adhesive tape, x 40	insulation tape, x 10	paper label, x 10	razor blade, x 10

On the nature of the separators.-  I have tried four kinds of separators. The self-adhesive paper labels, one black plastic electrical insulation tape, one brand of clear adhesive so-called “Scotch” tape and the razor blade itself. To determine their thickness I stick a sample of the tapes to the edge of a glass slide, and cut it level with a razor blade. I take pictures with the 10 x and 40 x objectives using the COL-D3 contrast disc that gives a very good optical separation of the components, and I measure the thickness with the calibrated measuring tool of the camera program. The razor blade was broken into two halves and one half was shaped as a V that can be put upright over a glass slide to offer the sharp edge to the objectives.

The results are confusing, not about the materials but about the thickness of the cut materials.

The limits for the thickness of the sections.- The Scotch tape (including the relatively wide layer of adhesive is 47-48 (roughly 50) microns thick. The adhesive paper label is 135 microns thick; the insulator tape is roughly 200. And the thickness of the razor blade itself turns out to be 100 microns.

I make one more measurement: the width of the cutting edge (not its thickness). It turned out to be 300 microns wide. So the sharp edge is an isosceles triangle with a base of 100 microns, and a height of  300 microns. The edge is at 50 microns from each side of the blade.

But only rarely have plant sections been under 100 microns, and normally they are of 125 to 150 microns. I think this is the result of the sharp edges being V-shaped. Thus the edge is in the center of the 100 microns blade (50 microns away from any face). If you put in a separator of 50 microns the gap between the sharp edges (not the blades) are 50+50+50 = 150 mic. Theoretically this is the thinnest section you can cut if the blades are parallel (and were not flexible, thus allowing thinner or thicker sections). Do not think of putting the blades together without a separator. Most sections are not hard enough to separate the blades.

I tried several methods to make the internal sides of the sharp edges more or less parallel and closer, but not one of them was successful.

The most easily cut and thinnest sections were obtained using half a razor blade as a separator. And this turned out to be also the easiest way to build the slicer. But some materials, and especially the longitudinal sections of stems, cut better if the separator is the adhesive tape.

So as stated in the first part of this article, now I currently use two versions of the slicer.

The best razor blades. These are the most rigid ones. Try several trade marks if you can. Flexible blades can be separated by the incoming section. This ends in a wedge profile. To study the anatomy of a plant this is not important. But it is for photographic recording. Anyway, even with flexible blades and by making 3 or 4 attempts with different pressures and speeds, should give you a useful section.

Appropriate materials. Highly lignified materials need to be sliced very slowly and with a firm pressure. You may experience great difficulties trying to make sections of Gramineae stems that are very hard. Very soft materials tend to collapse if the blades are too close. The best materials are the medium lignified ones. 

Generally the thinnest section is the best. But after several attempts you should be convinced that a medium thick section is perfect for many tasks. If it is cylindrical (not wedged) the surface can be studied at high powers successfully. And it allows a good use of the COL, DF and RhQ filters. Many times these are useless when the section is too thin.                                                                                            

Preserving materials. If you are collecting far away from your laboratory or if you want to preserve some materials for future studies, you can fix them. The simplest fixative recommended for botanical materials is 70% ethyl alcohol. Of course some organs, like petals for example can wilt, but their anatomy should be preserved. For anthers, ovaries, root tips, and any reproductive organ Methacarn 95% could be a better choice. After 1 or 2 hours in Methacarn, change to 70% alcohol for one hour and preserve in a second change of 70% alcohol, better with a 1-2% of glycerin. Don’t forget to adequately label your sample.

 Shown left is a section of a basil stem from the exterior epidermis to the central pith parenchyma, captured at x 40 and half reduced, brightfield, no staining,

Amateur staining.- Professional staining, as you have discovered if you've just read the references, calls for metachromatic dyes like o-toluidine, or for safranin, or double stains such as carmine-green or similar. Or  phloroglucynol for lignified tissues. All are difficult to buy or very high priced items.

x-section of the stem of a wild tropical vine. Hypochlorite - gentian violet.	x-section of the lamina of one Aptenia leaf, with a dense palisade parenchyma, and a spongy mesophyl. Hypochlorite - gentian violet.

I have found that a very common and easy to buy dye is gentian violet, a former disinfectant for the babies “mugget”, and other infections. Here it is sold as a one percent solution in water. It is very stable; 20 ml is a provision for all your life, because it is used at a drop for every 10 or more milliliters of water. This working solution, which also keeps very well, stains the schlerenchyma and fibers a dark violet, the xylem a more deep color with a red tinge, and the cambium, phloem and the collenchymas a slight purple. The cuticle of the epidermis is also deep colored. As discussed later, also methylene blue can be resorted to. 

Contrast Discs.- If you cannot get gentian violet (also known as crystal violet) you do not need to renounce  technicolor. Your best choice is to take recourse of the contrast discs. The “dispersion staining” that Ted Clarke has appropriately described for DF contrast disks at high powers, is a common characteristic well known to amateurs working at 4 or 10x.

The behavior of the contrast discs is so dependent on the thickness and the nature of the materials that you must play around with your own discs to try the best effect. I have more than 30 different ones. I never know which of these will do the best job with a particular section. But the darkfield stops, some Rheinberg filters and some modifications of the Nomarski simulation filters proposed by Wim van Egmond give outstanding images. The COL filters normally behave only as special darkfield stops.

Section of the wall of an hybiscus young fruit. The white points in the endodermis are little crystals of calcium oxalate. 10x, Darkfield stop.	A longitudinal section of the stem of Aptenia. Idioblasts, charged with acicular raphids show a red glow. At right the epidermis. The orange band in the center is the xylem of the vascular bundle. 10x, with a RhQ filter.

 The better Rheinbergs are blue centered (12-15 mm diameter) with the exterior ring in different tints of yellow or orange. A very useful modification is a disc with a first clear ring 2 mm wide, a red ring 3 mm wide, a clear one of 2 mm, a blue one of 2 mm and a black center 12 mm in diameter. Centered or a little displaced it gives many useful nuances. Another remarkable Rheinberg is the Quadrant’s Rheinberg (RhQ).

The original van Egmond filters are black discs, with a marginal transparent crescent and a circular blue or purple center. They are a combination of darkfield with Rheinberg and oblique lighting and give its best results with discrete objects like fibers, spicules, sand and the like, especially if they are of high refractive indexes.

I replace the black backgrounds for colored ones (deep blues and deep reds for example) and make the centers of a diameter similar to the darkfield disk for the objective in use, in a contrasting color. I leave the transparent crescent unchanged. Its best performance is with relatively thick sections.

These contrast discs (darkfield, Rheinberg and Nomarski simulators) allow the optical differentiation of the different tissues, in medium thin sections, simulating stained sections. They are really very useful (for the 4x and the 10x and with limitations up to the 40x objectives) to give variety and gaiety to the photographed sections. Resolution suffers a little with the 40x. A word of warning: the colors most useful for the visual rendering of sections are aggressive for the sensor of my photographic camera and gives a very bad rendition when compared with the visual image. But they behave very well in direct view. My Col-D3 with a yellow background gives strange results. I see the image in yellow nuances, but the camera records them as many different blue tints. So be prepared, in case your camera behaves in the same way. Judging by published comments most cameras, including the high priced ones, share this problem.

Anyway the difference between a brilliantly colored section and a gray and more opaque one is really outstanding.

Acicular raphids in an idioblast. Aptenia. Brightfield. No stains.	Calcium oxalate crystals in cortical parenchyma of Aptenia. Brightfield.	Epipremmnum x section in brightfield, Iodine added to stain starch.

Uncolored sections.-  If you mount your just made sections without any subsequent treatment in glycerin or PVA-G you can make a profound and informative study of the anatomy of an almost living material. Glycerin and PVA-G act as preservatives and even the chlorophyll lasts for many days unchanged. All vegetable tissues are easily recognizable by their morphological traits, and the arrangements of the studied organs are very characteristic. You can discover the idioblasts with its secreted crystals, see even the nucleus of many cells, and the plastids containing oils and starch. If you add to the glycerin a trace of iodine, the starch will be colored blue and you can easily discover the areas of starch production. All this is lost if you void the cells of its contents with hypochlorite. Note: don’t try to add iodine to sections made from materials recently exposed to high levels of sunlight; you risk having a mostly blue and illegible specimen. Try materials exposed to low sun intensities. 

Microwave ovens use. -  Mounting in glycerol, or even in PVA-G can exert on the living cells an excessive osmotic pressure. Delicate materials, like algae, tender hairs, epithelium cells, fungal hyphae and fruiting bodies, and so on, can collapse. You can of course use some fixatives and dehydrating routines for a lengthier mounting in glycerin. But a useful rapid technique is to put the materials or sections, collected in a small Petri dish, or even the recently made slides, on the turntable of the microwave oven and apply a 12 to 20 seconds period of radiation at full power. This enhances the infiltration of the mounting media, evaporates water, gets rid of air bubbles, and the cells become turgid again. Experiment to find the best timing for your own oven. Mine is a 700W model. You can extrapolate suitable times for your oven wattage.

x-section of the leaf of a wild vine treated with a mixed technique: Hypochlorite-gentian violet viewed through a COL-D3 contrast disk. The background was inserted in PhotoPaint.