A method for staining pollen tubes in pistil

A quadruple staining procedure has been developed for staining pollen tubes in pistil. The staining mixture is made by adding the following in the order given: lactic acid, 80 ml; 1% aqueous malachite green, 4 ml; 1% aqueous acid fuchsin, 6 ml; 1% aqueous aniline blue, 4 ml; 1% orange G in 50% alcohol, 2 ml; and chloral hydrate, 5 g. Pistils are fixed for 6 hr in modified Carnoy's fluid (absolute alcohol:chloroform:glacial acetic acid 6:4:1), hydrated in descending alcohols, transferred to stain and held there for 24 hr at 45 +/- 2 C. They were then transferred to a clearing and softening fluid containing 78 ml lactic acid, 10 g phenol, 10 g chloral hydrate and 2 ml 1% orange G. The pistils were held there for 24 hr at 45 +/- 2 C, hydrolyzed in the clearing and softening fluid at 58 +/- 1 C for 30 min, then stored in lactic acid for later use or immediately mounted in a drop of medium containing equal parts of lactic acid and glycerol for examination. Pollen tubes are stained dark blue to bluish red and stylar tissue light green to light greenish blue. This stain permits pollen tubes to be traced even up to their entry into the micropyle.     

Differential Staining of Aborted and Nonaborted Pollen

A single staining solution was made by compounding it in the following order (dyes were from British Drug Houses): ethanol, 10 ml; 1% malachite green in 95% ethanol, 1 ml; distilled water, 50 ml; glycerol 25 ml; phenol, 5 gm; chloral hydrate, 5 gm; acid fuchsin 1% in water, 5 ml; orange G, 1% in water 0.5 ml; and glacial acetic acid, 1-4 ml. For best results in differentiation to give green pollen walls and red protoplasm, the staining solution should be acidified with glacial acetic acid. The amount of acid to be added depends upon thickness of the pollen walls: for very thin-walled pollen, 1 ml; for moderately thin walls, 2 ml; and for thick-walled or spiny-walled pollen, 3 ml of acid. For pollen inside non-dehiscent anthers, 4 ml of acid should be used. Staining is hastened by flaming the slide (for loose thin-walled pollen) or by immersing thick-walled pollen or anthers for 24-48 hr at 50 C. In the typical stain, aborted pollen grains are green; nonaborted, red. The method is useful for pollen inside nondehiscent anthers if these are small and not too deeply coloured naturally. The stain is very durable, especially if the coverslips are sealed with param wax. The staining solution will keep well for about a month. It is useful both for angiosperms and gymnosperm microgametes.     

Staining Germinating Pollen and Pollen Tubes

Germinating pollen on stigmas and pollen tubes in styles of Antirrhinum, Brassica, Oenothera, Raphanus, Rosa, solatium and Tagetes spp. were prepared for examination as follows: The styles were fixed in ethyl alcohol-acetic acid 3:1 for 1 hr, and hydrolyzed at 60°C for 5 to 60 min (depending on the species) in 45% acetic acid. The stigma with its attached strand(s) of stigmatoid tissue was then dissected out under a stereoscopic microscope, placed in a few drops of a staining solution made by dissolving 150 mg of safranin O and 20 mg of aniline blue in 25 ml of hot 45% acetic acid. After 5-15 min in this stain, the tissue was placed in a fresh drop of stain on a microscope slide and gently squashed under a cover glass. Because of a gradual precipitation of the aniline blue component, the stain had to be filtered regularly before use. However, a staining solution could be kept at room temperature for several weeks.     

A Versatile Stain for Pollen Fungi, Yeast and Bacteria

A versatile stain has been developed for demonstrating pollen, fungal hyphae and spores, bacteria and yeasts. The mixture is made by compounding in the following order: ethanol, 20 ml; 1% malachite green in 95% ethanol, 2 ml; distilled water, 50 ml; glycerol, 40 ml; acid fuchsin 1% in distilled water, 10 ml; phenol, 5 g and lactic acid, 1-6 ml. A solution has also been formulated to destain overstained pollen mounts. Ideally, aborted pollen grains are stained green and nonaborted ones crimson red. Fungal hyphae and spores take a bluish purple color and host tissues green. Fungi, bacteria and yeasts are stained purple to red. The concentration of lactic acid in the stain mixture plays an important role in the differential staining of pollen. For staining fungi, bacteria and yeasts, the stain has to be acidic, but its concentration is not critical except for bacteria. In the case of pollen, staining can be done in a drop of stain on a slide or in a few drops of stain in a vial. Pollen stained in the vial can be used immediately or stored for later use. Staining is hastened by lightly flaming the slides or by storing at 55±2 C for 24 hr. Bacteria and yeasts are fixed on the slide in the usual manner and then stained. The stock solution is durable, the staining mixture is very stable and the color of the mounted specimens does not fade on prolonged storage. Slides are semipermanent and it is not necessary to ring the coverslip provided 1-2 drops of stain are added if air bubbles appear below the coverslip. The use of differentially stained pollen mounts in image analyzers for automatic counting and recording of aborted and nonaborted pollen is also discussed.     

Aceto-Iron-Haematoxylin-Chloral Hydrate for Chromosome Staining

Aceto-iron-haematoxylin can be used combined with the clearing agent chloral hydrate for the squash method. The stain is prepared by dissolving 2 gm of chloral hydrate in 5 ml of a stock solution of 4% haematoxylin and 1% iron alum in 45% acetic acid, which has been allowed to ripen for 24 hr to 1 wk. Heat must not be used to hasten solution. The material (fixed in 1:3 acetic-alcohol) is put on a slide, the fixative removed and a drop of stain added; if necessary the material is crushed before the cover slip is placed in position. The preparations are now carefully heated until a slight colour change occurs. Squashing needs more pressure than in other techniques. This stain gives best results in zoological and botanical material not requiring hydrolysis, e.g., leucocytes, ascites cells, and cells undergoing spermatogenesis and microsporogenesis. Well-spread and selectively stained mitotic and meiotic figures can be obtained.     

A Protargol Method for Staining Nerve Fibers in Frozen or Celloidin Sections

Extensive experimentation with protargol staining of neurons in celloidin and frozen sections of organs has resulted in the following technic: Fix tissue in 10% aqueous formalin. Cut celloidin sections IS to 25 μ, frozen sections 25 to 40 μ. Place sections for 24 hours in 50% alcohol to which 1% by volume of NH₄OH has been added. Transfer the sections directly into a 1% aqueous solution of protargol, containing 0.2 to 0.3 g. of electrolytic copper foil which has been coated with a 0.5% solution of celloidin, and allow to stand for 6 to 8 hours at 37° C. Caution: In this and the succeeding step the sections must not be allowed to come in contact with the copper. From aqueous protargol, place the sections for 24 to 48 hours at 37° C. directly into a pyridinated solution of alcoholic protargol (1.0% aqueous solution protargol, 50 ml.; 95% alcohol, 50 ml.; pyridine, 0.5 to 2.0 ml.), containing 0.2 to 0.3 g. of coated copper. Rinse briefly in 50% alcohol and reduce 10 min. in an alkaline hydroquinone reducer (H₃BO₃, 1.4 g.; Na₂SO₃, anhydrous, 2.0 g.; hydroquinone, 0.3 g.; distilled water, 85 cc; acetone, 15 ml.). Wash thoroly in water and tone for 10 min. in 0.2% aqueous gold chloride, acidified with acetic acid. Wash in distilled water and reduce for 1 to 3 min. in 2% aqueous oxalic acid. Quickly rinse in distilled water and treat the sections 3 to 5 min. with 5% aqueous Na₂S₂O₃+5H₂O. Wash in water and stain overnight in Einarson's gallocyanin. Wash thoroly in water and place in 5% aqueous phosphotungstic acid for 30 min. From phosphotungstic acid transfer directly to a dilution (stock solution, 20 ml.; distilled water, 30 ml.) of the following stock staining solution: anilin blue, 0.01 g.; fast green FCF, 0.5 g.; orange G, 2.0 g.; distilled water, 92.0 ml.; glacial acetic acid, 8 ml.) and stain for 1 hour. Differentiate with 70% and 95% alcohol; pass the sections thru butyl alcohol and cedar oil; mount.     

Haematoxylin-Phloxine-Alcian Blue-Orange G Differential Staining of Prekeratin, Keratin and Mucin

This is a modification of Kreyberg's stain with Alcian blue 8GS used to stain acid much while phloxine B and orange G stain keratin and prekeratin. Procedure: Dewax formalin-fixed paraffin sections in xylene and hydrate through alcohol. Stain in Mayer's haemalum, 10 min; blue in tap water; wash in distilled water; stain in 1% phloxine, 3 min; wash in running water, 1 min; wash in distilled water; stain in 0.5% aqueous Alcian blue in 0.5 acetic acid, 5 min; wash in distilled water; stain in 0.5% orange G dissolved in 2.0% phosphotungstic acid, 13 min; dehydrate quickly in 2 changes of 95% alcohol and 2 changes of absolute alcohol; clear in several changes of xylene; mount in a synthetic resin. Acid mucopolysaccharides are stained turquois blue; prekeratin and keratin are orange to red orange.     

The Demonstration of Beta Cells in Pancreatic Islets and Their Tumors

The stain is applied routinely to tissues fixed in 10% buffered formalin (pH near 7.0) or in Bouin's fluid. Bring paraffin section to water as usual and mordant 72 hr in 5% CrCl₃ dissolved in 5% acetic acid. Wash in water and in 70% alcohol and stain 6 hr. Formula of staining solution: new fuchsin, 1% in 70% alcohol, 100 ml; HCl, conc., 2 ml and paraldehyde, 2 ml, mixed together and added to the dye solution; let stand 24 hr before use. After staining, wash in running tap water 5-10 min, rinse in distilled water and counterstain if desired. Dehydration in alcohol, clearing and covering completes the process. When the paraldehyde is obtained from a freshly opened bottle, standardized staining times can be used and thus eliminate the necessity of differentiating individual slides. The granules of beta cells stained deep blue to purple and were demonstrated in the pancreatic islet of man, dog, mouse, frog, guinea pig and rabbit.     

Staining Paraffin Sections with Protargol 6. Impregnation and Differentiation of Nerve Fibers in Adrenal Glands of Mammals

A series of experiments with protargol staining of nerve fibers in mammalian adrenal glands has yielded the following procedure: Fix-1-2 days in a mixture of formamide (Eastman Kodak Company) 10 cc, chloral hydrate 5 g., and 50% ethyl alcohol 90 cc. Wash, dehydrate and embed in paraffin. Cut sections about 15 and mount on slides. Remove the paraffin and run down to distilled water. Mordant 1-2 days in a 1% aqueous solution of thallous (or lead) nitrate at 56-60°C. Wash thru several changes of distilled water and impregnate in 1% aqueous protargol (Winthrop Chemical Company) at 37-40°C. for 1 to 2 days. Rinse quickly in distilled water and differentiate 7-15 seconds in a 0.1% aqueous solution of oxalic acid. Rinse thru several changes of distilled water for a total time of 0.5 to 1.0 rain. Reduce 3-5 rain, in Bodian's reducer: hydroquinone 1 g., sodium sulfite 5 g., distilled water 100 cc. Wash in running water 3-5 min. and tone 5-10 min. in a 0.2% gold chloride solution. Wash 0.5 min. or more and reduce in a 2% oxalic acid solution to which has been added strong formalin, 1 cc. per 100. (Caution. This last reduction is critical and over-reduction can spoil an otherwise good stain; 15-30 seconds usually suffices, and the sections should show only the beginning of darkening to a purplish or gray color.) Wash, fix in hypo, wash, dehydrate and cover.     

Toluidine Blue-Alizarin Red S Staining of Cartilage and Bone in Whole-Mount Skeletons in Vitro

Cartilage and bone of the developing skeleton can be reliably differentiated in whole-mount preparations with toluidine blue-alizarin red S staining after FAA fixation. The recommended staining procedure is based chiefly on the use of newborn white and Swiss-Webster mice, 4-9 days postnatal, but was tested also on mice and rats 3-8 wk of age. Procedure: Sacrifice, skin, eviscerate, remove body fat, and place specimens in FAA (formalin, 1; acetic acid, 1; 70% alcohol, 8) for approximately 40 min. Stain in 0.06% toluidine blue made in 70% ethyl alcohol for 48 hr at room temperature. Use 20 volumes of stain solution to the estimated volume of the specimen. Destain soft tissues in 35% ethyl alcohol, 20 hr; 50%, 28 hr; and 70%, 8 hr. Counterstain in a freshly prepared 1% aqueous solution of KOH to which is added 2-3 drops of 0.1% alizarin red S per 100 ml of solution. Each day for 3 days, transfer the specimen to a fresh 1% KOH-alizarin mixture, or until the bones have reached the desired intensity of red and soft tissues have cleared. Rinse in water, and place in a 1:1 mixture of glycerol and ethyl alcohol for 1-2 hr, then transfer the specimen to fresh glycerol-alcohol for final clearing and storage. Older mice and rats require procedural modifications: (1) fixation for 2 hr, (2) 0.12% toluidine blue, (3) maceration for 4 days in 3% KOH-alizarin, and (4) preliminary clearing for 24 hr in a mixture of glycerol, 2; 70% ethyl alcohol, 2; and benzyl alcohol, 1 (v/v) before placing in a 1:1 alcohol-glycerol mixture.     

A Phloxine-Azure-Hematoxylin Sequence for Differential Staining of Cells in Pancreatic Islets

Sections of 6 μ from tissues fixed in Susa or in Bouin's fluid (without acetic acid) and embedded in paraffin were attached to slides with Mayer's albumen, dried at 37 C for 12 hr, deparaffinized and hydrated. The sections fixed in Susa were transferred to a I₂-K₁ solution (1:2:300 ml of water); rinsed in water, decolorized in 5% Na₂S₂O₃; washed in running water, and rinsed in distilled water. Those fixed in Bouin's were transferred to 80% alcohol until decolorized, then rinsed in distilled water. All sections were stained in 1% aqueous phloxine, 10 min; rinsed in distilled water and transferred to 3% aqueous phosphotungstic acid, 1 min; rinsed in distilled water; stained 0.5 min in 0.05 azure II (Merck), washed in water; and finally, nuclear staining in Weigert's hematoxylin for 1 min was followed by a rinse in distilled water, rapid dehydration through alcohols, clearing in xylene and covering in balsam or a synthetic resin. In the completed stain, islet cells appear as follows: A cells, purple; B cells, weakly violet-blue; D cells, light blue with evident granules; exocrine cells, grayish blue with red granules.     

Staining and Coating Roller-Tube Tissue Cultures

Decanted roller-tube tissue cultures are fixed either by oven drying (60-63°C) for 3 hr or by methyl alcohol for 5 min and stained within the tube with Harris hematoxylin (diluted 1:1) and 4.4% alcoholic eosin. Oven drying before staining emphasizes cytoplasmic detail, whereas methyl alcohol produces distinct nuclear detail. After dehydrating and clearing by alcohols and xylene, 1.0 ml of Fisher's Permount is pipetted into the roller tube which is held at a 5-10 angle, rotated until the tissue sheet is covered, and placed in a paraffin oven at this angle for 16-20 hr. With a few exceptions, a majority of tubes showed no tissue drying and only minimal fading after 4-6 mo.     

Modified clearing method to show sieve tubes in minor veins of leaves

Leaf samples of Glycine max and numerous other dicotyledonous species were cleared by a common, well established procedure modified by using more concentrated (10% w/v) aqueous NaOH, and by leaving samples in NaOH for 2-4 weeks and in chloral hydrate for 3 days, all at room temperature. A single dye, chlorazol black E (1 g/100 ml absolute ethanol), is used to stain for 3-6 min. Samples are mounted with the lower epidermis upward. Sieve tubes in favorable material can be seen in minor veins and vein endings.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

Differentiation of Mastocyte Granules by Tartrazine Counterstaining After the Periodic Acid-Schiff Procedure

When the periodic acid-Schiff stain is followed by a counterstain of 1% tartrazine O in 1% acetic acid, mastocyte granules turn to a brick-orange color. Pharyngeal and intestinal mucus remain unchanged and gastric mucus is only slightly affected. The best results were obtained after fixation in Cajal-De Castro fluid (95% ethanol, 50 ml; distilled water, 50 ml; chloral hydrate, 5-10 gm; and nitric acid, 5 ml).     

A Hematoxylin Staining Procedure for Maize Pollen Grain Chromosomes

Studies of postmeiotic chromosome behavior have been impeded by the thick exine and abundant starch grains of maize pollen. Staining pollen grain chromosomes with acetocarmine is tedious and gives inconsistent, often unsatisfactory results. A hematoxylin stain, used in conjunction with the clearing agent chloral hydrate, has been successfully used by the authors to stain chromosomes, nuclei and sperm cells of the maize pollen grain. An ethanol-formaldehyde fixing fluid is used to fix and preserve the pollen samples. The procedure, which is rapid and simple, gives excellent preparations with both fresh and fixed material. Stained preparations do not get darker with time, as is typical of other hematoxylin stained materials.     

A hematoxylin staining procedure for maize pollen grain chromosomes

Studies of postmeiotic chromosome behavior have been impeded by the thick exine and abundant starch grains of maize pollen. Staining pollen grain chromosomes with acetocarmine is tedious and gives inconsistent, often unsatisfactory results. A hematoxylin stain, used in conjunction with the clearing agent chloral hydrate, has been successfully used by the authors to stain chromosomes, nuclei and sperm cells of the maize pollen grain. An ethanol-formaldehyde fixing fluid is used to fix and preserve the pollen samples. The procedure, which is rapid and simple, gives excellent preparations with both fresh and fixed material. Stained preparations do not get darker with time, as is typical of other hematoxylin stained materials.     

Adaptation of Mallory's Trichrome Stain to Embryonic and Fetal Material

Based upon results of an investigation of the role of phosphotungstic acid in connective tissue staining, the Mallory trichrome stain was adapted to sequential application of all three dyes, thus making it usable on embryonic and fetal material. Ten to twelve day postconception mouse fetuses were formalin fixed and paraffin embedded. Staining was as follows: (1) 1% aqueous acid fuchsin for 5 min followed by not more than 30 sec in running tap water; (2) 2% aqueous phosphomolybdic acid (PMA) for 10 min followed by a 2 min running tap water wash; (3) staining in 0.5% aniline blue in 8% acetic acid for 10 min, followed consecutively by 30 sec in running tap water, 2% aqueous PMA for 2 min, and 30 sec in running tap water; (4) 2% orange G in 8% acetic acid for 5 min, and rinsing for 30 sec in running tap water. Dehydration in ethanol, t-butanol, acetone, or by blotting followed by 1:3 terpineol-xylene, clearing in xylene and mounting, completed the procedure. The 30 sec tap water rinses can optionally be replaced by 1-2 min in 8% acetic acid. Sections can be made redder by increasing acid fuchsin staining time, or increasing time in the first PMA; red can be decreased by decreasing staining time, increasing time of the 2 min tap water wash, or decreasing time in the first PMA. Blue or orange staining can be increased or decreased by varying staining times in these solutions. Sharper differentiation may be obtained by increasing the time in PMA.     

A NEW CLEARING-SQUASH TECHNIQUE FOR THE STUDY OF OVULE DEVELOPMENT IN ANGIOSPERMS

The simple, efficient method described here for the study of ovule and megagametophyte development in angiosperms provides for the extension of investigation beyond the limits imposed by the traditional but arduous section technique. Excised pistils previously fixed in FPA₅₀ and stored in 70 % ethanol are placed in a clearing fluid composed of lactic acid (85 %), chloral hydrate, phenol, clove oil, and xylene (2:2:2:2:1, by weight). After 24 hr, ovules dissected from the ovularies are transferred with some of the fluid to a slide, covered so that the cover glass is supported laterally by two permanently affixed covers, and examined with phase contrast optics. The unique action of the clearing fluid permits the study of cellular structure with the phase oil objective focused at any focal plane within the ovule. Downward focusing thus reveals a series of optical sections in the sagittal, frontal, or transverse plane depending on the orientation of the ovule. Orientation can be altered by a slight shifting of the cover glass on the lateral support mounts. The ovules become quite fragile in the clearing fluid. Pressure applied to the cover glass gradually breaks the ovule apart without disrupting the structural integrity of individual cells. This squash procedure provides for extending observations to cytological features of megasporocytes, megaspores, and megagametophytes previously identified in intact ovules. The new method is applied here to the study of ovule development in two unrelated species, Cassia abbreviata Oliver var. granitica Bak. f. (Leguminosae) and Ludwigia uruguayensis (Camb.) Hara. (Onagraceae). For best results, the ovules of Ludwigia must be pretreated in lactic acid (85 %) for 24 hr prior to application of the clearing fluid. Other methods for pretreatment likely will be required as the technique is applied to a wider range of flowering plant species.     

An improved method for clearing and staining free-hand sections and whole-mount samples

BACKGROUND AND AIMS: Free-hand sectioning of living plant tissues allows fast microscopic observation of internal structures. The aim of this study was to improve the quality of preparations from roots with suberized cell walls. A whole-mount procedure that enables visualization of exo- and endodermal cells along the root axis was also established. METHODS: Free-hand sections were cleared with lactic acid saturated with chloral hydrate, and observed with or without post-staining in toluidine blue O or aniline blue. Both white light and UV light were used for observation. Lactic acid was also used as a solvent for berberine, and fluorol yellow for clearing and staining the samples used for suberin observation. This procedure was also applied to whole-mount roots with suberized celllayers. KEY RESULTS: Clearing of sections results in good image quality to observe the tissue structure and cell walls compared with non-cleared sections. The use of lactic acid as a solvent for fluorol yellow proved superior to previously used solvents such as polyethylene glycol-glycerol. Clearing and fluorescence staining of thin roots such as those of Arabidopsis thaliana were successful for suberin visualization in endodermal cells within whole-mount roots. For thicker roots, such as those of maize, sorghum or tea, this procedure could be used for visualizing the exodermis in a longitudinal view. Clearing and staining of peeled maize root segments enabled observation of endodermal cell walls. CONCLUSIONS: The clearing procedure using lactic acid improves the quality of images from free-hand sections and clearings. This method enhances the study of plant root anatomy, in particular the histological development and changes of cell walls, when used in combination with fluorescence microscopy.