutting of bigger samples. In addition, specialized holders were developed,
fittin into the microtome clamps, which enable working with increment cores of
5 mm and 10 mm diameter (Figs. 3.52 and 3.53), and also with micro-core samples
(e.g., collected with a puncher; compare section 2.1, Fig. 2.5) (Figs. 3.54 and 3.55).
The maximum length of cores is limited to 6 cm.
Fig. 3.50. Commercial sharpening machine.
Fig. 3.52. Holder for cores with 5 mm diameter.
Fig. 3.51. The microtome, type Reichert, is well
designed, but is no longer produced.
Fig. 3.53. Holder for cores with 10 mm diameter.

33
Fig. 3.54. Holder for micro-core samples with
2 mm diameter.
Fig. 3.56. Micro sections made from Pucher cores from Eucalyptus sp., normal light and polarized light.
Fig. 3.55. Stained sections from puncher samples.
500 µm 500 µm

34
3.6.2.1 WSL-Core-microtome
The core-microtome (weight: 14 kg; length: 80 cm) is designed to prepare planed
surfaces on increment cores up to a length of 40 cm (Gärtner & Nievergelt 2010).
Compared to sanding, the procedure commonly used to prepare surfaces on cores,
the cells of the annual rings remain open and are not fille with dust. The cell walls
are smooth after cutting, and thus clearly visible. This microtome was not designed
for producing micro sections, though it is perfectly suited to cut perfect surfaces
from increment cores. Important Note: The core-microtome is suitable for the
preparation of samples to be used in isotopic analysis.
Fig. 3.59. Scanned images of a Quercus petraea core. The upper image shows the core after cutting
without further preparation. In the lower image, the core was stained with a black felt marker and
then the cells were filled with white chalk to enhance the contrast for image analysis (modified after:
Gärtner & Nievergelt 2010).
◂ Fig. 3.57. Core-microtome,
side view. Developed by:
Arthur Kölliker; Design and
Production: Sandro Lucchi-
netti, Schenkung Dapples,
Zürich.
Fig. 3.58. Photos taken of the earlywood portion of a Larix decidua ring at two magnifications (cell
walls not stained). White areas on the images are caused by reflections of water on the core surface,
because the images were taken right after cutting (modied after: Gärtner & Nievergelt 2010).

35
3.6.2.2 Microtome type GLS1
Functional microtomes, designed to cut micro sections of wooden specimen, are
commonly bulky and heavy. In contrast, the design of the GSL1 combines maxi-
mum stability, minimal weight and reduced size for a fully functional, small and
lightweight microtome (30 x 14 x 12 cm, 3.6 kg). This special design allows using
the device not only in a laboratory, it can easily be transported to remote research
areas, enabling to cut micro sections right after sampling (Gärtner et al. in review).
The fact that the clamp of the sample holder is fixe tightly on the sledge enables
cutting all kind of wooden specimen, ranging from soft balsa wood to hard ebony
wood. The fixe position of the clamp is also a little disadvantage, because wooden
blocks have to be perfectly oriented before cutting, a fina tuning of the sample ori-
entation is not possible. With this microtome design, the sample is pulled over the
blade, which is mounted in a fixe oblique position and the carrier with the clamp
runs along one rail (Fig. 3.60).
Fig. 3.60. Microtome GSL1. Design: Sandro Lucchinetti; Production: Schenkung Dapples, Zürich.
Fig. 3.61. Cutting a wooden block using the GSL1. Fig. 3.62. Section of dense material: Rhizophora
mangle, cross section, cut with the GSL1-micro-
tome.
100 µm

36
Fig. 3.64. Micro-core holder mounted on the microtome clamp.
Fig. 3.63. Core holder for the microtome GSL1.

37
3.6.2.3 WSL-Lab-microtome (modied Reichert-type)
The lab microtome (Figs. 3.65 and 3.66) was developed based on the design of an
old Reichert microtome (see Fig. 3.51), which is no longer available on the market.
But it is not just a copy of the old device, it differ in some essential details. The
plate to  x the knife holder on is  xed on two sledges which are guided by two
rails. These parallel sledge guidance rails have no internal play, so the only possible
movement is forward and backward. There is no movement possible in any other
direction. This guarantees maximum stability while cutting, the knife cannot be
lifted upwards when the sample is very dense . For this, the lab-microtome is suited
for cutting all types of plant tissues ranging from soft herb stems or bark, to very
dense tropical wood. The stability of the entire construction even enables cutting
so called “Nördlinger” sections (Bubner 2008) (Fig. 3.67).
The knife (or removable blade holder) is mounted to a holder, which is adjustable
in the vertical and horizontal planes. The sample holder is manually tightened and
adjustable. Fine adjustments of the clamp allow for the correction of the sample
orientation for all sample types, e.g., wooden blocks, cores, cork-mounted soft tis-
sues, or very small, bent shoots. The microtome is also equipped with a semi-auto-
mated system that lifts the sample in a constant manner, where the uplift of the
sample is adjustable between 5 and 30 μm.
◂ Fig. 3.65. WSL-Lab-microtome (modi-
fied Reichert microtome). Design: San-
dro Lucchinetti; Production: Schenkung
Dapples, Zürich.
▴ Fig. 3.66. Back side of the lab-microtome show-
ing the semi-automatic sample transport con-
struction.

38
Fig. 3.69. Very large stained thin section of a larch (50 mm in diameter).
Fig. 3.68. Two examples for “Nördlinger” sections cut with the Lab-microtome. The sections can be
dried after cutting between two felt layers. Because of their thickness (60 μm) they are stable enough
to directly analyze their structure.
Fig. 3.67. Original of a “Nördlinger” section. Hermann Nördlinger, Professor of Forestry in Hohenheim,
Germany, in 1852–1888, made thin sections from 1100 tree species for macroscopic observations (Bub-
ner 2008). He distributed them commercially in volumes containing 50 species each.

39
3.6.3 Seconing with microtomes and observaon without staining
For all specimen it has to be guaranteed that the sample is correctly oriented in the
sample holder. Depending on the purpose, you can cut transversal (cross), radial
or tangential sections. For cross sections, the sample has to be positioned with the
fibe direction at 90 degrees relative to the blade. For radial and tangential sec-
tions, the fibe direction has to be parallel to the blade, whereas for radial sections
the cut has to be done parallel to the rays, directed towards the pith (or center of
growth).
3.6.3.1 Material of normal sness
Clamp the sample very tightly in the sample holder and cut the surface to a fla
plane before you intend to take a micro section, using water on a painting brush to
keep the surface wet. Without adding water to the surface, the cell walls tend to
break while cutting because they are more or less brittle. When adding water to the
top of the sample, the cell walls get moist and therefore more flexible
When the surface is plane and all saw marks are gone, bring the blade in a position
where it is definitel sharp or change the blade (Figs. 3.70 and 3.71). Now, hold the
brush on top of the wet object and pull the blade slowly in a steep angle (pulling
angle) under the brush (Fig. 3.72). By using water even on top of the blade, the
section glides and does not roll-up on the blade, and can be transported from the
brush to a glass slide (Fig. 3.73). Water or glycerol are perfect gliding liquids for sec-
tions, although on soft tissues, sectioning requires the use of ethanol 96% or 99% as
a gliding solution, which helps stabilizing the tissue.
Fig. 3.70. Position of the pulling angle of the blade
in relation to the object. The optimum angle of
the knife is around 45 degrees, but depending on
the species of your sample and the desired qual-
ity of your section, this angle should be adjusted
until an optimal section is achieved.
Fig. 3.71. Position of the angle of inclination of the
blade in relation to the object. For hard woods,
e.g., tropical woods the incline should be fairly
steep. In soft material, e.g., archeological wet
wood, the incline should be as at as possible.

40
Fig. 3.73. ‘Swimming’ or gliding a section from
the blade onto the glass slide.
Fig. 3.72. Keeping the section flat with a painting
brush on the blade.
If the section rolls while cutting, you can prevent this to a certain degree by placing
the brush on top of the section to keep it flat If the sample rolls anyway or even
after cutting, put a needle or the tip of a pair of tweezers into the roll and massage
the section carefully onto your finge tip to unroll it and harden it with a drop of
ethanol. If the section remains flat place it from the finge tip directly onto the
slide (Figs. 3.74a–d). Do not stain the section when it is rolled, as the result would
be irregular stain-patterns.
Fig. 3.74. a) The tip of a pair of tweezers must be placed in the center of the rolled section. b) and c)
Massage the rolled section on the finger tip over the conical tweezer tip. Stabilize the section in a flat
position on the finger tip by putting a drop of absolute ethanol on it. d) Put the flat section onto the
slide and start the staining process.
a
c
b
d

41
3.6.3.2 Seconing conifers
Perfect sectioning of conifers is a challenge because secondary walls are only lightly
bonded with primary walls especially in earlywood tracheids. Because of this, it is
absolutely necessary to use a sharp blade to avoid splitting o the primary walls
and bending the cell lumen area inwards (Fig. 3.75). The break-away of the second-
ary walls can also be avoided by using the full length of the blade for sectioning.
In addition to the above-mentioned procedure, cell lumina can be fille with a
non-Newtonian fluid The principle of a non-Newtonian flui is that it changes its
consistency from a viscose flui to a solid as soon as it is placed under mechanical
pressure. Loïc Schneider, of the WSL Dendroecology group, created such a flui by
adding some drops of water and glycerol to corn-starch (Schneider & Gärtner, in
press). The consistency of the mass is right if the paste “crumbles” when stirred
and flow when at rest. Then it should be liquid enough to just flo into the cells
when applied to the surface of the sample. When the blade presses against the
surface the flui acts as a solid and therefore stabilizes the cells (Fig. 3.76).
If you intend to electronically measure cell wall and cell lumina dimensions, this
simple procedure saves a lot of laborious effor to digitally correct the images.
Fig. 3.75. Problematic section: pulled-out sec-
ondary walls in the earlywood cells of a conifer.
Pseudotsuga menziesii. Section courtesy by Jodi
Axelson.
Fig. 3.76. Perfect section: Secondary walls in the
earlywood are connected with the primary walls.
Pseudotsuga menziesii. Section courtesy by Jodi
Axelson.
100 µm 100 µm

42
3.6.3.3 Seconing of very so material
3.6.3.4 Stabilizaon and seconing of
waterlogged archeological wet wood
Sections of very soft tissues often get compressed at the very slight flang between
the blade and the holder. If the blade is not fully inserted into the holder, the gliding
space is larger and the section glides without any mechanical resistance onto the
blade (Fig. 3.77).
When a high-quality section is ready, you should immediately label the slide either
with a soft pencil on a self-adhesive label or with a permanent fin marker directly
on the glass (Figs. 3.78 and 3.79). Important Note: Labels written with ball pens
and/or permanent fin markers will dissolve in ethanol.
Handmade sections using razor blades are useful for wood identificatio (see sec-
tion 3.3). For studying large surfaces, or decomposed samples, e.g., structures
with degraded cell walls, the material can be soaked with polyethylene glycol 1500
(PEG = Carbowax). The Carbowax flake must be fluidize at temperatures of 60°C.
Then, anatomically prepared blocks with a size of approximately 1 cm3 are placed in
the viscous liquid of the Carbowax (Fig. 3.80). The material has to be saturated for
at least 24 hours at 60°C in an oven. At temperatures of 20°C the saturated blocks
are stable enough to be sectioned with the microtome. Since Carbowax is water-
soluble, use glycerol or ethanol instead of water for sectioning. Once a successful
section is made, follow all the steps described in section 4. If the sections are thin
enough, anatomical characteristics can be recognized in detail (Figs. 3.81–3.84).
Important note: For these types of samples, the pulling and inclination angles of
the blade must be minimal.
Fig. 3.77. Protrude disposable blade fixed in the
blade holder. Soft sections slip onto the flat sur-
face of the blade.
Fig. 3.79. Temporarily labeled slides with a soft
pencil on self-adhesive paper labels, ball point ink
dissolves in ethanol and xylene.
Fig. 3.78. Temporary labeled slides with a perma-
nent marker, be aware that it dissolves in ethanol
or Eau de Javelle.

43
Fig. 3.84. Radial section. Preserved are the rays
(red) and degradation products in ray cells as well
as hyphae of fungi in the vessel (blue) and tertiary
walls of bers (bluish).
◂ Fig. 3.80. Polyethylene glycol 1500. Commercial
flakes at room temperature (right). Heated PEG
solution at 60°C containing wet wood samples
(left).
▾ Figs. 3.81–3.84. Sections of waterlogged arche-
ological soft wet wood preserved with polyethyl-
ene glycol (Carbowax) and stained with Astrab-
lue/Safranin, Alnus sp.
Fig. 3.81. Cross section of a large radially split sec-
tion.
Fig. 3.82. Cross section of an annual ring bound-
ary. Secondary walls of fibers are degraded. Pre-
served are primary and tertiary walls.
Fig. 3.83. Tangential section. Preserved are the
rays, and almost completely degraded are the
vessel-walls.
500 µm 50 µm
50 µm 50 µm

44
If you are simply interested in determining the age of the section, and in recogniz-
ing the annual ring boundaries, staining is not mandatory. Without staining, ana-
tomical details can be recognized by closing the condenser (Figs. 3.85 and 3.86), or
using polarized light (Figs. 3.87 and 3.88).
▾ Figs. 3.85–3.88. Unstained sections of very soft herb stems.
Fig. 3.85. Daucus carota, 2-year-old semi-ring
porous polar root, normal light transmission.
Fig. 3.86. Bellis perennis, 2-year-old vessel-less rhi-
zome, normal light transmission.
Fig. 3.87. Centaurea jacea, 4-year-old polar root,
polarized light. Fig. 3.88. Medicago lupulina, 2-year-old polar root,
polarized light.
100 µm
100 µm
250 µm
250 µm
3.6.3.5 Observaon of so herb stems

45
Fig. 3.89. Two slides with sections of herbs, preserved in glycerol since one year.
Fig. 3.90. Slide holder with unstained sections
and temporarily labeled slides. Fig. 3.91. Stack of slide holders in the horizontal
position.
3.7 Storing glycerol-preserved secons
If the sections don’t have to be stained, they can be covered with glycerol (glyc-
erin) and a cover glass, which prevents the sections from drying out (Fig. 3.89).
Such preparations can be stored for months in commercial slide holders (Figs. 3.90
and 3.91). The holders must be stored horizontally, otherwise the cover glasses
move sideways.

46
3.8 Maceraon and measuring axial cell dimensions
Normally, the length of vessels or ray cells can be measured on thin radial sections
(Fig. 3.92). By observing sections you precisely know in which intra-annual position
the measured vessels occur, i.e., in the earlywood or latewood. Maceration is nec-
essary for measuring fibe length. For ecological studies it is therefore important to
isolate a slice within the annual ring (e.g., in the earlywood or the latewood). For
technical analysis, e.g., for pulp studies, the samples must represent a mean of the
whole wooden block.
After comparing differen maceration methods proposed in the literature, we rec-
ommend the following method.
Dispensing the mazeration solution:
1 part hydrogen peroxide (H2O2) + 1 part concentrated acetic acid + 1 part water
Fill small Erlenmeyer flask with 50 ml of the maceration solution and add several
wood pieces not larger then a match in diameter. After boiling the material (Fig.
3.93) for 3–6 hours, the wooden pieces are completely bleached (delignified and
the middle lamella of the fiber are dissolved (Figs. 3.94–3.97). Take small pieces
with a tweezer and put them onto a water-permeable, non-staining tissue and
wash the sample with water by using a pipette. When the solution does not smell
anymore, tease the sample until the fiber are visible. Now put a drop of Astrablue
on the tissue with the fibers Dehydration and embedding is described in sections
4.9 and 4.10.
With Eau de Javelle (bleach) it is possible to macerate fiber-nest of wasps (Fig.
3.98). The nest-tissue decays after soaking for a few minutes with bleach. After
rinsing the fibe solution with water, it can be dehydrated and stained with Astra-
blue.
Fig. 3.92. Aspect of vessel length (indicated by
the two arrows) in a radial section of a Mediter-
ranean dwarf shrub. Pallenis spinosa.
Fig. 3.93. Mazeration of wood samples in an Erlen-
meyer flask, containing a solution of hydrogen
peroxide, acetic acid and water, on a heater.
100 µm

47
Fig. 3.98. Macerated bers of a wasp-nest.
Fig. 3.94. Macerated long tracheids of a conifer, Pinus cembra.
Fig. 3.97. Macerated fibers (1), rays (2) and vessels
with helical thickenings (3) of a tree, Tilia sp.
Fig. 3.96. Macerated short fibers, vessels and
ray cells of a broadleaf dwarf shrub, Empetrum
nigrum.
Fig. 3.95. Macerated tracheids of Pinus cembra
with fenestrate ray-tracheid pits and bordered
pits.
250 µm
50 µm
100 µm
250 µm
1
3
2
500 µm

48
3.9 Preparaon of texle bers
Fibers of all textiles, e.g., clothes, lines, ropes and artificia tissues, are already mac-
erated. Their structure can be analyzed by just picking some fiber out of the tissue
using, e.g., a pair of tweezers. After dehydration on a glass slide and embedding
in Canada balsam as described in section 4.8, most structural details are visible in
polarized light. Therefore, staining of the fiber is not necessary. The following fi -
ures illustrate images taken using polarized light from unstained fibers extracted
from textiles and ropes (Figs. 3.99–3.102).
Fig. 3.99. Isolated Nylon bers from a shirt. Fig. 3.100. Isolated cotton ber from a shirt.
Fig. 3.101. Bundle of fibers in a string of flax of
Central Europe. Fig. 3.102. Bundle of fibers in a string of agave
from Mexico.
250 µm
100 µm 100 µm
100 µm

49
3.10 Preparaon of technically composed wood
Plywood is the result of a technical procedure, where several veneers are glued
together in opposite  ber directions (Figs. 3.103 and 3.104). The main goal of this and
most other arti cially recombined wood parts or components is the elimination of
anisotropy.
Even plywood can be sectioned with microtomes and stained like common wood
samples. Microscopic slides allow the identi cation of the tree species used because
the structure is clearly visible. Furthermore, the penetration of the glue into the wood
structure at the contact surfaces becomes visible (Figs. 3.105 and 3.106).
Sectioning particleboards without embedding succeeds only in the direction of the
longitudinal axis of  bers and chips. The staining process is the same as for normal
wood.
Fig. 3.103. Cottonwood-Plywood consisting of 3
layers, stained with Safranin only.
Fig. 3.104. Birch-Plywood consisting of 5 layers,
sectioned in opposite directions and stained with
Safranin only.
Fig. 3.105. Glued joint of the Cottonwood-Ply-
wood where a huge amount of glue has been
used (irregular, intensely red areas, see arrows).
Fig. 3.106. Glued joint of the Birch-Plywood with a
minimal use of glue (arrows).
250 µm 250 µm
1 mm 1 mm

50
Fiber boards consist of chemically macerated wood and mechanically destroyed
fiber (Figs. 3.107–3.112). Wooden hardboard plates consist of common wood bits
and pieces, which have been compressed under high pressure (Fig. 3.113). Section-
ing these types of artificia wood is possible in all directions.
Fig. 3.107. Particle boards consist of thin, axially
sectioned wood chips, which are oriented in all
directions. Section stained with Safranin only.
Fig. 3.108. Radial chip of a cottonwood stem.
Microscopic anatomical features are well pre-
served and can be used for wood identification.
Reticulate pits (arrow) in the homogeneous ray
are an indication for Populus sp.
Fig. 3.109. Tangential chip of cottonwood stem. Fig. 3.110. Foreign wood type with crystals
(arrows) in a cottonwood chip board.
1 mm 100 µm
100 µm 50 µm

51
Fig. 3.112. Fiber board with decomposed pits (1) in
conifer tracheids and glue (2).
Fig. 3.111. Isolated and broken bers are characteristic for pressed ber boards. Polarized light.
Fig. 3.113. Wooden hardboard is characterized by
bent rays and deformed vessels and fibers. Popu-
lus sp.
1
2
50 µm
500 µm
250 µm

52
4. Fixaon, bleaching, staining,
embedding, cleaning, labeling and storing
4.1 Eect of staining, and safety instrucons
Samples have to be stained, dehydrated and embedded in order to be permanent.
Staining enhances the contrast of cell-wall structures and cell contents for observa-
tions and/or image analyses. Depending on the dye used, differen cell types and
cell contents can be distinguished.
Preparing staining soluons
»When preparing the liquid dyes, always follow the safety instructions. Powders
are poisonous!
»For preparing the solutions needed for staining, you need a balance, plastic or,
if possible, glass bottles with a minimum capacity of about 200 ml, inhalation
protection, and gloves.
»Before you start the procedure, make sure to cover the table around the bal-
ance and below the dye-powder boxes with some paper tissue. It is also best
to wear old clothes, gloves and goggles. If you only loose a single grain of the
powder, it might stain everything it gets into contact with, so be careful. More-
over, it is severely irritating to the skin if it gets in direct contact.
»Before starting the preparation process, bring the bottles, chemicals (Fig. 4.1),
and instruments into a logical order (Fig. 4.2).
Fig. 4.1. Holder with chemicals for preparation of
permanent slides. It contains dyes, ethanol and
xylene and corresponding pipettes.
Fig. 4.2. Preparation tools: gloves, paint brushes,
tweezers, pipettes, needle, paper knife, marker,
slides and cover glasses.

53
The entire preparation process (staining and dehydration) can be done on the glass
slide finall used for the permanent slide (Fig. 4.3). Use a pipette fille with the
chemicals needed for the respective process (i.e., one for the dye and another for
ethanol 75%) to begin the dehydration process and to rinse the section (Figs. 4.3–
4.5). Collect all liquids coming o the slide in a waste box, preferably of glass, as
some chemicals, i.e., xylene, dissolves some types of plastics (Fig. 4.6)! If you have
to work many hours with chemicals it is best to do so under an extractor hood or
near an open window (Fig. 4.7).
Fig. 4.3. Schematic view of the staining/cleaning process. Use a pipette for the chemicals and run the
liquids over the section. For removing the surplus stain you can also place the pipette on top of the
section and pump the ethanol through. By doing so you ensure that the dye is also removed from very
small cell lumina.
Fig. 4.5. Dye on a slide with a piece of tissue and
very small micro sections.
Fig. 4.4. Dye on a slide with micro sections.
Fig. 4.6. Waste box that is resistant to Eau de
Javelle, ethanol and xylene.
Fig. 4.7. Extractor hood.

54
4.2 Fixaon of cell contents
For cytological studies, plant tissues must be soaked with a fix tion solution (Fig.
4.8). This procedure is necessary because without fixatio the cell contents includ-
ing the nuclei degrade in the ethanol when dehydrating the samples after staining.
There are several possible procedures:
Fresh material should be preserved in FAA solution or ethanol 40%. FAA solution
and ethanol preserve cell contents but cannot be stained with Astrablue/Safranin.
Dispensing the FAA solution:
5 parts ethanol 96% + 1 part acetic acid + 1 part formaldehyde + 3 parts distilled water
If nuclei and protoplasts are to be visualized by dyes, they must be preserved
through an additional fixatio (Nawashin solution) (Fig. 4.8). The two possible pro-
cedures are:
»The samples have to be soaked for 3–6 days (Fig. 4.9). Afterwards they must be
stored in a glass or plastic bottle containing 40–60% ethanol .
»Micro sections of fresh material, or samples preserved in FAA or ethanol, have
to be covered for a few minutes with Nawashin solution. Staining is possible
after carefully rinsing them with water (Fig. 4.10).
Dispensing the Nawashin solution (Purvis et al. 1964):
10 parts of 1% chromic acid + 4 parts 4% formaldehyde + 1 part acetic acid
Fig. 4.8. Nawashin solution in a
bottle and small glass contain-
ers with ethanol 75%, for perma-
nent sample storing.
Fig. 4.10. Fixation of micro sec-
tions with the Nawashin solu-
tion on the slide.
Fig. 4.9. Fixation of cores with
the Nawashin solution in test
tubes.

55
4.3 Bleaching: Destrucon of cell contents and
dark-stained cell wall components
If unstained sections are brownish, this coloration indicates that the wooden sam-
ple contains phenols (Fig. 4.11). Although the presence of phenols is an indication
for plant physiological processes (e.g., heartwood formation, compartmentaliza-
tion (Shigo 1984)), they should be removed before staining the section if one is only
interested in the structure of the cells. This is because phenols, like lignin, stain red
in Safranin and for this blur the structure of the cell walls.
To dissolve the phenols, put a few drops of Eau de Javelle (bleach) on the slide
until the sections appear light (for about 5 minutes) (Fig. 4.11, right). Eau de Javelle
available for laboratories is a 13% solution. Common bleach as it is available at retail
contains around 5% of Javelle. When using this, you should cover the samples a few
minutes longer. Before staining, rinse the slide well with water until you can no
longer smell the bleach.
The application of Eau de Javelle to micro sections before staining in general has a
positive effec for image analyses, because bleaching also enhances cell wall con-
trasts (Fig. 4.12).
Eau de Javelle (bleach) is composed of:
sodium hypochlorite (2 NaOH + Cl2) / potassium hypochlorite (2KOH + Cl2)
Fig. 4.11. Left: Unbleached micro section (Vitis
vinifera). Many cell walls contain brownish sub-
stances, i.e., phenols. Right: Bleached micro sec-
tion (Vitis vinifera). Cell walls appear light and
transparent.
Fig. 4.12. Left: Unbleached micro section (Vitis
vinifera) after staining with Safranin/Astrablue.
Blue and red colors are not well di erentiated.
Right: Bleached micro section (Vitis vinifera)
after staining with Safranin/Astrablue. Blue and
red colors are well dierentiated.
500 µm 500 µm

56
4.4 Staining cell walls
Safranin/Astrablue stains mainly cell walls. Basically, Safranin stains red and cre-
ates a very good contrast to further analyze the cell structures using various image
analyses programs. Many other dyes are recommended in various textbooks (e.g.,
Rudzin 1999, Mulisch and Welsch 2010), but in our opinion, Safranin/Astrablue cre-
ates the best contrasts between differen cell-wall types (see section 4.2). Safranin
stains lignifie cell structures red, and Astrablue stains unlignifi d structures blue
(Figs. 4.13 and 4.14). Safranin/Astrablue also stains gelatinous fib rs and makes vari-
ous chemical compositions visible (Fig. 4.15).
When mixing both solutions (not the powder!) use a ratio of 1:1. There is no need
to stain using one dye after the other, instead they can be mixed, staining differen
structures in the section simultaneously.
Dispensing the Safranin solution:
0.8 g Safranin powder in 100 ml distilled water
»Never use common tap water for this procedure. Close the bottle tight and
shake it until the solution is dark red and no clumps are visible.
»When shaking the solution, you create a certain amount of foam. This foam
does not affec the stain and it will disappear after a few hours.
Dispensing the Astrablue solution:
0.5 g Astrablue powder in 100 ml distilled water + 2 ml acetic acid
Without the acetic acid the dye will not remain stable.
»Never use common tap water for this procedure. Close the bottle tight and
shake it until the solution is dark blue and no clumps are visible. The Astrablue
powder requires a more intense and longer shaking than Safranin to remove all
the clumps.

57
Fig. 4.15. Safranin/Astrablue stained xylem with
tension wood of a tree (beech). Red stained are
the primary walls of fibers and the secondary
walls of fibers in the latewood zone. Dark blue
stained are the gelatinous fibers and the axial
parenchyma cells.
Fig. 4.13. Safranin/Astrablue stained flower stalk of a Gramineae. Lignified (red) are the radially ori-
ented beams of fibers, the vessel cell walls and the external walls of the epidermis. Unlignified (blue)
are most parts of the cortex and the phloem within the vascular bundles.
Fig. 4.14. Safranin/Astrablue stained stem of an
annual climbing Cucurbitaceae. Lignifie (red) are
the xylem of vascular bundles and fibers in the
cortex. Unlignified (blue) are the rays and major
parts of the bark.
500 µm 250 µm
250 µm

58
4.5 Staining cell contents
Picric-Anilinblue is an excellent dye to visualize organic cell contents. Protoplasts,
nuclei and even chromosomes appear dark blue and cell walls remain yellow (Fig.
4.16). With the detection of living and dead cells, heartwood and sapwood bound-
aries become clear, as nuclei are only present in rays of the sapwood, never in
heartwood.
This staining can be recommended for studying xylogenetic processes and the
detection of sapwood/heartwood boundaries.
Staining procedure: Put some drops of Picric-Anilinblue on sections on a slide. Heat
them for a few seconds to approximately 80°C (Fig. 4.17). Clean the section with
water and dehydrate it with ethanol 96%, ethanol absolute and xylene. Embed the
section in Canada balsam.
Dispensing the Picric-Anilinblue solution:
1 part saturated Anilinblue + 4 parts saturated picric acid (trinitrophenol),
dissolved in 95% ethanol
4.5.1 Nuclei and protoplasts
Fig. 4.16. Picric-Anilinblue stained xylem/cambium/
phloem zone of a conifer (Abies alba), cored at the
end of the growing season. Blue stained are the
protoplasts including nuclei and the unlignifi d cell
walls. Yellow stained are the lignied cell walls.
Fig. 4.17. Heater for warming the Picric-Anilinblue
solution on slides.
50 µm

59
Starch grains indicate where carbohydrates are stored within the tissue (Eades
1937).
Lugol’s solution (Lugol’sche Lösung = iodine and potassium iodide solution) has a
brownish color (Figs. 4.18 and 4.19).
Put a drop of Lugol’s solution on the section on a slide. After a few minutes, starch
grains appear brown to violet and lignifie cell walls yellowish. The staining effect
disappear after a few hours. Making permanent slides is not possible. Preserve the
sections for observations in glycerol.
Starch grains are also visible in polarized light. They appear as light circles with dark
Maltesian crosses (Fig. 4.20).
Dispensing Lugol’s solution:
10 g of potassium iodide + 5 g of iodine in 100 ml distilled water
4.5.2 Starch grains
Fig. 4.18. Dark stained starch
grains (arrows) with the Lugol‘s
solution in rays of a conifer
(Abies alba).
Fig. 4.19. Dark stained starch
grains (arrows) with the Lugol’s
solution in rays of a broadleaf
shrub (Corylus avellana).
Fig. 4.20. Unstained starch grains
(arrows) reflec ed in polarized
light.
50 µm 50 µm50 µm

60
4.6 Staining cell walls and cell contents
For studying the timing of xylogenetic processes it is necessary to recognize cell
contents as well as the lignificat on of cell walls. A combination of Astrablue/Safra-
nin and Picric-Anilinblue dyes yields excellent results. Protoplasts appear blue and
nuclei dark blue, unlignifie cell walls light blue and lignifie cell walls red (Figs. 4.21
and 4.22).
Stain the section on the slide with Astrablue/Safranin for 3–4 minutes, rinse the dye
with water and then put some drops of Picric-Anilinblue on the section and heat it
until the slide gets hot (approximately 80°C). Now rinse out the dyes with water,
dehydrate, and embed in Canada balsam.
Fig. 4.22. Astrablue/Safranin and Picric-Anilinblue
stained radial section of a conifer (Pinus sylves-
tris). Dark blue stained are nuclei (1), light blue
tori of pits (2), and reddish are cell walls of axial
tracheids (3) and ray-tracheids (4).
Fig. 4.21. Astrablue/Safranin and Picric-Anilinblue
stained cross section of the xylem of a dwarf
shrub. Dark blue stained are nuclei (1), light blue
secondary walls of fibers (2), and red parts of pri-
mary cell walls of bers and vessels (3).
1
1
4
3
3
2
2
25 µm 25 µm

61
4.7 Staining hyphae, bacteria and decomposed cell walls
Recognition of hyphae is important for the interpretation of wood degradation.
Well approved is Picric-Anilinblue for staining hyphae. Picric-Aniliblue stains hyphae
and bacteria dark blue, lignifie cell walls yellowish and de-lignifie cell walls blue
in various intensities (Figs. 4.23–4.26).
Dispensing of the solution of Picric-Anilinblue is described in section 4.5.1. Staining
procedure: Put some drops of Picric-Anilinblue on sections on a slide. Heat them for
a few seconds to approximately 80°C. Clean the section with water and dehydrate
it with ethanol 96%, ethanol absolute and xylene. Embed the section in Canada bal-
sam.
Staining with Astrablue/Safranin and Picric-Anilinblue creates contrasts, which
show cell wall degradations and hyphae (Fig. 4.27).
Fig. 4.23. Dark blue stained
hyphae with Picric-Anilinblue.
Fig. 4.27. Secondary walls of fibers in the xylem
of a broadleaf tree. White rot fungi decomposed
and de-lignified mainly the secondary walls of
bers. Hyphae grow spiral-like in the rays.
Fig. 4.26. Blue stained red rot caverns in latewood
tracheids. Lignied parts appear yellow.
Fig. 4.25. Blue stained bacteria
in tracheid pits of a drift wood
conifer, Larix sp.
Fig. 4.24. Blue stained hyphae
in a vessel of a de-lignified wet
wood.
25 µm
100 µm 50 µm 25 µm
50 µm

62
4.8 Dehydraon
Dehydration is necessary to prepare permanent slides for the following reason: The
refraction index of water (n = 1.33) is lower than that of microscopic cover glass
(n = 1.523). The finalize section will at the end be embedded in Canada balsam, a
resin having more or less the same refraction index (n = 1.51–1.54) as the glass of the
micro slides and the cover glass.
When the samples have been covered by the stain for 3–5 minutes, they are ready
to be processed. Remember: all further steps occur on the slide (Figs. 4.28 and
4.29). First, use water to wash away the surplus stain until the water running over
the sample is more or less clear. The use of water for the firs basic cleaning helps
save ethanol.
Now you can start the dehydration process by using ethanol 75%. When this runs
clear o the sample use ethanol 96%. To really remove the stain from the cell lumina
and inter cellular spaces you can place the pipette directly onto the sample and
pump the ethanol through the sample. Do this until the ethanol runs more or less
clear. If the sections are rolled, it is possible to unroll them (see section 3.6.2). To
finaliz the dehydration process, rinse the sample with anhydrous ethanol.
Dispensing anhydrous ethanol:
95 ml ethanol 96% + 5 ml 2,2 dimethoxypropane (acetone dimethyl acetal)
Finally, prepare the section for embedding. Rinse the section with some drops of
xylene or Histoclear. If the liquid runs clear, you can continue embedding the sec-
tions in Canada balsam. If the liquid gets milky, repeat the dehydrating process with
anhydrous ethanol, and finall rinse with xylene again.
Fig. 4.29. Dehydration of tiny sections of grasses
on a tissue.
Fig. 4.28. Dehydration of a section of wood.

63
4.9 Embedding micro secons
Canada balsam and Euparal have excellent optical qualities but are slightly acidic.
Therefore, some stains for cell contents bleach out after several years. Further-
more, Euparal has a slight shrinking tendency. In our opinion, Canada balsam is well
suited for all stains presented here. Colors remain stable over decades and shrink-
ing does not occur. The only little drawback, despite the fact that xylene has to be
used to prepare the section before embedding in Canada balsam, is the slight oxi-
dation (yellowish to brownish color) of the balsam at the edges of the cover glass
after several years. However, this does not disturb the optical clarity. A very nice
example for the durability of micro sections embedded in Canada balsam is a micro
slide prepared by Nördlinger in 1878 (Fig. 4.30). For all other embedding resins, we
have no long-term experience.
The preparation of large sections is differen from preparing small sections. For a
large section (i) put a small drop of Canada balsam on top of the section, (ii) place
the edge of the cover glass on one side of the object without touching it and (iii)
fold down the cover glass slowly to fully cover the object. For small sections pre-
pared on a tissue, put a drop of Canada balsam on the glass and turn the tissue with
the sections towards the Canada balsam (Figs. 4.31 and 4.32). The small sections
stick in the Canada balsam and the tissue can be taken away. The sections are now
ready to be covered with the cover glass (Fig. 4.33).
When the cover glass is placed on top of the sections, you should start moving the
tweezers on top of the cover glass while exerting a little pressure to it. In doing
so, the section will be flattene below the glass, excessive Canada balsam will be
pressed out and you avoid the enclosure of air bubbles (Figs. 4.34 and 4.35).
Fig. 4.30. A 135-year-old micro slide of an unstained stem of a silver fir [Abies pectinata (syn.: Abies
alba)]. The section is preserved in Canada balsam. The section is transparent, only the borders around
the cover glass appear brownish.

64
Fig. 4.35. Covered section. The Canada balsam
spreads itself over the whole surface of the glass.
Fig. 4.31. Tissue with small sections next to the
glass with a drop of Canada balsam. Fig. 4.32. Turning the tissue with the sections
towards the Canada balsam.
Fig. 4.33. The small sections are trapped in the vis-
cous Canada balsam. Fig. 4.34. Covering the section with the cover
glass. Enclosing air bubbles can be avoided by
putting the glass down from one side to the
other.

65
4.10 Drying and stabilizing
Slides are placed between two heat-resistant plastic strips on an iron plate. Mag-
nets on the plastic surface press the sections fla (Figs. 4.36–4.38). Without the
plastic strips, dried Canada balsam would stick to the plate and it would not be pos-
sible to remove the slide without destroying it. If you have no magnets available,
use any small, but heavy objects (i.e., box with glass slides) to place on top. The
magnet (or weight) is needed to prevent the cover glass from being lifted during
the drying process. The slide is dried out after a few days to weeks at room temper-
ature or after 12–24 hours in an oven with a temperature of 60°C (140°F) (Fig. 4.39).
Fig. 4.36. Place the embedded sample (slide) on top of a heat resistant plastic strip on a metal plate
(1). Place a second strip on top and fix it with a magnet (or other weight) (2). Repeat the procedure for
all other slides (3). Always have a (heat resistant) plastic strip below and above the slide!
Fig. 4.37. Slides on an iron plate between plastic
strips, and weighed down with magnets.
Fig. 4.38. Serial production of sections ready to
be dried.
Fig. 4.39. Serial production of sections in an oven.

66
4.11 Removing from plasc strips and cleaning slides
When removing the slides from the oven, make sure that you do not touch the
magnets or slides before they are cold. As long as the slides are warm, the Canada
balsam is still liquid and the samples will be ruined when removing the plastic strips
or even the magnets.
As soon as the slides are cold you can start to remove the magnets very carefully.
For best results press down on the covered slide with two fing rs and slowly pull
o the magnet. Then take the upper plastic strip covering the slides and slowly pull
it o laterally. Make sure to not pull it upwards (Fig. 4.40).
After removing the slides from the plastic strips, the glass needs to be cleaned, i.e.,
the Canada balsam which squeezed out on top and below the slides during the dry-
ing process in the oven needs to be removed.
For removing the Canada balsam, never use chemicals. The easiest and fastest way
to clean the slides is using a razor blade or a paper knife (Fig. 4.41). Place the blade
fla on the slide and just scrape the dried balsam o the glass.
Fig. 4.41. Cleaning dry slides with a paper knife.Fig. 4.40. Cold micro slides after removing the
magnets (upper image). Note that the Canada
balsam has spilt over the slides. Always pull o
the plastic strip sideways in order to not break
the slides (lower image).
sideways
upwards


67
Fig. 4.42. Various types of labeling.
4.12 Labeling
4.13 Operang a line for staining, dehydrang and
embedding
This is a very important step in preparing micro sections in order to avoid mix-ups of
your samples. The (basic) labeling should already be done after cutting the section
and placing it on the glass slide. But be aware that during the staining and dehydra-
tion process there is a risk that the label will be dissolved. Make sure that you place
the writing on one side of the glass so that you can hold the slide with your  ngers on
this side during the staining to embedding process. As soon as the slide is  nalized,
i.e. the slide is cleaned, you should decide what long-term labeling your slides require.
Modern, self-adhesive labels will last for decades on the slide (Fig. 4.42).
In most cases you do not stain just one sample. Although this is possible, the more
common situation is that you have to process many samples, which are covered by
glycerol, in a row. To avoid waiting for each single sample covered by the dye, you
can open up a line. This can be done as follows:
»You take the first slide, remove the glycerol from the sample by rinsing it with
water, cover the section with the stain and place it on the desk.
»Then take a second slide, remove the glycerol and also cover this section with
the stain and place it next to the firs slide.
»Wait for another 1–2 minutes, then take the first slide and start the preparation
process (washing o the stain, dehydrating, rinsing with xylene, embedding)
and place the slide between the plastic strips and load it with a magnet or any
other weight.
»Then take a new (third) slide, remove the glycerol, cover it with the stain and
place it next to the second slide, which is now covered by the stain for about
4–5 minutes.
»Take the second slide and start again the dehydration and embedding process
and fi it between the plastic strips.
»Take a new slide, cover it with the stain, place it on the desk, and continue with
the slide already covered by the stain ….. and so on.

68
4.14 Repairing cracked (embedded) slides
4.15 Final storage
Sometimes it can happen that you drop a slide, or you accidently break one for
another reason. In most cases, these cracked slides can be repaired. Take the bro-
ken slide and place it in xylene for about 10 hours. This dissolves the Canada bal-
sam and you can take out the micro section and embed it onto a new glass slide
(Figs. 4.43 and 4.44).
Important Note: Remember to soak the sample in a glass container as xylene can
dissolve plastics.
There are differ nt possibilities to finall store the embedded slides. They can be
stored in prefabricated aluminum plates within special shelves (Figs. 4.45 and
4.46), or, more commonly, in special slide boxes (Figs. 4.47 and 4.48) that can be
stacked on any shelf (Fig. 4.49).
Fig. 4.44. Cracked slide in a petri dish containing
xylene. The glass it has to be covered because the
xylene evaporates quickly.
Fig. 4.43. Cracked slide.

69
Fig. 4.48. Storing slides in prefabricated card-
board boxes.
Fig. 4.49. Storing the slides in boxes protects them from dust. Changing the arrangement is hardly
possible. A digital classication system is needed to nd slides.
Fig. 4.46. Final storage of slides on the aluminum
plates in special shelves. Changing the arrange-
ment is always possible.
Fig. 4.47. Storing slides in prefabricated plastic
boxes.
Fig. 4.45. Storing slides in prefabricated alumi-
num plates.

70
4.16 Digital classicaon of slides
4.17 Processing me
When working with wood anatomical slides it is important to make a classificatio
of all characteristics visible on the section. Basically, there is a need to document
the following things:
» number of slide,
» species,
» family,
» collected part of plant,
» life form,
» plant height,
» site characteristics,
» vegetation zone,
» location,
» altitude,
» geographical coordinates,
» collection date,
and possibly information on
» sample provenance,
» special features,
» distribution, etc…
Recommended for anatomical classification are
» the IAWA system for softwoods (conifers) (Richter et al. 2004),
» the IAWA system for hardwoods (Wheeler et al. 1989), or
» the system for dicotyledonous herbs, shrubs and trees
(Schweingruber et al. 2011).
It is difficu to give any exact information about the time needed to process one
sample. The following times are estimates based on own experience over a longer
time.
The processing times for Astrablue/Safranin stained permanent slides for an expe-
rienced person are:
Cutting:
Cross sections of branches: approx. 20–30 sections / hour
Cross, tangential and radial sections: approx. 8 samples (3 sections each) / hour
Cross sections of difficu samples: approx. 5–10 sections / hour
Staining and embedding:
When working in a line as described in section 4.13: 10 slides / hour
Fixation and additional staining with Picric-Anilinblue takes 2–3 minutes longer per
slide.

71
5. Preparaon errors
Cutting radial sections requires a very precise orientation of the sample in relation
to the cutting direction (see section 3.2). A good orientation can be achieved when
concentrating on the ray orientation of the sample. The rays have to be placed par-
allel to the cutting direction (Fig. 5.1). As a result the rays in the micro section will
be quite long and clearly visible (Fig. 5.2). In case the ray orientation is not parallel,
but oblique to the cutting direction, the rays in the micro section appear short (Fig
5.3) and specifi features will not be visible in most parts.
Fig. 5.2. Perfect radial split reveals long sections
of rays.
◂ Fig. 5.1. Splitting parallel (left
side) and oblique (right side) to
rays.
▾
Fig. 5.3. Oblique splits reveals only short sections
of rays. The rays also appear short when rays are
radially bent within the stem.
ray orientation
direction of cutting
▾
▾
▾
▾
1 mm 1 mm

72
Fig. 5.6. Slides with inconsistent thickness are a result of blades too loosely fixed in the knife holder
of the microtome.
Fig. 5.5. Traces on the slide of are a result of a
blade with notches. The damage is irreparable.
Fig. 5.4. Traces on the surface of the sample are
the result of blades with notches. Change the
blade.
Straight lines or even grooves on surfaces and sections are a result of notches in
the blade (Figs. 5.4 and 5.5). If such traces appear on surfaces, change the blade or
move the blade in the holder. Inconsistent thickness of sections is a consequence
of not perfectly stabilized blades (Fig. 5.6), or potentially the wood being too hard
for the blade. Irregularly-formed secondary walls in the earlywood of conifers indi-
cate a dull blade or a badly guided blade (Fig. 5.7).
Trapped air bubbles in the slides appear dark and are extremely disturbing (Figs.
5.8–5.10). Large air bubbles can be massaged out from under the slide cover before
the slide is dried. Slight warming of a dried slide may expand them so that they
disappear.
Some irregularities can be cleaned in the pictures with Adobe Photoshop, or other
image processing software.
250 µm

73
Fig. 5.8. Air in a large part of the slide. Put the
slide in xylene for a day and cover the section
again with sucient Canada balsam.
Fig. 5.10. Air bubbles in cork cells. Thee bubbles
remain in the cells even after heating.
Fig. 5.12. Holes can be old saw marks.
Fig. 5.7. Irregularly-formed secondary walls in the
earlywood of a conifer as a result of a dull knife.
The secondary wall is split off the primary wall.
The damage is irreparable (see section 3.6.2). To
avoid, use a new blade and non-Newtonian uid.
Fig. 5.9. Air bubble in a ray cell. Heat the slide for
a few seconds until the air bubbles escape and
press the cover glass with a magnet on an iron
plate until the slide is cold (see section 4.11).
Fig. 5.11. Small dots (water droplets) are a result
of insufficien dehydration. Put the slide in xylene
for a day, go back to the dehydration process to
ethanol 96%, ethanol absolute and xylene. If the
xylene runs clear, water is no longer present.
250 µm
250 µm
250 µm
25 µm
100 µm 1 mm

74
6. Microscopic observaon
6.1 Polarized light
New student microscopes with magnification of 40x, 100x, 200x, 400x (and
1000x) can be used for most stem anatomical studies. Extremely informative is the
use of polarizing filters No matter which type of microscope you use, the filter
are always placed above and below the slide. You can switch between normal and
polarizing light by rotating one of the filter (Figs. 6.1–6.4).
Polarized light can be used to differentiat types of cell walls (S1, S2, S3) based on
the orientation of their micro fibrils All cell walls consist of more or less net-like,
unstructured micro fibrils primary (S1) and tertiary (S3) cell walls, as well as cell
walls of parenchyma cells disappear in polarized light. On the contrary, secondary
cell walls (S2), consisting of more structured cell walls, more or less oriented in the
same direction, light up in polarized light (Figs. 6.5–6.8).
Fig. 6.2. Placing polarizing filters on top of the
light source and between the objectives and the
oculars. If the top part of the microscope is not
removable, place the lter directly on the slide.
Fig. 6.1. Student microscope with a removable
top part.
Fig. 6.4. Rotating the polarizing filter on the light
source.
Fig. 6.3. Covering the polarizing filter with the top
piece of the microscope.

75
Fig. 6.6. E ect of polarized light in a liana stem
(Vitis vinifera): Lignified cells are red, unlignified
cells are bluish or disappear. Crystals are white.
Fig. 6.8. E ect of polarized light in a stem of a tiny
herb (Cerastium semidecandrum): Cells with sec-
ondary walls in the xylem appear light. Peripheric
cortex cells have lignified secondary walls and
appear red.
Fig. 6.5. E ect of normal light in a liana stem (Vitis
vinifera): Lignified parts are red, unlignified parts
are blue.
Fig. 6.7. E ect of normal light in a stem of a tiny
herb (Cerastium semidecandrum): Lignified parts
are red, unlignied parts are blue.
250 µm
250 µm
250 µm
250 µm

76
6.2 Measuring cell dimensions
6.3 Photography
Measuring cell dimensions, such as cell diameters or vessel lengths, is possible with
a scale integrated in the ocular of the microscope. With this scale, you can directly
see and document the relative dimensions of the cell types and document it. For
measuring cell dimensions in millimeters or microns, the scale in the ocular has to
be calibrated with a stage micrometer.
Place the scale under the microscope and compare the dimension of the eyepiece
micrometer, and calibrate it for all magnification available (Figs. 6.9 and 6.10).
Cell dimensions in photographs can be calibrated with photographs of the stage
micrometer.
Modern microphotographic systems are equipped with automatic measuring sys-
tems.
All companies dealing with microscopes provide sophisticated photographic sys-
tems. Since all of them have their own specificatio we will not describe them here.
Please contact the company. If you want to document some observations without
any intention of publishing it, it is possible to take a photograph with any digital
camera through the eyepiece of the microscope.
Fig. 6.10. Comparing the scale of the eyepiece
micrometer with the metric scale of the stage
micrometer The bent lines represent the border
of the eld of view in the microscope.
Fig. 6.9. Stage micrometer with a resolution of 10
microns.

77
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78
Acknowledgements
We thank Jodi Axelson for assisting with editing this booklet; Cornelia Krause,
Achim Bräuning and Dieter Eckstein for their helpful comments on a firs draft of
the booklet and last but not least we are grateful to all participants of the wood
anatomy course in the last years who directly or indirectly helped developing the
idea for this booklet.