Rapid Preparation & Examination of Plant Sections with Microscopy

Posted by Bon Hui on

by Dr. Robert Berdan
August 30, 2022
Fig. 1. Above is a hand cut section of an Aspen twig that was stained with Toluidine blue. Bright-field light microscopy 50X.


Free-hand sectioning of living plant tissues with a single edge razor blade often provides an adequate method for rapid and inexpensive microscopic observation of plants internal structure. Moreover this simple technique often results in high quality images (A. Lux et al., 2005). Alternative methods to cut plant sections include the use of a hand microtome, vibratome, and a freezing microtome. Only free-hand sections and the hand microtome will be described here. The number of research papers using free-hand sections is increasing, along with the use of fluorescent stains. I describe some simple methods for free-hand sectioning of plant tissues in order to take photomicrographs using various light microscopy techniques. These techniques can be used in the laboratory or the classroom. For detailed methods to staining methods see the references.


Equipment required


⦁ Plants – vegetables from your garden, fridge, weeds, leaf stems, pine needles, and shrubs
⦁ Single edged razor blades or disposable microtome blades
⦁ Fine detail paint brushes to gently move the cut sections
⦁ Petri dishes, watch glass or small bowls with water
⦁ Microscope glass slides, and No 1 or 1.5 cover glass
⦁ Forceps, dental wax
⦁ Staining solutions: Toluidine blue, Safranin O, Aceto-carmine, and, Potassium iodide
⦁ Pasteur pipettes, water bottles with distilled water and tap water
⦁ Paper towels
⦁ Microscope slides
⦁ Stereo microscope (optional but I like to see the sections clearly while cutting)
⦁ Light microscope with bright field (optional illumination includes phase contrast, dark-field, or fluorescence)


Methods used to cut and stain sections


Some plants will cut easier than others and your sections will get better with practice. Try to cut the stems, roots and leaves at right angles. You can break the single edge razor blades in half and put tape on the broken half. Sections can be cut by hand by holding the stem in one hand between your thumb and first finger and cutting the sections with a single edge razor by drawing it toward your thumb nail (see YouTube video). I use a special razor blade holder (Fine Science tools) to hold part of the razor blade and cut plant sections on top of dental wax while examining them with a stereo microscope. Some sections will be too thick, so always cut several sections and choose the thinnest – it takes practice to get good thin sections.

Fig. 2. The above photo shows a wide variety of cutting blades and tools that can be used to cut thin sections of plants.


In addition to cutting plant sections by hand, hand-microtomes are relatively inexpensive and can help you obtain thin sections consistently. Hand microtomes can use a variety of razor blades, cutter blades and even disposable microtome blades. I found cutting with disposable razor blades worked the best – see Fig. 2. The blade is pulled at an angle to slice the tissue and the sections then transferred to a bowl of water using a fine brush. It's important to cut through the tissue with a single, even stroke of the razor blade. Hand trimmers cost from $100 to $300. My favorite hand microtome was the American Optical Microtome circa 1957 (Fig. 4.). Blades should be cleaned with ethanol and wetted with water before sectioning (R. Berdan, 2020).

Fig. 3. The microtome on the left has a clamp that allows it to be fixed to a table. The middle microtome uses a disposable cutting blade and disposable single edge razor blades. The sample is clamped into the middle of the microtome and by rotating the base you can move it up in small increments. Samples are placed in the center and a micrometer gear at the bottom is used to raise the sample in 5 micron increments. On the right is a simple microtome that can even be home made. You can use a razor blade, cutter blades, double edged blades or the older shaving blades with a handle to cut sections. These microtomes cost $50 to $200.

Fig. 4. American Optical Hand Microtome circa 1950's I purchased from E-Bay. The sample is clamped down in the middle and a microtome blade is slid across the top rails to cut thin sections. At the bottom of this hand microtome there is a dial that moves the sample up in small increments of 5 microns.


After cuttings sections, they should be transferred to a small dish of warm water where they will float and flatten. The sections can then be transferred to a microscope slide with a fine brush. The sections can be viewed after mounting on a glass slide with water and a coverslip. The slides can then be viewed with a light microscope using bright-field, phase contrast, dark-field, or fluorescence.

Some woody stems need to be softened before cutting. This can be done by boiling the twigs in water, or water with 30% glycerol. Plant tissues can also be fixed (e.g. Farmer’s fluid – 1 part glacial acetic acid and 3 parts 95% ethanol). Sections can be viewed unstained or after staining with Toluidine blue, Safranin O, Astra blue or Nile blue. Toluidine blue (0.5% w/v in water) is commonly used and stains different plant components different colours. It is a metachromatic stain.

Some tissues, like leaves or small stems need support before you can cut thin sections. This can be done by placing leaves into a cut in a carrot stem then cutting both the carrot and leaf. Other substances can also provide support such a potato or soft wood like balsa.

Studying wood sections can reveal information about climate change and the amount of rainfall (dendroclimatology). Pieces of wood larger than several millimeters are often cut using a sliding microtome, but small woody twigs can be sectioned by hand.


Clearing

Some plant tissues such as leaves, stems and seeds can be seen more easily if they are cleared (made transparent) before or after sectioning (Sharma, N. 2017). Ethanol and glacial acetic acid (7:1) or bathing the tissue in 1M sodium or potassium hydroxide (NaOH or KOH) for a couple of hours can clear some plant tissues. Some researchers use Lactic acid for clearing. Lactic acid must be removed if using a fluorescent microscope because it is autofluorescent.


Staining plant sections

There are a wide variety of methods to stain plant tissues depending on what an investigator is looking for. To start, I recommend examining plant sections without staining and trying different microscopy illumination methods. Many plant sections will exhibit autofluorescence when illuminated with blue excitation light due to the presence of chlorophyll and polyphenolic compounds. A good first stain to try on your sections is Toluidine blue (O’Brien, et al. 1964). Toluidine blue stains different plant tissues different colours e.g.
⦁ Pectin – red or reddish purple
⦁ Lignin – blue
⦁ Phenolic compounds – green to blue
⦁ Thin-walled parenchyma – reddish purple
⦁ Collenchyma – reddish purple
⦁ Lignified elements and sclerenchyma – green


Colours generated by Toluidine blue can provide information on the nature of the cell walls and other parts of the plant. Staining times for plant tissue can vary from a few seconds to hours, so there is some trial and error involved in getting optimum results. You can remove stain by washing it in the solvent.


Common stains used to stain plant tissues:


⦁ Toluidine blue – 0.1% w/v in water
⦁ Safranin 0 – 0.5% w/v in 50% ethanol (less than one minute ), 95% ethanol is available at liquor stores called Everclear 190 proof
⦁ Fluorescent Yellow 0.01% w/v in polyethylene glycol PEG 400
⦁ Acetocarmine or Aceto-orcine stains chromosomes in growing root tips
⦁ Potassium iodide – stains starch grains blue-black in colour
Once the sections are stained, they are rinsed in solvent. If the tissues are to be mounted permanently they need to be dehydrated with 50%, 75%, 95% ethanol, then placed into xylene or some other solvent that is miscible with the mounting media like Canada Balsam. If the sections are to be viewed immediately they can simply be placed on a microscope slide in water and covered with a coverslip. To make semi-permanent slides, you can put clear nail polish around the edges of the coverslip to prevent or reduce evaporation of the water for a few days.

Fig. 5. Popular tree branch sectioned with the American optical microtome and stained with Toluidine blue.  Focus stack 100X.

Fig. 6. Popular branch stained with Safranin O, Bright field microscopy - sectioned with American Optical table top microtome shown in Fig. 4. 100X.

Fig. 7. Hand cut cross section through a leaf vein from Cannabis sativa male 200X Dark-field microscopy.

Fig. 8. Cannabis sativa leaf stem in cross section cut by hand using a razor blade and stained with Safranin O. Left: Bright-field microscopy Right: Dark-field microscopy.

Fig. 9. Section of a blue spruce branch (Picea pungens) cut with a double edged razor blade and then stained with 0.1% Toluidine blue in water. The section is a panorama of 4 regions that were focus-stacked and then blended into a panorama. The section was photographed by bright-field microscopy at 50X. The branch was taken from a Blue spruce tree in my front yard.

 

Summary

Making sections of plants to observe with a microscope is relatively simple with single edge razor blades or a hand microtome, but takes practice to get good sections. The sections can be prepared quickly and reveal a surprising amount of information. Plant sections can be examined unstained or after staining and permit high quality photomicrographs. Some common stains used with plant tissues have been described here. A variety of hand microtomes and various cutting blades are also shown. With practice it is possible to cut reasonably thin sections 15-50 microns thick to examine the different cell types and identify various vascular components in the stems, leaves, and roots. These procedures are commonly used in research, and the classroom. If the techniques are used in the classroom a basic first aid kit for cuts should be readily available.

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References:

Lux A., S. Morita, J. Abe and K. Ito (2005) An Improved Method for Clearing and Staining Free-hand Sections and Whole–mount Samples. Annals of Botany 96: 989-996. DOI https://doi.org/10.1093/aob/mci266 PDF

Sharma, N. (2017). Leaf Clearing Protocol to Observe Stomata and Other Cells on Leaf Surface. Bio-101: e2538. 10.21769/BioProtoc.2538. DOI: 10.21769/BioProtoc.2538.

Young E.C. (1998) A Beginner’s Guide to the Study of Plant Structure. Pages 125-142 in Tested studies for laboratory teaching. Vol 19 (S.J. Karcher, Editor) Association for Biology Laboratory Education (ABLE), 365 pages. Chapter 9. https://www.ableweb.org/biologylabs/wp-content/uploads/volumes/vol-19/9-yeung.pdf

O’Brien, T.P., N. Feder, and M.E. McCully (1964) Polychromatic staining of Plant cell walls by toluidine blue 0. Protoplasma 59: 367-373.
http://www.cas.miamioh.edu/~meicenrd/Anatomy/Ch4_Histology/Polychromatic%20staining%20of%20plant%20cell%20walls%20by%20toluidine%20blue%20O.pdf

R. Berdan (2020) The Microscope Beauty of Plants and Trees - How to prepare and photograph plant sections for viewing with a microscope. Canadian Nature Photographer. https://www.canadiannaturephotographer.com/microscopicplants1.html

YouTube video: Beyond the Bean Making cross-sections by hand (stems) and using Toluidine blue 0.
https://www.youtube.com/watch?v=GlCOQijkIKc

 

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