Bryum caespiticium - peristome
Over time, every bryologist develops ways of getting the most out of their fieldwork and microscopy. Many techniques are passed informally from one person to another and may not be written down anywhere, but they can make a huge difference to the successful study of bryophytes. In this section, we share some of our favourite tips and techniques – we hope they’re useful!
The leaves of certain mosses have an annoying tendency not to lie flat on a slide, meaning that upper leaf features are often obscured. Falling in this category are the slightly curved leaves of many acrocarps, including Dicranum, Dicranella, Grimmia and Ceratodon. When keying out Didymodon it is important to be able to see whether the cells of the costa in the upper part of the leaf on the ventral (upper) surface are isodiametric or elongate and also, for Didymodon vinealis and related species, to see if there is a translucent groove in the apical part of the leaf. When leaves of such species are stripped off stems and floated under a coverslip, they nearly always orient themselves the ‘wrong’ way up.
One way to get uppermost part of leaves to lie flat on the slide is to take a whole dry stem, place it on a dry slide under a stereo microscope near a drop of water and, pinning the stem down with a forefinger, cut the uppermost leaves transversely above their mid-point with a razorblade. When the half-leaves are pushed into the droplet, you will find they lie flatter than if they were still attached to their lower portions. In Didymodon, this technique can also be combined with cutting leaf sections, by making the first cut around 4/5 way up leaf tip and then slicing back against the finger to produce a series of very thin nerve and lamina sections.
When examining the branch leaves of Sphagnum species, it is often necessary to view or measure pores or hyalocysts visible on the convex outer leaf surface. Unfortunately, because such leaves tend to be very concave, they have a strong tendency to float concave-side up when placed in a drop of water. You can correct this by floating the leaves in a drop of water on a coverslip, placing the slide on the coverslip and then turning it over so that the coverslip is uppermost. This technique can work for Didymodon leaves too.
To avoid getting fingerprints on coverslips, some people keep them on a thin piece of foam; it’s then easy to press finger and thumb into the foam on either side of the coverslip to pick it up. As a further innovation, you could glue a small piece of foam to your stereo microscope stand for this purpose. Read David Wagner’s article about coverslip handling.
Coverslips and slides can be reused many times if kept clean – if they do get dirty, simple apply a few drops of Isopropyl Alcohol (IPA) and wipe gently with a tissue or clean lens cloth.
It is a good idea to have a couple of sizes of coverslip available. It’s surprising what a difference it can make searching for a few leaf sections if you use a small circular coverslip. But a large one is good for examining a number of leaves, especially big ones.
Stain Sphagnum leaves with 1% Methylene Blue, Crystal Violet (or another stain of your choice) to see hyalocyst pores etc. more easily. Staining of the leaves of other species such as Dicranum will also make porose cell walls more visible. Add a tiny amount of stain via a fine dropper (either purchased or made from a recycled eye drops bottle) to a drop of water on a slide and then add a coverslip. Even distribution of the stain under the cover slip can be encouraged by carefully using a tissue to dab the edge of the coverslip to pull liquid across. If you’ve used too much stain, add more water and draw through by using a tissue on the opposite side of the coverslip.
For staining Leucobryum sections or similar small specimens, try placing a drop of stain next to the water containing your sections on the slide. Then drag a small amount of stain into the water using a needle and repeat until it looks dark enough.
Branch leaf, showing pores
Soil around the rhizoids of small acrocarps such as Bryum, Dicranella and Pohlia often obscures rhizoidal gemmae needed for identification. To remove the soil, half-fill a small sample pot with water, add a small number of stems and soil (it helps if particularly solid lumps are gently broken up with forceps first), screw the lid down firmly and let it soak for at least 10 mins. Then give it several vigorous shakes; if the soil is heavy clay more may be needed. This procedure will detach a lot of soil from the rhizoids and when the plants are then viewed in clean water under a stereo microscope any rhizoidal gemmae should be more obvious. This technique also works quite well to remove encrusted sediment from the leaves of aquatic species such as Rhynchostegiella teneriffae.
Other techniques for removing soil and dirt from specimens on a slide include the use of a very fine paintbrush (place the stem or leaf in water, grasp the base with forceps and gently brush the dirt off). A needle dropper bottle with a very fine nozzle can also be used to wash soil off specimens under a dissecting microscope.
Rhizoidal tubers
If you have access to Potassium hydroxide (KOH), it can be used to clear cell contents and make cell walls easier to see (useful for determining whether cell walls are porose, and for viewing trigones in liverworts).
** Take care when using any chemicals. KOH is a caustic chemical and can cause severe damage such as burning or ulcers, on contact with skin. **
Prepare a 10% solution of KOH. This can be kept in a plastic bottle (an old eye-dropper bottle is ideal) for many months. Place a few leaves in a drop of KOH and leave for 5 minutes, then transfer the leaves to a fresh drop of water, add coverslip and view.
To progress in the study of bryophytes, mastery of techniques of leaf, capsule and stem sectioning is important. It is something that improves with practice, so do not despair if your first few sections are not what you hoped for. And, if you get the chance, observe how experienced bryologists do their sectioning; there is often more than one technique so it’s important to find one that works for you.
As a general rule, sectioning requires concentration, a steady hand, and good tools – a reasonable stereo (dissecting) microscope with a large enough stage to work on, a pair of fine-tipped dissecting forceps or needles and a sharp razorblade. For best results when sectioning leaves, always use a brand-new razorblade as they get dull surprisingly quickly. Used blades can however be used for cutting capsules and other thick material. There is no need for special blades – you can use the replacement blades for old-fashioned (reusable) men’s razors sold by many high street and online retailers. To get more life out of these blades, some people cut them in half.
A comparison of blade sharpness for various different brands can be found on the Refined Shave website (thanks to Richard Zander for his article in the Bryological Times 2022 for this tip).
Leaf sections, showing guard cells in 2 tiers
This technique is good for leaves and stems when material is plentiful – and a good one for beginners as it requires less dexterity than some others. Please be careful when handling razor blades!
Cutting sections of leaves of mosses works best when the leaves lie more or less appressed to the stem. Some species which have leaves that stick out or curve away from the stem when moist e.g., Didymodon, Polytrichum can be left to dry, when leaves often lie more appressed. For these species, select a single, well-grown dry shoot and place it on a slide under a stereo microscope. Align the shoot in the middle of the slide so it is parallel to the long side of the slide and its uppermost leaves are pointing directly at a drop of water no more than 1cm away. The trick is to keep the shoot dry but to push the sections off into the droplet as you work. Press the tip of a forefinger down onto the shoot so that only the tips of the uppermost leaves are visible. Then, looking down the microscope, take a new razorblade and cut across the shoot, pushing back against your finger. Aim to get multiple slices by slicing repeatedly in the same place, pushing the arisings to the droplet of water with the blade as you go. Doing this, you will end up with a considerable quantity of unusable material in the droplet, but there should also be some good (thin) sections. If you don’t succeed the first time, don’t be disheartened. Improvement definitely comes with a little practice. With species that have naturally crisped or curled leaves when dry, the ‘finger’ method can also be used with hydrated (but not wet) plants.
You could also replace the finger with a slide to hold down the shoot. This has the advantage of offering a straight edge to slice against, but the slide is generally a little harder to move if you are aiming for multiple sections.
If you’re just starting, it’s a good idea to practice on something easy first. Polytrichum leaves tend to be quite large, straight and stiff so they’re relatively easy to handle. The leaf sections are interesting too!
Ken Adams described his technique for cutting thin sections in Field Bryology 88, 2006, and has provided an illustrated version to download below. As BBS Librarian he also sells the mini blades he uses, details are on the Library, Sales and Loans page.
Section cutting using the ‘floating slide’ method
This is a simple and quick method that works well for Sphagnum leaf sections and needs no special equipment other than a small pair of sharp scissors. It is also suitable for Polytrichum and may be worth trying on other shoots that are not too small or delicate.
If all else fails, you might like to try a technique invented by a lichenologist, John Skinner, and described in a download from the BLS website. This involves mounting leaves (or whole stems if you have plenty of material) in Gum Arabic, letting it dry until firm (about 30 mins), and then using your favourite technique for guiding a razor to cut sections. It takes a little longer than freehand sectioning, but avoids some of the frustration!
Download article from the BLS websiteTo examine the peristomes and exothecial cells of e.g., Bryum, Orthotrichum/Lewinskya, Ulota, soak a well-developed capsule in water. Then dab it off with a tissue and place it on a slide under a stereo microscope. To examine the peristome and cells around the annulus, hold the capsule firmly with forceps and cut transversely across it not far below the peristome. Discard the lower part of the capsule and any seta before cutting the peristome and upper part of the capsule in half longitudinally and moving the halves carefully to a new (dry) slide. If any spores are present most will be left behind at this stage. Carefully use the forceps to position the peristome halves so that one shows the inner peristome uppermost and the other shows the outer peristome. Take particular care not to touch the very fragile inner peristome. Put a coverslip on top and irrigate it gently from the edge using a fine water dropper. Air bubbles can be reduced by adding a tiny amount of domestic surfactant e.g., soap, shampoo or washing-up liquid to the water to reduce surface tension.
For some of our Ulota species, it is necessary to examine inner peristome teeth, and if you have ever tried doing this, you will have discovered that when moist, the outer peristome teeth spring up and hide the endostome teeth.
One way to overcome this is to cut a dry capsule in half lengthways, place half with the inner peristome uppermost and half with the outer peristome on top. Place a coverslip on the dry slide then press down firmly and place a drop of water at the edge of the coverslip and allow it to seep under. By holding the coverslip down throughout, the outer peristome teeth should remain reflexed, allowing you to examine the inner teeth. When you’ve done that, you can add a little more water so that the coverslip lifts slightly and allows the exostome teeth to straighten.
A few images of sections under the high power microscope showing what you are aiming for.
This is good for putting a precise amount of water (or undiluted stain) on a slide to mount leaves etc. It is also handy for washing soil and other debris off specimens. Make sure you buy one with a very fine needle. Some even come with a selection of different needle tips.
Needle dropper bottle
These are similar to normal microscope slides but have a small well in the centre, and are designed for the examination of larger specimens. For example they are useful for viewing whole shoots of liverworts to see underleaves and leaf orientation, or whole shoots of small acrocarps when searching for male organs, or rhizoidal tubers.
2 examples of cavity well slides
These can be used to gently remove dirt and air bubbles from leaves, tubers etc. Sets of variously sized brushes can be bought very cheaply on Amazon and are perfectly good enough as long as the bristles don’t fall out!
Use to illuminate a slide from above. This can be useful for finding tubers in a mass of soil on a slide, or for viewing more detail of opaque objects under high power.
Use of an LED torch with a compound microscope
