TECHNICAL TIPS ON THE USE OF THE PRECISE DOUBLE RAZOR BLADES SLICER
I bring together here several tips about the design and performance of the slicer described in the first article. I add only the minimum technical information for the pictures, because in successive articles I hope to present short illustrated monographs of some plant species, studied with the aid of the slicer. A formulary with laboratory protocols is also a future project.
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.
Using the Scotch tape as a separator I make the razor blade sandwich and put it upright under the 4 x and the 10 x objectives. The width of the gap between the blades is roughly 50 microns as it can be suspected.
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.
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
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.
after several attempts you should be convinced that a medium thick
perfect for many tasks. If it is cylindrical (not wedged) the surface
studied at high powers successfully. And it allows a good use of the
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.
I have found that a very common and easy to buy
dye is gentian violet, a former disinfectant for the babies “mugget”,
infections. Here it is sold as a one percent solution in water. It is
stable; 20 ml is a provision for all your life, because it is used at a
for every 10 or more milliliters of water. This working solution, which
very well, stains the schlerenchyma and fibers a dark violet, the xylem
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
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
around with your own discs to try the best effect. I have more than 30
ones. I never know which of these will do the best job with a
section. But the darkfield stops, some Rheinberg filters and some
modifications of the Nomarski simulation filters proposed by Wim van
give outstanding images. The
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.
Uncolored sections.- If you mount your
just made sections without
any subsequent treatment in glycerin or PVA-G you can make a profound
informative study of the anatomy of an almost living material. Glycerin
PVA-G act as preservatives and even the chlorophyll lasts for many days
All vegetable tissues are easily recognizable by their morphological
traits, and the arrangements of the studied organs are very
discover the idioblasts with its secreted crystals, see even the
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
exposed to high levels of sunlight; you risk having a mostly blue
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.
Please report any Web problems or offer general comments to the Micscape Editor.
Micscape is the on-line monthly
magazine of the Microscopy
site at Microscopy-UK