Experimental Anatomy of Plant Development Laboratory 1

Laboratory Techniques

Purpose: For the student to become familiar with the sectioning and staining techniques used with plant materials and comfortable with the use of the microscopes. These skills are foundational to the course and will be used extensively throughout. The materials provided will below serve as reference materials for the course. Activity 1. Review rules and techniques for the use of the microscopes. To best accomplish the other goals, students will be divided into groups and will rotate through the following activities: Activity 2. Sectioning and staining. A variety of plant materials, stains and sectioning materials have been provided. Students should explore the use of these materials to perform the sectioning techniques below. Perform each technique listed under “Sectioning Techniques” to make each “Plane of Section” possible, stain the sections with a variety of stains provided and make drawings of your work in your lab notebook.

Plant material: maize coleoptiles, Garlic (Alium) root tips, Coleus stems, petioles and leaf laminae. Sectioning and mounting materials: Plain glass microscope slides, cover glasses, single-edge razor blades, fine forceps, fine dissecting needles, pipette, soft watercolor brush, 3H pencil, drawing paper.

Activity 3. Use of the vibratome. Activity 4. Use of the fluorescent microscope.

I. Anatomical Drawings When you make anatomical drawings, you develop several useful skills including the ability to interpret complex information, identify diagnostic features that distinguish among similar structures, and represent and communicate this information in visual form. The purposes of a drawing are to convey information, and to provide you with a record of what you have seen for future reference.. To best accomplish this, use the following steps when planning your drawings:

1. Select the magnification and field according to what you are asked to illustrate. Given the same prepared slide, you might be asked to illustrate a) a cell type, b) the arrangement of cells within a tissue, or c) an arrangement of tissues within an organ (i.e., a diagram). The resulting drawings would be very different.

2. Include details that distinguish the subject from other similar structures (diagnostic

features). Given the assignments above, you drawings might include a) details of the individual cells, b) outline of individual cells with enough detail to distinguish among cell types or c) you may not need to draw individual cells at all. If the point is to show how vascular bundles arranged, you need only outline the boundaries of vascular bundles.

3. Represent form, proportion, and spatial relationships accurately. 4. Use insets when information at more than one level of organization needs to be

conveyed. 5. Label all distinguishing features.

Each drawing should be labeled with the - plant material used - type of sectioning and plane, if appropriate - fixation and staining - magnification and microscopy used

II. Sectioning techniques A. Planes of sectioning Solid material should be sectioned in several planes in order to discover the distribution of the various tissues within it. The complete investigation of axial structures, such as stems or roots, normally requires a transverse (cross) section at one, or more, levels; and radial longitudinal, and tangential longitudinal sections at different depths from the surface to the center. Foliar structures generally require transverse, and paradermal sections; and vertical longitudinal sections may occasionally be necessary. B. Treatment of sections When cut, sections of fresh material should be placed in water; and those of preserved materials into alcohol. However, it may be necessary to place sections of some fresh materials into alcohol to get ride of the air within them. In order to avoid unnecessary handling of the sections, all treatments should be carried out on the slides on which the sections will ultimately be mounted. When the liquid surrounding the section is to be changed, merely add a pool of the new liquid at one end of the slide. Pull the section into the new liquid with a dissecting needle, and pour the old liquid off the other end of the slide. Some techniques may require especially long rinses. In those cases, the sections may be rinsed more thoroughly in a shallow dish. The best instrument for moving sections when using a dish is a small watercolor brush. Sections of preserved material should be mounted in a drop of dilute glycerin. Sections of fresh material should be mounted in water or dilute glycerin. It is not necessary to spend a great deal of time trying to make a perfect mount. A few air bubbles in the final mount are generally not a problem, as long as they can be recognized and don’t obscure the areas of interest. The size and frequency of such bubbles may be reduced by mounting sections in water containing a wetting agent such as glycerin or detergent (“wet” water). Clearing This technique is especially useful for examination of the intact vascular systems of leaves and floral parts. However, it may also be resorted to as a means of making thick freehand sections more transparent. The easiest method is to clear tissue by incubating it in EtOH to remove hydrophobic pigments including chlorophyll. This may take several hours.

Many such plant parts of either fresh, preserved or dry materials may be cleared sufficiently by warming them in chloral hydrate. Others may require treatment with NaOH as follows: Clear in 5% NaOH in a Petri dish in an oven (the time varies from one to several day depending on the material). Wash 3 to 5 times in distilled water carefully with a

pipette. If more clearing appears to be necessary, use a saturated aqueous solution of chloral hydrate for 24 hours. Wash again in distilled water. Staining Two or three drops of stain can be transferred by pipette to a spot plate, ceramic ring slide, or depression slide. Transfer the specimen by forceps to the drop of dye and wait 1-several minutes (depending on the dye concentration). Transfer the specimen to a drop of solvent (water or dilute ethanol) of the same concentration used to make up the dye. Let stand a minute. Alternatively, sections can be placed in gooch crucibles, and moved among small beakers, syracuse watch glasses, or coplin jars if production staining is necessary. Mounting Take a clean dry microscope slide and place on it a small meniscus of calcium chloride solution. Transfer a stained specimen to the calcium chloride and wait until the specimen mixes well with the salt solution. Drop a clean cover slip onto the specimen. C. Sectioning techniques. Epidermal Peels The superficial tissues of many plant parts (especially leaves) may be peeled away in strips thin enough for microscopic examination. To make such a peel, break or cut the surface of the plant apart. Then, grip the epidermis with forceps at one of the cut edges, and pull the outer tissue layer back away from the cut. The resulting epidermal peel should be mounted in water containing a wetting agent; or in alcohol if it is very hydrophobic. Macerations The three-dimensional form of a cell is most easily seen when the cell is separated from the surrounding cells of the tissue. Macerating fluids accomplish this through a hydrolysis of the middle lamella. The following method is a gentle, but effective, technique: Cut small pieces of the tissue into a mixture of 1 part Hydrogen peroxide, 4 parts distilled water, and 5 parts glacial acetic acid. Cook the mixture in a 56-60 degree oven for 24 hours. If further macerating is needed, replace the old fluid with a fresh mixture and cook the tissues for another 24 hours. Repeat the process until the material is mostly colorless, and may be easily teased apart with a dissecting probe. When the maceration is

complete, rinse the tissues in water in an uncovered container. Stain in 0.25% Safranin in water and mount in dilute glycerin. Squashes Material can be squashed on a slide for cytological examination. This technique is most often used for chromosome counts and examination of mitotic structures. Dissect away non-meristematic tissues, chop the meristem with a scalpel, place a coverslip over the tissue, place paper towels on the coverslip and apply vertical pressure through your thumb. Free-Hand Sectioning Material should be kept moist while sectioning. Liquid should be kept on the razor blade, so that the sections float as they are cut. In general, it is inadvisable to take particular care over individual sections. Better results are usually obtained by cutting a large number of slices rapidly, and sorting out the best ones. Sections of uniform thinness are usually not necessary. Wedge-shaped slices which taper from opaque, overly thick margins to ultra-thin edges will sow useable areas of the proper thickness. When cutting longitudinal sections, it is important to use a short piece of material not much longer than wide. It is impossible to cut satisfactory longitudinal sections of any considerable length by the freehand technique. Flexible structures, such as leaves, require some support during sectioning. Many leaves will yield good transverse, and vertical longitudinal sections if rolled or folded so that 10 or more thicknesses are cut at each stroke. If some extra support is necessary, the material may be inserted into the cut and of a young carrot which has been pickled in alcohol. The material and the surrounding carrot tissue are then cut at the same time. This technique should produce results superior to those of the more classical elderberry pith method. To obtain paradermal sections of a leaf, bend it over a finger and cut small slices off the curved surface. Sections of dry material should first be soaked in alcohol or hot water to soften it and remove air from the cells. Mount sections on clean slide in a drop of water. To apply the coverslip, hold it at an angle and touch the water drop with one edge. Lower the coverslip slowly to avoid air bubbles. Semi permanent mounts can be made by fixing tissue in phosphate-buffered glutaraldehye, mounting in glycerol jelly and sealing the coverslip with nail polish.

Vibratome Sectioning: An Alternative to Paraffin Methods The vibratome is an instrument that is similar to a microtome but uses a vibrating razor blade to cut through tissue. The vibration amplitude, the speed, and the angle of the blade can all be controlled. Fixed or fresh tissue pieces are embedded in low gelling temperature agarose. (Some have had success without using the agarose to embed.) The resulting agarose block containing the tissue piece is then glued to a metal block and sectioned while submerged in a water or buffer bath. Individual sections are then collected with a fine brush and transferred to slides of multiwell plates for staining. For most plant specimens, stems are the easiest to section both transversely and longitudinally. Leaf cross sections are relatively easy to cut, although sometimes the blade of the leaf tends to rotate as it is cut, resulting in imperfect cross sections. Roots tend to be the most difficult organs to section due to their wood texture. Softening root tissue or using roots grown on tissue culture medium may be preferable. All of the parameters associated with vibratome sectioning, including fixation and cutting settings, depend greatly on the source of tissue being sectioned and will vary with different species, organs, and tissues. It is therefore necessary to optimize these parameters for your own specimens. Fixation conditions vary depending on tissue type and the target antigen so it is best to consult the literature to obtain some starting ideas for your system. After fixation, the tissue pieces are washed briefly in 50 mM PIPES buffer (3 changes over a period of 30 minutes). Tissues are then embedded in 5.0% low gelling temperature agarose (Sigma Type XI, A-3038) and allowed to cool until solid. Small 5 ml plastic disposable microbeakers can be used as molds. Several tissue pieces can be embedded in each mold. It is important to have the tissue surrounded by agarose on all sides when it is finally solidified. A small block of agarose containing a tissue piece is cut out and trimmed in such a way as to yield proper final orientation of the tissue when placed on the metal block holder. After the fine trimming is completed, the agarose block is glued onto a metal block holder with super glue, allowed to dry for about 5 min, and then sectioned on the vibratome. See vibratome demonstration for more details on vibratome sectioning.

D. Stains, Reagents and other visualization techniques The student should become familiar with the use of the following simple techniques and

apply them in the study of laboratory materials: “Wet” water Used for preparing wet mounts of specimens for general observations with a minimum of trapped air. Dilute glycerin Used for preparing wet mounts for general observations when rapid drying of the mount is undesirable. Chloral hydrate Used to clear whole structures or sections which are not otherwise transparent. Mount the specimen in a few drops of chloral hydrate under a cover glass. Warm the mount over alcohol lamp until it seems more transparent. Do not allow the fluid to boil violently. Observe the mount directly, or rinse and remount in glycerin. This treatment renders cell walls visible, but removes most cell contents. Starch grains are dissolved, but crystals of calcium oxalate is not. Lignin also remains and tests positive with phloroglucinol. Aniline blue Used to stain callose in the phloem; and to stain nuclei and nucleoli. Place fresh hand sections in IKI for about a minute, then rinse in water. Stain in aniline blue for about 5 minutes, then briefly rinse in IKI. Mount in water or dilute glycerin. (IKI may be omitted if too much starch is present causing blackening of the tissues.) Callose and nuclei stain blue; starch grains blue-black; cell walls yellow-brown. Safranin A general stain for cell walls, it is especially useful for staining

macerated tissues. Stain for a minute in Safranin, then rinse thoroughly with water. Mount in water or dilute glycerin.

Toluidine blue A metachromatic dye which gives a multicolored stain in fresh hand sections. Stain in toluidine blue for about 10 seconds to one minute, then rinse thoroughly in water. Mount in water. Carboxylated polysaccharides such as pectic acids stain pinkish purple. Macromolecules with free phosphate groups such as nucleic acids stain purple or greenish blue. Polyphenolic compounds such as lignin and tannis stain

green, greenish blue, or bright blue. Hydroxylated polysaccharides such as cellulose and starch are unstained. Thus, cells with primary walls (parenchyma, sieve elements, collenchyma) stain pinkish purple due to the polyuronides (pectic compounds) in their walls. Lignified secondary walls of sclerenchyma, xylem elements, and phloem fibers will stain green to blue, as will some other unlignified cells with phenols in their primary walls. Phloem fibers stain bright blue, xylem elements a greenish color. Cambial cells and some phloem cells are almost unstained. Polarized Light Some of the components of plant cells are wholly, or partly

crystalline. Any such structures may be detected by the use of polarizing filters. One filter (polarizer) is placed between the microscope illuminator and the specimen to polarize the light. Another polarizing filter (the analyzer) is placed between the specimen and the viewer’s eye. One of the polarizing filters is then rotated, causing the field of view to alternately darken and brighten. When the field is dark, the two polarizing filters are crossed and screen out all light passing through amorphous (isotropic) structures. A crystalline structure in the specimen will rotate some of the polarized light reaching it from the polarizer, allowing light to pass through the analyzer to the viewer’s eye when the tow polarizing filters are crossed. Thus crystalline structures will appear to shine when viewed with crossed polarizing filters.

Micro chemical Tests Calcium carbonate Crystals composed of calcium carbonate will dissolve when wet

mounted in 33-1/3% acetic acid. Calcium oxalate crystals will not. Cellulose Soak fresh sections in IKI for at least 15 minutes. DO NOT COVER THE TISSUES DURING THIS SOAKING PERIOD. Drain off the excess IKI. Then add a drop or two of 65% sulphuric acid (H2SO4) directly to the sections, and apply a cover glass. Cellulose will become blue in color. Lignin will appear yellow to orange in color. Note: This test will ultimately destroy the specimen. Make all other necessary observations before adding the acid.

BEWARE OF THE ACID. DON’T ALLOW ANY TO CONTACT THE MICROSCOPE.

Lignin Prepare a wet mount of sectioned or cleared material in phloroglucinol in HCL. Lignin will stain red. BEWARE OF ACID. DON’T ALLOW ANY TO CONTACT THE MICROSCOPE. Lipids Place fresh hand sections or epidermal peels in 50% EtOH for about a minute. Stain with sudan IV for about 5 to 20 minutes, then rinse in 50% etOH for about a minute. Mount the stained sections in glycerin. Waxes, fats, and oils will stain red-orange. Nucleic Acids DAPI (4'-6-Diamidino-2-phenylindole) is a very easy method for

detecting mitotic figures if a fluorescence microscope with a DAPI filter cube is available. It is excited by UV light and fluoresces blue. It forms fluorescent complexes with natural double-stranded DNA, showing a fluorescence specificity for AT, AU and IC clusters. Because of this property DAPI is a useful tool in various cytochemical investigations. When DAPI binds to DNA, its fluorescence is strongly enhanced, but there is also evidence that DAPI binds to the minor groove, stabilized by hydrogen bonds between DAPI and acceptor groups of AT, AU and IC base pairs. Use at 1 µg/ml in buffer. Stain for 5 min. It is not necessary to wash after staining as the dye only fluoresces when bound to DNA. Fix tissues in 3:1 ethanol: acetic acid. Tissues that lack a cuticle work best (i.e. roots).

The Feulgen technique selectively stains DNA. The reaction consits of

two steps: Fixed material must be hydrolyzed for 8-12 min with 1N HCl. Hydrolysis times may vary for different tissues and will affect the intensity of staining. Hydrolysis can be done in 1N HC1 at 60° C for 10 min (8 and 12 min time points should also be tested for optimal staining) or at room temperature in 5N HC1 for 30 — 60 min. Afterwards, the material is immediately transferred into Schiff's reagent at room temperature (for at least 30 min or until the tissue stains deep purple). The material is then squashed in acetocarmine or aceto-orcein. It is recommended that the material be analyzed the same day, however, it can be kept at 4 C for a several days if necessary. (Acid hydrolysis removes purin bases from the DNA, thereby unmasking free aldehyde groups. The aldehyde groups then react with Schiff's reagent, which results in the purple staining. RNA is not hydrolyzed by the HCl treatment and, thus, the reaction is DNA-specific.)

Immunolocalization of PCNA - PCNA, proliferating cell nuclear antigen, is required for DNA replication.

Pectin Place hand sections in freshly prepared ruthenium red for 5 minutes or longer. Mount the stained sections in water. Pectic compounds will stain dark pink. Proteins Stain fresh hand sections in picric acid for several minutes, then rinse thoroughly with 100% EtOH. Mount in 100% EtOH. Proteins will stain yellow. Starch Stain fresh sectioned or peeled material in IKI for a few minutes. Rinse and mount with water. Starch grains will stain blue-black. The brown staining of some organelles and inclusions do not constitute a positive test for starch. Tannins Prepare a wet mount of fresh hand sections in ferric chloride or ferric sulfate. Tannins will form blue or green-black precipitates with these ferric salts.

NOTE: These reagents contain hydrochloric acid. BEWARE OF THE ACID. DON’T ALLOW ANY TO CONTACT THE MICROSCOPE.

PLANT TISSUE STAIN RESPONSES WITH CALCIUM CHLORIDE USED AS MOUNTANT DYE LIGNIN CELLULOSE COLLENCH METACHROMASY DIFFERENTIATION Cresyl viol. Acet. viol blue yell. tan red tan +++ +++ Thionin blue rose tan reddish +++ +++ Toluidine blue blue no no + +++ Methylene blue (1) green; blue no no; pink ++ +++ Hematoxylin (2) red; orange tan; grey red tan +++ +++ Iodine KI yellow rose rose +++ +++ Safranin O red yell. tan yell. tan + +++ Trypan blue no; blue blue blue - ++ Erythrosin no pink pink - ++ Phloxine no pink pink - - Malachite green blue-green no no - ++ Fast Green FCF green no no - +++ Orange G orange tan tan + ++ Celestine blue B no viol. blue viol. blue - ++ Aniline blue blue no no - ++ Schiff’s reagent red pink no no - +++ Methyl green green no; grey no; grey + +++ Chlorazol black no grey blue + ++ Fluorescein yellow yellow yellow - + Aniline blue black no blue blue - + Crystal violet violet; no violet violet - + (1) Dye leaches into mountant; doesn’t hurt image. (2) May fade in 2-4 weeks.