Plant Form and Function
Specimen Preparation Techniques
Creating Free Hand Sections of Plant Specimens
OVERVIEW
The study of the substructure of most plant organs requires
cutting sections. The most simple way to prepare them is free-hand
with a razor blade. Plant tissues offer particularly good
materials for free-hand sections because they are, in a way,
“pre-embedded” in a cell wall (i.e. their contents
stay put more readily than those in cells of animals because
the cell membrane is protected by the cell wall). This advantage
of sectioning plant material in comparison to animal tissues
was recognized early on by microscopists, and by the 1800’s,
free-hand sectioning had been developed to a fine art by European
Botanists. Indeed, in the 1880’s one could refer to
detailed instructions on making free hand sections of pollen
grains in the popular literature! The most detailed instructions
for making good hand sections have readily available in the
European Botanical literature. However, hand-sectioning is
not as commonly used in North America – either for teaching
or research.
Despite widely available literature in English, several generations
of students of botany in North America have not learned this
simple and very effective technique for preparing scientifically
valuable specimens for studying the substructure of plants.
Actually, this has been a handicap to botanists who are not
plant anatomy specialists, but who none-the-less frequently
have questions about the structure or organ they are working
with. Unfortunately these questions often go unanswered or
unasked because the botanist does not have the time or expertise
to prepare sections by more intensive methods of embedding
followed by sectioning with a microtome.
As Lindauer (1972) remarked, hand sections are NOT inferior
substitutes for specimens embedded and prepared using a microtome.
In fact they are of value in their own right as preparations
for study. Thin sections observed at high resolution can yield
a surprising amount of information – particularly about
cell wall structure.
But there are many advantages to making and using hand sections.
You know exactly where the section was taken from in the intact
plant organ and therefore have a much better perspective for
mentally reconstructing the three dimensional plant structure.
Moreover, observations of cells and tissues are not distorted
by the fixation process, or the shrinkage that is inevitable
in the dehydration and embedding processes. And when a polychromatic
dye, such as toluidine-blue-0 is used, one can learn not only
about the anatomy of the tissue, but also about the nature
and distribution of some of the large macromolecules present
in the specimen. Apart from these research benefits, many
hand-sections stained with toluidine-blue-0 are exceedingly
colorful and aesthetically pleasing due to the wide range
of polychromasy and because natural pigments such as chlorophyll
and anthocyanins also are present.
You will work in a small group (2-3) people to accomplish
the following objectives. Please note that you should be keeping
your notes in your lab notebook individually. See more about
the lab notebook under the assignments listed on the syllabus.
OBJECTIVES
1. To learn and master the technique of free-hand sectioning
2. Prepare thin sections of a variety of plant materials
3. To interpret the anatomical features made from free-hand
sections
4. To appreciate this technique as a valuable tool for studying
the substructure of plants.
LAB NOTEBOOK
Prepare sketches/drawings from your specimens and label the
cells you are looking at. Capture 2 of your best images digitally,
print them out to be affixed in your notebook, and save the
digital images according to the format for this course.
INTRODUCTION AND BACKGROUND INFORMATION:
For cutting herbaceous plant tissues, the best blades for
the job are “Blue Blades” manufactured by Gillette
Co. To make the best possible sections it is imperative that
new blades are used, and that only a few sections are made
with each part of a blade edge. For making thin sections of
woody tissues, it is better to use a single edge blade such
as the “Gem” brand.
To learn the free-hand sectioning method, you will start
with fresh herbaceous material. Stems are the easiest material
to work with when you are learning this technique. With practice,
most people will soon be able to cut difficult preparations
(e.g., longitudinal sections of roots; cross sections of leaves).
Regardless of the type of section you are making, it is important
that both the edge of the razor and the face of the tissue
being cut are moistened with water before sectioning. It is
almost impossible to cut thin sections of very hard tissues
like seeds. But even a thick section of a hard tissue can
yield a lot of anatomical detail with a dissecting microscope.
LAB INVESTIGATION:
METHODS
Supplies
1. Supply of degreased double-edged razor blades (preferably
Gillette Blue Blades). Degrease in xylene, store in 100% ethanol
2. Small metal spatulas for handling sections
3. Three four-inch watch glasses (or the equivalent). Two
will be used for soaking cut sections, and one for rinsing
stained sections.
4. One two-inch watch glass (or equivalent) for staining.
5. Paper towels
6. Clean slides and coverslips
7. 0.05% aqueous solution of toluidine-blue-O
8. hand lens
9. cheap cosmetic mirror
10. desk lamp or equivalent source of directed white light
11. supply of fresh carrots
SAFETY INFORMATION
Caution: Broken pipettes, coverslips and slides, and razor
blades are very sharp. Dispose of them in the “sharps”
container and NOT the regular trash can.
Procedure:
Either work near a sink or have two bowels available –
one for wetting fingers and tissues, and the other one for
discarding unwanted sections and stain solution.
A. Thin-Hand Sectioning
1. Hold tissue with thumb and first two fingers of one hand.
Position the tissue so that the region you want to cut is
just beyond the edge of the tops of the thumb and forefinger.
It is important not to grip the tissue and blade too tightly
– remain relaxed so your hands do not cramp.
2. Thoroughly wet the blade, tissue and fingers with water
from the tap or the bowl (unless there is some special reason
for sectioning dry tissue).
3. Make the first cut quickly and smoothly to produce a fresh
surface in the desired plane. Use no more than the first 25%
of the razor edge to make the first cut – discard the
section it produces. Use the remainder of the edge for the
rest of the sections you will keep.
4. Make each subsequent cut slowly and deliberately –
resting the blade in the tip of the thumb and the top or side
of the forefinger. Use a sliding action so that the blade
moves through the tissue at an angle (DO NOT USE A CHOPPING
ACTION). Control is usually better if the cut is made toward
the body but see what works best for you. With favorable material
quite a lot of sections can be made – even serial sections
– with one edge of the blade. The act of cutting is
controlled as much by touch as by sight. When the edge of
the blade is sharp, the thinness of the section you are cutting
can be sensed in advance by the resistance you feel with the
initial touch of the blade. Thicker sections will have a higher
resistance to the blade than a thinner section. For some tissues,
you may find it easier to make the cuts under a dissecting
scope.
5. Do not let the section(s) dry out. If there is difficulty
in making thin sections, do not hesitate to cut off another
thick section so that you can begin your work again from a
fresh smooth section. Do not be dismayed if your first efforts
produce a lot of incomplete sections or uneven sections. Also,
do not use pieces of plants that are too large in area. Instead
use sectors of the stem or root – you can still interpret
a lot of information from small sectors of a plant organ that
has a large area.
6. Remove sections from the razor blade by floating them off
into one of the two watch glasses containing tap water. The
edge of the blade will be dulled if you rub it on the edge
of the watch glass to remove the sections. Use a small spatula
to detach sections that adhere to the edge and for manipulating
sections during subsequent sectioning. Unstained sections
can be seen more easily if a watch glass is placed over a
black piece of paper or other dark background.
7. Allow sections to soak for at least two minutes to remove
debris from cut cells.
8. Rewet tissue, blade and fingers and continue sectioning
until sufficient material sections have been cut or the razor
blade gets dull.
9. Change the razor blade edge if resistance seems to increase
when cutting (herbaceous tissue will yield almost no resistance
to a sharp blade). As a blade dulls, an initial sensation
of greater resistance is followed very quickly by tearing
of fibrous vascular tissue, crushing the epidermis, and failure
to completely sever the section from the uncut surface. It
will be obvious that it is time to change the blade by this
point!
Special technique for cutting thin sections through leaves:
Moisten two razor blades and grip them tightly together. Use
the pair to slice into a fresh leaf spread on a paper towel.
Sections will lie between the blades. Sometimes it can be
helpful to use a piece of thin paper as a spacer between the
gripped edges of the blade.
10. Transfer selected sections to the small watch glass with
the toluidine-blue-0 stain. Stain for about 30 – 60
seconds. The optimal time will vary and depends on the material
being stained. Check the sections being stained under a dissecting
scope to see if all the tissues have achieved a satisfactory
intensity of staining. If the stain in the watch glass begins
to evaporate at the edge of the watch glass, throw it out
and wash the glass before adding new stain. Precipitated stain
will not wash off new sections. As a helpful option, hold
the watch glass over a mirror and adjust the position of a
lamp so that its light reflects upward through the staining
solution. In this way you can watch the process of the staining
reaction even in the intensely colored dye!
11. To rinse the sections, transfer to the third watch glass
and move them about gently in the tap water for about 60 seconds.
The rinse time is not critical since the stain does not bleed
rapidly from the tissue (at least as long as the pH of the
tap water is about 4.0). View these sections with the dissecting
scope and select the best ones for mounting on a microscope
slide. Discard the unwanted sections and replace the wash
water.
12. Mount sections in clean tap water (see instructions for
preparing the slide below).
Interpreting the Color of Macromolecules: Cells impregnated
with lignin or other phenols, and tannin-rich vacuoles stain
green, turquoise, or bright blue. Polysaccharides rich in
carboxyl groups or sulfate groups stain pink or reddish purple.
DNA stains green and purplish blue. RNA stains purple or purplish
blue (however these materials usually are lost from the hand
sections of fresh material).
B. Mounting Specimen on Microscope Slide
1. Select a clean slide and coverslip. Coverslips often are
dusty as they come from a box. Breathe on the coverslip and
polish it up by rubbing with a tissue. Once polished, handle
it only by its edges to avoid finger prints. Set the clean
slide and coverslip on a clean, dry paper towel – rest
the coverslip against the slide at an angle so you can pick
it up again without getting finger prints on it.
2. Put a drop of clean tap water about 5 mm in diameter on
the center of the slide.
3. Transfer the thin sections to the drop. Do not mount more
sections than will fit in about a 1/3 the area of the coverslip.
Once the sections are in position, hold the coverslip by its
edge at an acute angle to the slide and touch the edge of
the coverslip to one edge of the drop. Lower the coverslip
SLOWLY, supporting it as you do so with the spatula. It is
essential that you do not let it drop suddenly or you may
knock the prepared sections right off the slide! Moreover
you are likely to trap air bubbles if you drop the coverslip
onto the sections and water drop. You can avoid trapping air
bubbles by allowing the advancing front of the water film
to move slowly over the surface of the sections as the coverslip
is lowered.
4. Make certain that both the top and bottom of the slide
and the top of the coverslip are dry before placing them on
the stage of the microscope. Use a paper wick to remove water
from the top of the coverslip. Be particularly careful not
to wet the microscope stage to avoid damaging the condenser
lens and having your slide stick to the stage so that it is
difficult to move.
5. It is best to use water for all temporary mounts. You can
put a ring of nail polish or freshly melted petroleum jelly
around the coverslip when it is necessary to keep the preparation
for a few days. You can also make optically superior temporary
mounts that will still show the full range of colors if 1)
the stained sections are allowed to dry ever so briefly on
a moist paper towel and 2) then the sections are evacuated
and later mounted in paraffin oil.
C. Common Problems and Fixing Them
1. Coverslip is afloat: Too much mounting water. Remove excess
with absorbent paper applied to the edge of the coverslip.
2. Some sections still afloat even after the mounting water
has been absorbed: One of the sections is too thick and holding
the coverslip up off the other sections. Break the seal on
the preparation by gently lifting the coverslip off of the
slide. The edge of a dulled razor blade works well for this
task. NOTE that the scattered drops that form on a slide when
the seal is broken make it nearly impossible to remount a
coverslip without bubble entrapment so it generally is best
to start the mounting process all over again. Return all the
sections back to the water dish. Either discard the thick
section or mount it separately (thick sections can be valuable
for information in many cases). Dry both the slide and coverslip
before remounting.
3. Meniscus is evident at one edge of the slide: Too little
mounting water. Add more to the edge of the coverslip –
one or two little drops should be sufficient.
4. Coverslip slopes at one side and it is impossible to remove
the meniscus with additional water: You have a thick wedge-shaped
section on the slide. Start over with a better section.
5. Bubbles are present within the tissue even when all reasonable
precautions have been taken: This is a common problem in highly
aerated tissues such as stems of aquatic plants, rushes, sedges
and swamp grasses. This is likely caused by the highly hydrophobic
nature of the surface of the internal air cavities in these
plants. Alternative 1: Place the sections you made into freshly
boiled water and cooled water prior to mounting. Alternative
2: evacuate the air in the tissues in 10% aqueous ethanol
to improve penetration of the liquid and to allow air to escape.
You need to do this treatment prior to staining as the ethanol
would otherwise destroy the color.
6. Part of the section is seen as a surface view: This is
frequently common with leaf sections which can fold over –
especially grasses. Sections too thick relative to width have
a tendency to come to rest on their surfaces (epidermis facing
up). To avoid this cut thinner sections. Alternatively, cut
sections to include the mid-rib which will help hold the rest
of the leaf section flat.
7. Sections always flush to the edge of the coverslip during
mounting: This can be do to the use of too much mounting water
so use less. Sometimes it happens when the sections contain
too much air so evacuate them as described in 6 above. However,
sometimes it is due to surface tension effects that arise
from the influence of hydrophobic coverslips or the cutinized
epidermis of section surfaces. This latter case can be an
infuriating problem. A coverslip moistened by breathing on
it can help. Alternatively add a little tiny drip of saliva
to the water on the mount – stir it with a toothpick
before adding the sections and coverslip.
DISPOSAL - CLEAN UP
Wash and dry all slides when you are done and place them in
the separate glass containers near the sink! Turn off your
microscope, cover it and put it back into the microscope cabinet.
Clean and dry your workspace before leaving lab.
RESULTS
Include the following information in your lab notebook: Drawings,
sketches, printouts of your best images, and list of file
names made according to the course format.
ACKNOWLEDGEMENTS AND REFERENCES
These instructions on preparing free-hand sections were modified
from material developed by Dr. Greg Brown, Botany Department,
University of Wyoming.
Top |
