TECHNIQUE
Study of the Pottiaceae entails techniques
common to the study of most mosses with a few modifications that address the
need to examine features of internal anatomy, use of color reactions to KOH
solution, or to make permanent microscopic mounts of delicate tissue. Zander
(1979g, 1980a, 1983b) and others (e.g. Lightowlers 1981; Long 1982b; Frahm
1981, 1990a) have detailed aspects of this, but a review is appropriate here.
Packets are folded from 16-pound substance
100 percent rag or buffered paper to 10 cm in height and 14 cm in width using
an easily constructed device consisting of a square board to which is affixed a
low ledge below with a metal template the bottom of which is screwed to the
ledge, held parallel and about 3 µm above the board. The template is slightly
less than 10 by 14 cm in dimension. Packets can be folded quickly and
accurately about this flange. Subpackets for fragments are made from thin rag
or buffered paper (we use a rag tracing paper) and may be cut to various sizes
and folded into square or triangular packets. Glue for labels is made from
polyvinyl alcohol dissolved in water and used as refill for commercially
available “roll-on” glues. Commercially available polyvinyl acetate glue is
also adequate and perhaps less liable to degradation through crosslinking. (A
planchet of polyvinyl acetate glue was dried in 1970 and to date retains its
original elasticity after about 20 years exposed to fluorescent lighting.
Evidently any plasticizer included in the formulation is not volatile or degraded
in this time frame.) Water-based glues are, of course, subject to weakening in
very humid environments.
In annotating specimens, care must be taken
to use inks that do not fade with time. This is also true with making slide
labels, which may fade surprisingly quickly when exposed to sunlight. India ink
is, of course, ideal. Attaching annotation labels to packets may be done with
glue or with plastic or plastic-coated paper clips (to avoid rust marks). It is
helpful to keep some insect-mounting pins available for attaching annotation
labels to specimens borrowed from herbaria that require such pinning.
Certain standard and some modified
microdissecting tools are used in the study and processing of specimens. Fine
watchmaker's forceps may be further sharpened with a file. Dissecting probes
are re-constructed with sewing needles of medium size and little flexibility
inserted into a standard wooden or composition probe handle to replace the
original comparatively coarse and bendable needle. Single-edge razor blades for
sectioning should be discarded after five to ten uses since they dull fast. One
holds a leaf or stem crosswise with a stiff dissecting needle, then slices the
material with a razor blade held longitudinally against the far side of the needle,
meanwhile rolling the needle slowly towards oneself to gradually expose uncut
portions of the material. Practice (and a relatively fresh blade) makes this
technique quite effective, even for very small leaves. Remember to scrape off
sections (especially stem sections) adhering to the razor blade with a
dissecting needle after cutting. The usual pair of compound and dissecting
microscopes are needed, but using an additional illuminator with the dissecting
microscope for fine dissections rather than just a single lamp will prove
surprisingly advantageous for observation of fine features.
Round cover slips of 18-mm size and the
thickest weight (number 2) are used for routine examinations in water or KOH
solution because of ease of handling and cleaning for reuse. Square 18-mm cover
slips of medium thickness (number 1) are used for making permanent mounts
because they are less expensive and can be luted (i.e., sealed to prevent
evaporation) more easily. To avoid annoying spills, a box of cover slips may be
glued to the microscope base or to the bottom of a low, flat tray.
Stains are not standardly used, although they
might be worthwhile in examining the pores in the basal laminal cells of some
taxa, notably species of the genus Tortula. Color reactions to alkali
provide, however, important data, and a two percent potassium hydroxide
solution is kept at hand in a plastic squeeze bottle. Potassium hydroxide
solution helps hydrate dry plants (cf. G. Smith 1971, p. 2). Hagen
(1929, p. 14) suggested the use of a 10 percent KOH solution soak for the
leaves of Pottiaceae species to enlarge papillae and make them more easily
visible; this effect has not been evident to me, at least at the two percent
concentration recommended. Glass containers should not be used because KOH
reacts with the walls to form a precipitate. Dilute hydrochloric acid is a
valuable reagent to help determine, from observation of gas bubbles, whether or
not a collection was made from a calcareous habitat. It will also help clean
plants of limy incrustations.
Pohlstoffe (Wagner 1981; Christy 1987) may be
used to hydrate specimens quickly for examination. The formula for the stock
solution is one part di-octyl sodium sulfosuccinate, 24 parts methanol and 75
parts water. A few drops of the stock solution are added to a dropping bottle
of distilled water to make a fine wetting agent. Mounts on microscope slides
for examination at high magnification should, however, always be made in one or
two drops of two percent potassium hydroxide solution. This brings out
characteristic color reactions in leaves and other plant parts. This also helps
remove the operculum (Lauridsen 1972) to reveal the peristome. Heating the
slide with a butane cigarette lighter (if the flame does not touch the glass
slide, there is no soot) aids in loosening the operculum, and placing a cover
slip on the preparation before heating lessens evaporation. If the peristome
persists in being broken off at the base when removal of the peristome is
attempted, allow an intact capsule to soak in a mixture of KOH and Pohlstoffe
15 to 20 minutes or longer. Since Pohlstoffe will eventually precipitate from
basic solutions, one or two drops of a concentrated (4 g in 20 cc water) stock
solution of sodium N-lauroylsarcosine (trade name “Gardol”) may be added to the
bottle of KOH solution as an effective surfactant, although small amounts of
any commercial detergent may substitute.
Taking time to make a good microscopic
preparation will save time later during identification. My standard method is
to place one or two drops of two-percent KOH solution on a microscope slide,
and add a stem of the plant to the slide. The material is cleaned of debris (at
this time one can search for rhizoid-borne propagula) and the laminal color
reaction to the KOH solution is noted. If the material is to be transferred to
an acidic mountant like lactophenol gel (see below), a drop or two of dilute
hydrochloric acid solution is now added. The best preparation consists of one
slide with free leaves that have been removed from the stem, especially near
the apex to reveal the axillary hairs, and sections of the stem and leaf, and a
second slide with the capsule showing the peristome, spores and perichaetial
leaves. Taking care to flip some of the leaves so that the ventral side is
uppermost will save time later. When making permanent slides, two cover slips
may be mounted on each slide with a slide label on the left for consistency.
Permanent slides may be made with Hoyer's
solution (Anderson 1954) or polyvinyl lactophenol (Frahm 1990a), these being
very convenient mountants that need not be luted, especially if sufficient
glycerine is used in formulating Hoyer's solution to counteract dry air in the
storage area.
Many species of Pottiaceae, however, have
large, delicate laminal cells that collapse in Hoyer's solution or polyvinyl
lactophenol. Lactophenol gel (Zander 1983b), which does not collapse the cells
of most species, may be used instead for sensitive species or as a standard
mountant for all species. It has a high index of refraction. The formula for
lactophenol gel is:
30 cc lactic
acid (= 2–hydroxypropanoic acid)
15 g phenol,
crystal (= carbolic acid)
15 cc
distilled water
6 g methyl
cellulose, powder (= cellulose methyl ether, of viscosity 25 cP in 2% solution
or lowest viscosity available)
35 cc
ethylene glycol (= 1,2–ethanediol).
Note, February 3, 2003: This formula is
problematic because it evaporates too fast. Try glycerine jelly (any
microscopic technique book has the formula), which lasts as long as Hoyer's
Solution. If you head a specimen in glycerine jelly, the leaf cells, which
originally collapse, re-inflate and the plants look natural.
Mix the phenol with the lactic acid,
dissolving it with gentle heat. Add the water and stir. Heat to just boiling
(use a fume hood). Add the methyl cellulose powder and stir vigorously into the
hot solution to dissolve (reheat if necessary). Add the ethylene glycol last.
Pour into a glass cylinder and let stand to allow bubbles and undissolved
material to rise. After a day or two, remove any floating particles and pour
the clear liquid into a storage bottle. A small bottle with an applicator wand
built into the lid or a plastic squeeze bottle with a fairly wide aperture (4 µm)
is used to place a drop or two on a slide. Specimens incrusted with carbonate
deposits must first be soaked in a drop or two of dilute hydrochloric acid to
prevent bubble formation in the lactophenol gel.
Specimens soaked in KOH should also be
neutralized with a drop or two of dilute HCl before mounting in lactophenol
gel. A precipitate may appear when overly moist plants are placed in
lactophenol gel but this is redissolved on stirring. Plants may be manipulated,
and leaves and stems sectioned, while in the gel.
The slides prepared as noted above, after
placement of the cover slip, are adequate as semi-permanent mounts for one or
two months. Permanent mounts may be made by sealing the cover slip to the
slide. So as to further solidify the gel and help prevent migration of the
plant material, water portion of the mounting fluid may be allowed to evaporate
somewhat by leaving the freshly mounted slide exposed in a dry place, such as a
fume hood, overnight, but sealing the slide immediately upon preparation is
generally best. To seal the slide, clean cover slips and slides must be used to
insure a complete bond. Clear fingernail polish, although commonly used as a
lutant, is generally inadequate since pyroxylin (nitrocellulose) does not
adhere well to glass.
A good lutant is poly (ethyl methacrylate)
with butyl benzyl phthalate as plasticizer (as KrystalonÔ, Harleco, Gibbstown, NJ 08027 USA), an
artificial balsam with good long-term adherence to glass. Another, more easily obtainable
sealant with excellent adhesion to glass, and good flexibility when dry (but
somewhat reactive, so do not store slides in strong light) is the commercial
liquid “polyurethane” gloss finish for wooden floors, of various formulations
but based mainly on diisocyanates (such as tolylene diisocyanate). Isocyanates
adhere to glass extremely well apparently because they react with an always
present thin film of water strongly adsorbed to glass (Skeist 1962). Keep the
polyurethane container well sealed and, if necessary, add a few drops of
artist's drying retardant to the commercial product to slow the gelling during
polymerization caused by oxidation of linseed oil, a common additive. Any
lutant will stay liquid longer and be more easy to use if kept in a “balsam
bottle” or, better, in a small, disposable applicator bottle. Keep the bottle
more than half full to help exclude air. Also acceptable as glass sealants
include various commercially available silicone rubber glues or caulks, and
“hot-melt glue” applied with a glue gun, but these are difficult to work with
and leave a ridge that may obstruct the high-dry objective lens. Apply lutants
liberally to ensure a good seal; most seals can be scraped off in patches if a
morphological detail is obscured.
Disposable serviettes or wipers are important
to have on hand to clean tools and slides of reagents. A small fan is valuable
at times to disperse any acrid fumes.
Sexing specimens is often required for
identification, but this can be as difficult as sectioning. Poor lighting makes
this more difficult, and one should not be satisfied with only a single
standard illuminator for the dissecting microscope. Dry plants may show
bulbiform perigonia better than moist ones, but plants should be moistened
before removal or manipulation of leaves. Sexuality cannot be determined
accurately without location of antheridia (in perigonia, in antheridiate buds
or naked on the stem, and preferably as observed on several plants in the
collection to weigh degree of variation). Perichaetiate collections entirely
lacking antheridia are often assumed, with some degree of confidence, to be
dioicous, but note Steere's (1940) discussion of dichogamy in Syntrichia
princeps (as Tortula princeps). Perigoniate plants are
best looked for near the periphery of a clump of sporangiate plants. Most
genera show a distinct apical thickening of the plant apex in perigoniate
plants, but this is not always the case. Autoicous buds can be searched for by
carefully stripping leaves of archegoniate plants. Paroicy and synoicy is
determined by attentive dissection of perichaetia with fine forceps under high
magnification under the dissecting microscope with bright illumination.
Antheridiate buds or short perigoniate plants located on soil at the very base
of archegoniate plants are a sign of possible rhizautoicy, but this cannot be
definitely determined without dissection of the rhizoid system or cultivation
of single spores.
Measurements are made with a transparent
plastic metric ruler or a more accurate “Minitool” 5-mm ruler marked in 0.1 µm
increments. An ocular micrometer for microscopic use is of course essential.
Specialists in the taxonomy of particular
groups are often asked by other bryologists to render identifications of
unusual specimens or other puzzling collections. The accepted way to have this
done is to send a “duplicate for identification,” which is a labeled portion of
a collection that the specialist can retain for his/her herbarium. A name for
each specimen sent is then returned to the sender by the specialist. If
duplicates cannot be made, each specimen to be returned should be clearly
marked “unicate.” The cover letter should point out which specimens may be
retained by the specialist and which returned. Generally, a quick response may
be had if only one to three specimens are sent at a time; a package of several
specimens may be placed aside by a specialist pending availability of a larger
block of time for work on them. Identifications of large sets may not be
completed for many months, and, for these, preliminary inquiries of the
specialist should be made. Botanists preparing grant proposals that necessarily
involve more than a little identification work by others should consider
inclusion of support through subcontracting.