IJSEM Papers in Press. Published November 6, 2013 as doi:10.1099/ijs.0.057893-0
Running title: Observation and preparation of ciliates
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An update of “basic light and scanning electron microscopic methods for taxonomic studies of
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ciliated protozoa”
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Wilhelm Foissner
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Universität Salzburg, FB Organismische Biologie, Hellbrunnerstrasse 34, A-5020 Salzburg, Austria
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Content category: Methods
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Correspondence:
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Wilhelm Foissner, FB Organismische Biologie, Universität Salzburg, Hellbrunnerstr. 34,
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A-5020 Salzburg, Austria
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Telephone number: +43 (0)662 8044 5615
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FAX number: +43 (0)662 8044 5698
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e-mail: [email protected]
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Running title: Observation and preparation of ciliates
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This is an update of the review by Foissner (1991). Since then, I improved some methods
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considerably. The following methods are described in detail: live observation, supravital staining with
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methyl green-pyronin, dry silver nitrate impregnation, wet silver nitrate impregnation, silver carbonate
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impregnation, protargol impregnation (three procedures), scanning electron microscopy, and
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deciliation. Familiarity with these methods (or modifications) is a prerequisite for successful
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taxonomic work. No staining method is equally appropriate to all kinds of ciliates. A table is provided
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which indicates those procedures which work best for certain groups of ciliates. A second table relates
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to the structures revealed by the procedures. Good descriptions usually demand at least live
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observation, silver nitrate and protargol or silver carbonate impregnation. Some instructions are
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provided for distinguishing mono- and dikinetids as well as ciliated and non-ciliated basal bodies in
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silvered ciliates. Further, I added a chapter on “Deposition and Labeling of Preparations”. All methods
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work not only with ciliates but also with many other heterotrophic and autotrophic flagellated and
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amoeboid protists. The brilliancy of the silver preparations has tempted some taxonomists to neglect
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live observation. However, many important species characteristics cannot be seen or are changed in
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silvered specimens. I thus consider all species descriptions based exclusively on silver slides as
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incomplete and of doubtful value for both α-taxonomists and ecologists.
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Running title: Observation and preparation of ciliates
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INTRODUCTION
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The micrographs of various freshwater, marine and soil ciliates published by my colleagues and
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myself during the past 25 years, most based on the few methods described here and by Foissner
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(1991), have been widely accepted as being of a high standard. All methods are modifications of
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techniques which initially did not work well in our laboratory, either because they were incompletely
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described or too complicated. This paper provides detailed step-by-step protocols for the methods
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used in our laboratory. For other techniques the reader is referred to the literature cited and to the
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methodological guides by Kirby (1950), Lee et al. (1985) and Dragesco & Dragesco-Kernéis (1986).
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The basic of most methods and the hunger for beautiful protist preparations, I learned from the
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literature cited, from laboratory manuals for the microscopic anatomy of plant and animal tissues (e.g.,
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Adam & Czihak, 1964; Romeis, 1968), and in the laboratories of Dr. N. Wilbert (Bonn University,
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protargol-and wet silver nitrate impregnation) and Prof. Dr. I. Dragesco (Clermont-Ferrand
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University, protargol impregnation). Most improvements were based on hard experimental work or,
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rarely, on good luck. As an example for both, I mention the use of albumized slides (luck) and of a
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chemical developer (many experiments) for the dry silver nitrate method of Klein (1926, 1943). The
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results of these modifications were so convincing that I could publish them in the “Mikrokosmos”, a
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journal for amateur microscopists, when I was just fifteen. Since that, I tried to improve the methods
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almost continuously, the last success being a simple technique for deciliation so that the often
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complex cortical and ciliary structures can be observed with outstanding clarity (Fig, 24–30).
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The photographic documentation profited greatly from the digital revolution. The images can be
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nicely improved and the problem of low focal depth can be overcome by stacking (Fig. 12–16).
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STRUCTURES REVEALED BY THE METHODS DESCRIBED AND INTERPRETATION
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OF SILVER STAINS
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There is no single method which can reveal all details necessary for a good description of a ciliate.
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Likewise, no staining method is equally appropriate to all kinds of ciliates. Good descriptions usually
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demand, as is evident from Tables 1 and 2, at least live observation, silver nitrate and protargol or
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Running title: Observation and preparation of ciliates
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silver carbonate impregnation. The brilliancy of the silver preparations has unfortunately tempted
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some taxonomists to neglect live observation. However, many important species characteristics cannot
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be seen or are changed in silvered specimens such that descriptions based exclusively on silver slides
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are incomplete and of doubtful value for α-taxonomists and ecologists. Especially the latter are usually
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not trained to correlate the silvered structures with the live appearance of the cell. Whenever it is
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possible, important and/or unusual details (e.g., species characters) should be documented not only by
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accurate drawings but also by photographs.
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Several species (e.g., in the genera Pseudoprorodon and Cyrtolophosis) persistently withstand our
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methods, their infraciliature and/or silverline pattern impregnates poorly or not at all. Further
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improvements of existing methods or new techniques should therefore be developed. Stimulating
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ideas can be found in the papers of J. Gelei (1934, 1935), G. Gelei (1939) and Horváth (1938).
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The methods described here work not only with ciliates but also with many heterotrophic and
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autotrophic flagellated and amoeboid protists.
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Usually, silver impregnation is undertaken to reveal the infraciliature (= ciliary pattern) and the
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silverline pattern (= lines revealed by silver nitrate, connecting basal bodies and other cortical
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organelles such as extrusomes and the cytopyge). Extrusomes, various structures associated with the
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basal bodies of the cilia (e.g., parasomal sacs, microtubular ribbons) and several cortical fibrillar
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networks (e.g., the infraciliary lattice) are sometimes also impregnated. This may render interpretation
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of silver stains difficult. It is beyond the scope of this paper to discuss this problem in detail; in fact,
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interpretation is different for each method (and even of slight variations!) and almost for each higher
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systematic category. Thus, I must refer the reader to some key-references (Foissner, 1977, 1981; G.
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Gelei, 1939; J. Gelei, 1934; Hiller, 1991; Hiller & Bardele, 1988; Klein, 1943; Peck, 1974; Tellez et
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al., 1982; Zagon, 1970). However, the differentiation of mono- and dikinetids must be mentioned
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because this is of basic importance for successful taxonomic work.
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Cilia and their basal bodies may be arranged singly (monokinetids) or in pairs (dikinetids); the basal
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bodies may be ciliated or non-ciliated (e.g., Foissner & Foissner, 1988). All silver methods described
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here reveal at least the basal bodies (cilia are usually not impregnated with silver nitrate methods!),
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but not each black or brown granule is a basal body of a cilium! Depending on the procedure used,
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other structures which have a similar size as the basal bodies are also impregnated (e.g., parasomal
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Running title: Observation and preparation of ciliates
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sacs, extrusomes). Thus, the arrangement and the state of the basal bodies cannot be unequivocally
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ascertained with silver methods and transmission electron microscopy is necessary (SEM is
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insufficient as it often does not reveal non-ciliated basal bodies!). The following procedure provides a
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rather reliable differentiation of mono- and dikinetids and of ciliated and non-ciliated basal bodies. (i)
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Study cells carefully in vivo, preferably with interference contrast. With some experience it is easy to
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see whether or not cilia are paired. (ii) Overstain cells with protargol and/or silver carbonate. In
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overimpregnated cells cilia are usually clearly recognizable. (iii) Non-ciliated basal bodies and/or
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parasomal sacs are often slightly smaller than ciliated basal bodies. (iv) Study the electron
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microscopic literature related to the species or group of species under investigation and try to correlate
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the stained structures with ultrastructural features.
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Insert Tables 1 and 2
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METHODS
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Observing living ciliates. Many physical and chemical methods have been described for retarding the
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movement of ciliates in order to observe structural details (for literature, see Foissner, 1991).
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Chemical immobilization (e.g., nickel sulphate) or physical slowing down by increasing the viscosity
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of the medium (e.g., methyl cellulose) are, in our experience, usually unsuitable. These procedures
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often change the shape of the cell or cause premortal alterations of various cell structures. The
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following simple method is therefore preferable: place about 0.5 ml of the raw sample on a slide and
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pick out (collect) the desired specimens with a micropipette (Fig. 1g–l) under a compound microscope
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equipped with a low magnification (e.g., objective 4:1, eyepiece 10×). If specimens are large enough,
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they can be collected from a Petri dish under a dissecting microscope. Working with micropipettes,
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the diameter of which must be adjusted to the size of the specimens, requires some training. The
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collected specimens are now in a very small drop of fluid. Apply small dabs of vaseline (Petroleum
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jelly) to each of the four corners of a coverslip (or on the slide; it is useful to apply the jelly by an
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ordinary syringe with a thick needle). Place this coverslip on the droplet containing the ciliates. Press
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on the vaselined corners with a mounted needle until ciliates become slightly squeezed between slide
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and coverslip (Fig. 1a–d). As the pressure is increasing, the ciliates gradually become less mobile and
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more transparent. Hence, first the location of the main cell organelles (e.g., nuclear and oral apparatus,
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contractile vacuole) and then the details (e.g., extrusomes, micronucleus) can easily be observed under
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low (× 100–400) and high (× 1000, oil immersion objective) magnifications.
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Running title: Observation and preparation of ciliates
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The shape of the cells is of course altered by this procedure. Therefore, specimens taken directly from
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the sample with a large-bore (opening ~ 1 mm) pipette must first be investigated under low
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magnification (× 100–400). Some species are too fragile to withstand handling with the micropipette
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and coverslip trapping without deterioration. Investigation with low magnification also requires some
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experience but it guarantees that undamaged cells are recorded. Video-microscopy is very useful at
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this point of investigation, especially for registrating body shape and swimming behaviour.
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A compound microscope equipped with differential interference contrast is best for observing living
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ciliates. If not available, use bright-field or phase-contrast; the latter is only useful for very flat
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species.
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Insert Fig. 1(a–l) here
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Staining Procedures. Although there are numerous methods for staining ciliates, most of the older
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procedures (e.g., hematoxylin; see Kirby, 1950 for an excellent compilation of protocols) have been
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outdated by silver impregnation techniques and electron microscopy. Various silver stains are
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available, but all need some experience and are usually individually modified. However, familiarity
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with the four silver methods described below is an absolute prerequisite for studying ciliates. These
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are thus described in great detail in order to give even beginners a fair chance to obtain usable slides.
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Apart from silver impregnation, various other staining techniques are useful for taxonomic work with
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ciliates, especially the Feulgen nuclear reaction and supravital staining with methyl green-pyronin in
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order to reveal, respectively, the nuclear apparatus and the mucocysts.
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Simple, viz., molecular formulae are given for the chemicals used, since usually only these are found
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in the catalogues of the suppliers (e.g., Merck). In a laboratory manual it is thus convincing to use this
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style too, instead of the more correct constitutional or structural formulae.
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Five plates of selected micrographs should show examples of excellent preparations. There are two
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ways to do this: either to use a few species and show them treated with all methods described or to
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select many species to give the reader an impression of the diversity. I decided the second way as it is
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possibly more convincing for beginners. However, two species (Colpidium colpoda, Figs 5, 6, 25, 29
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Running title: Observation and preparation of ciliates
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and Paramecium caudatum, Figs 8, 9, 12) are shown prepared with three, and several others with two
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different methods.
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I. Feulgen nuclear reaction. Descriptions of this method are to be found, e.g., in Lee et al. (1985) and
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Dragesco and Dragesco-Kernéis (1986). The Feulgen reaction reveals the nuclear apparatus very
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selectively. I use it only occasionally for α-taxonomic purposes because the nuclear apparatus usually
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stains well with protargol. As protargol often stains various small cytoplasmic inclusions, too, some
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caution is necessary, especially with multimicronucleate species. If in doubt, use the Feulgen reaction
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or an other nuclear staining method, such as that described by Larsen (1975).
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II. Supravital staining with methyl green-pyronin. This simple method is an excellent technique for
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revealing the mucocysts of many ciliates (those of tetrahymenids and rather many haptorids, however,
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usually do not stain). Mucocysts are stained deeply and very selectively blue or red, and can be
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observed in various stages of swelling because the cells are not killed instantly. The nuclear apparatus
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is also stained. Examples: Fig. 10 and Figs 8, 17, 38 in Foissner (1991).
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P r o c e d u r e (after Foissner, 1979)
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1.
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Pick out the desired ciliates with a micropipette and place the very small drop of fluid in the centre of a slide.
2.
Add an equally sized drop of methyl green-pyronin and mix the two drops gently by swiveling the
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slide or with a needle.
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R e m a r k s : If ciliates were already mounted under the coverslip, add a small drop of dye at one
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edge of the coverslip and pass it through the preparation with a piece of filter paper placed at the
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opposite end of the coverslip.
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3.
Place a coverslip with vaselined corners on the preparation and squeeze specimens slightly.
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R e m a r k s : Observe immediately. Cells die in the stain within some seconds or minutes.
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Mucocysts stain very quickly and many can be observed at various stages of swelling. To reveal
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the nuclear apparatus, cells should be fairly strongly squashed (= flattened). The preparation is
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temporary. After 5–10 minutes the cytoplasm often becomes heavily stained and obscures other
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details.
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Reagents
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Running title: Observation and preparation of ciliates
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1 g methyl green-pyronin (Chroma-company, Küferstrasse 2, P.O. Box 1110, D-73257 Köngen,
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Germany)
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ad 100 ml distilled water and filter
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R e m a r k s : This solution is very stable and can be used for years
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III. The “dry”silver nitrate methods. Because of the numerous problems with the basic dry Klein
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(1926, 1958) technique, Foissner (1976) and others (e.g., J. Gelei, 1934; Ruzicka, 1966) introduced
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some improvements. The dry methods (“dry” because cells are air-dried and not chemically fixed
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before being treated with silver nitrate) provide preliminary information on the somatic and oral
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infraciliature (= ciliary pattern) and are often best for revealing the silverline pattern (= lines revealed
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by silver nitrate and connecting basal bodies and other cortical organelles such as extrusomes and the
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cytopyge). Although the results vary highly, the method is worthwhile because it is quick and often
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produces excellent preparations, which can be well documented since the cells are flattened during
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dehydration. Only cortical structures are revealed. Examples: Fig. 8 and Figs 15, 19, 20, 23, 26, 35, 36
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in Foissner (1991).
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P r o c e d u r e (after Foissner, 1976 and recent experience)
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1.
Take 5–10 clean slides and spread a very thin layer of albumen over the middle third of each with
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a finger-tip. Dry for at least 1 minute.
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R e m a r k s : The egg-albumen (remove germinal disk! do not add glycerol) must have been kept
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open in a wide-necked flask for at least 20 hours; fresh albumen is often less satisfactory. It can
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be used for 2–3 days if the flask is subsequently sealed; do not, however, stir before use but skim
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the albumen from the surface with a finger-tip. To facilitate spreading breathe on slide so that a
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film of condensation is produced on which the albumen can glide. The albumen layer must be
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very thin and uniform but should not cover the cells.
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2.
Place a drop of fluid containing the ciliates on the albumized slide, spread with a needle (do not
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touch albumen layer!), and dry the preparation at room temperature.
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R e m a r k s : Even single specimens can be placed on the albumized slide with a micropipette. If
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necessary enrich ciliates by gentle centrifugation or by leaving sample to settle for a few hours,
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after which time oxygen depletion induces many ciliates to move to the water surface. The
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amount and chemical composition of the fluid with which the ciliates are air-dried as well as
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temperature and air humidity greatly influence the results. Therefore, 5–10 slides should usually
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Running title: Observation and preparation of ciliates
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be prepared simultaneously to vary these parameters, e.g. by washing cells with different amounts
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of distilled water or fresh culture medium. Washing cells with distilled water or spreading the
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drop to a very thin film is especially recommended with saline fluids, e.g. seawater, sewage, and
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soil. Temperature and humidity are varied using an ordinary hair-dryer.
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3.
Apply some drops of silver nitrate to the dried material for about 1 minute.
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R e m a r k s : Do not touch albumen layer with the pipette. The reaction time does not influence
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the results; a minute is adequate to impregnate and fix the cells on the slide.
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4.
Wash slide for about 3 seconds with distilled water and re-dry.
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R e m a r k s : Wash gently! Apply water current from the top third of the tilted slide so that water
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runs gently over the dried material. Leave slide tilted during drying.
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5.
Pre-develop the dried slide by exposing it for 5–60 seconds to a 40–60 watt electric bulb at a
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distance of 3–10 cm.
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R e m a r k s : Time and distance influence intensity of impregnation (see also next step).
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6.
Apply a few drops of developer to the dried preparation for about 60 seconds.
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R e m a r k s : The pre-development (step 5), the composition of the developer, and the material
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itself influence impregnation intensity and quality. The best ratio of these parameters must be
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evaluated in pilot experiments. If the developer is well adjusted, the albumen around the dried
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fluid turns brown-black; if the developer is too strong, the albumen appears black (add some
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component A [see Reagents] and/or reduce pre-developing time); if the developer is too weak, the
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albumen appears brownish or yellowish (add some components B and/or C and/or increase pre-
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development time).
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7.
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Pour developer off slide, rinse gently in tap water for 5–10 seconds, and immerse it in the fixative (sodium thiosulfate) for 5 minutes.
8.
Remove slide from fixative and rinse gently in tap water for 10 minutes changing the water three
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times.
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R e m a r k s : The fixative must be thoroughly removed to avoid bleaching of the impregnation.
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Preparations usually fade within a few weeks when the silver nitrate is reduced only by sunlight
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or an UV-lamp. Do not use distilled water, otherwise cells swell and eventually detach from the
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slide!
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9.
Air-dry the slide again and mount in synthetic neutral medium (e.g., Eukitt, Euparal).
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Running title: Observation and preparation of ciliates
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R e m a r k s : Slides should be tilted during drying. Mounting medium should be of medium
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viscosity. The preparation is stable, providing good fixation and careful elimination of the
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fixative.
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Reagents
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a) Silver nitrate solution (long term stability in brown flask)
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1 g silver nitrate (AgN03)
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ad 100 ml distilled water
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b) Developer (stable for about 1–3 days; must be renewed as soon as it turns dark brown or shows
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crystals; mix components in the sequence indicated)
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20 ml solution A
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1 ml solution B
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1 ml solution C
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Solution A (this is an ordinary developer for negatives; dissolve ingredients in the sequence
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indicated; stable for a year in brown bottle)
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1000 ml hot tap water (about 40 °C)
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10 g boric acid (H3B03)
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10 g borax (Na2B40–)
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5 g hydroquinone (C6H6O2)
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100 g anhydrous sodium sulfite (Na2SO3)
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2.5 g metol = methylamino-phenol-sulfate = (CH3NHC6H4OH)2 • H2SO4
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Solution B (this is a concentrated photographic paper developer; dissolve ingredients in the
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sequence indicated; stable for about 6 months when stored in fully filled cups
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100 ml distilled water
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0.4 g metol = methylamino-phenol-sulfate = (CH3NHC6H4OH)2 • H2SO4
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5.2 g anhydrous sodium sulfite (Na2SO3)
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1.2 g hydroquinone (C6H6O2)
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10.4 g sodium carbonate (Na2CO3)
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10.4 g potassium carbonate (K2CO3)
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0.4 g potassium bromide (KBr)
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Solution C (stable for several years)
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10 g sodium hydroxide (NaOH)
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Running title: Observation and preparation of ciliates
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ad 100 ml distilled water
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c) Fixative for impregnation (stable for years)
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50 g sodium thiosulfate (Na2S2O3 • 5 H2O)
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ad 1,000 ml distilled water
308 309
IV. The “wet” silver nitrate methods. The first wet (“wet” because cells are chemically fixed before
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being treated with silver nitrate) method was described by Chatton and Lwoff (1930, 1936). The
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technique became well known after Corliss (1953) published the version in use in the Paris laboratory
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of Fauré-Fremiet. It works well with many kinds of limnetic and marine ciliates, especially with
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hymenostomes (e.g., Tetrahymena, Paramecium, Cyclidium), holophryids (e.g., Holophrya), most
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colpodids (e.g., Colpoda, Bresslauides), and some hypotrichs (e.g., Euplotes). Less convincing results
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are usually obtained with peritrichs (e.g., Vorticella), heterotrichs (e.g., Spirostomum, Metopus),
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oligotrichs (e.g., Halteria), and most hypotrichs (e.g., Oxytricha, Urostyla). The wet methods provide
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valuable information on the somatic and oral infraciliature as well as the silverlines, which are,
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however, often rather faintly stained. The shape of the cells is usually well preserved, which is of
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advantage to the investigation but makes photographic documentation difficult. As with the dry
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methods, only cortical structures are revealed. Examples: Figs 11, 12–16 and Figs 7, 10, 16, 28, 29 in
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Foissner (1991). Modifications have been described by, e.g., Chatton (1940), Frankel & Heckmann
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(1968), Lynn et al. (1981) and Roberts & Causton (1988), who investigated some variables in detail.
323 324
The method described in 1991 and likely all modifications have several problems: (i) the gelatin, in
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which the specimens are embedded, is too weak, i.e., does not become sufficiently solid, and the
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preparations are lost or full of clouds; (ii) the gelatin becomes cloudy and/or gets sharp fissures above
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the individual cells, even if “good” gelatin is used; (iii) the impregnation is too faint, especially on the
328
“back side”, that is the side oriented to the microscope slide; and (iv) the impregnation bleaches more
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or less strongly within a few days or months.
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Problem (i) can be solved by using “Gelatin, from Bovine Bone”, Wako company, Japan. Problem (ii)
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occurs when the preparation becomes too warm, i.e., when the gelatin commences melting. Thus, keep
333
the preparation < 10 °C throughout the procedure. Problem (iii) is partially caused by the inability of
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the UV-light to penetrate sufficiently the gelatin and the cells. Thus, silver reduction must be done
335
from above and below (see step 12) and chemical development should follow UV reduction. The
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Running title: Observation and preparation of ciliates
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fourth problem, I solved only recently (see steps 13–15). It makes the protocol rather complex but is
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worth to be done because stable preparations can be obtained. Several slides should be prepared
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simultaneously from the same material. If only few specimens are available, these must be handled
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with micropipettes during steps 1–7 (difficult task!); for ample material a centrifuge may be used.
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Until dehydration (step 19), keep all solutions cold (< 10 °C) as warming causes clouds or even
341
detaches the gelatin from the slide. The Da Fano solution is of paramount significance because it
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decides about the strength of the impregnation. If too much is used, precipitations may develop; if too
343
less, the impregnation may become too faint. The method is not simple and requires experience. Since
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some steps must be done quickly, it is necessary to be well organized.
345 346
Procedure
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1.
348 349
If possible, concentrate ciliates by gentle centrifugation (the fixative is expensive) or collect individual specimens and drop them into the fixative.
2.
Put ciliates into Champy’s fluid and fix for 1–30 minutes.
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R e m a r k s : The ratio of material to fixative should be at least 1:1, better 1:2. The fixation time
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apparently does not influence the results greatly. We usually fix for about 10 minutes. Fixation
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should be carried out in a fume hood since osmic vapours are highly toxic.
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3.
Remove fixative by centrifugation or micropipette and postfix in Da Fano’s fluid for at least 5
354
minutes. Continue this replacement until the solution is the colour of Da Fano’s fluid (2–3 times
355
are usually enough).
356
R e m a r k s : Material can be stored in Da Fano’s fluid for years.
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4.
Place a very clean, grease-free slide on a hot-plate (35–60 °C).
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R e m a r k s : The slides must be grease-free (clean with alcohol and flame); even commercially
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pre-cleaned slides should be cleaned with a alcohol-moist cloth.
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5.
Place a small piece (about 2–4 mm in diameter) of gelatin in the centre of the warmed slide and
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allow to melt.
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R e m a r k s : Gelatin should have been stored in the refrigerator for at least one week before use.
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Fresh gelatin may cause cloudy silver deposits.
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6.
Quickly add an equal-sized or smaller drop of concentrated specimens to the molten gelatin and
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remove slide from hot-plate.
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R e m a r k s : Use as few Da Fano’s fluid as possible. Mix organisms thoroughly into the gelatin
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using a mounted needle.
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Running title: Observation and preparation of ciliates
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7.
Quickly spread the drop on the slide or remove excess fluid under the dissecting microscope with
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a warmed micropipette until ciliates remain just nicely embedded in a thin gelatin layer.
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R e m a r k s : Steps 6 and 7 must be done quickly to avoid hardening and/or desiccation of the
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gelatin; if gelatin solidifies during the procedure return the slide to the hot-plate for a few
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seconds. Excess fluid can be removed only if ciliates are large or very numerous. For small (
Running title: Observation and preparation of ciliates
1 2 3 4
An update of “basic light and scanning electron microscopic methods for taxonomic studies of
5
ciliated protozoa”
6 7
Wilhelm Foissner
8
Universität Salzburg, FB Organismische Biologie, Hellbrunnerstrasse 34, A-5020 Salzburg, Austria
9 10 11 12 13 14 15 16 17 18 19
Content category: Methods
20 21 22 23 24 25
Correspondence:
26
Wilhelm Foissner, FB Organismische Biologie, Universität Salzburg, Hellbrunnerstr. 34,
27
A-5020 Salzburg, Austria
28
Telephone number: +43 (0)662 8044 5615
29
FAX number: +43 (0)662 8044 5698
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e-mail: [email protected]
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1
Running title: Observation and preparation of ciliates
32 33
This is an update of the review by Foissner (1991). Since then, I improved some methods
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considerably. The following methods are described in detail: live observation, supravital staining with
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methyl green-pyronin, dry silver nitrate impregnation, wet silver nitrate impregnation, silver carbonate
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impregnation, protargol impregnation (three procedures), scanning electron microscopy, and
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deciliation. Familiarity with these methods (or modifications) is a prerequisite for successful
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taxonomic work. No staining method is equally appropriate to all kinds of ciliates. A table is provided
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which indicates those procedures which work best for certain groups of ciliates. A second table relates
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to the structures revealed by the procedures. Good descriptions usually demand at least live
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observation, silver nitrate and protargol or silver carbonate impregnation. Some instructions are
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provided for distinguishing mono- and dikinetids as well as ciliated and non-ciliated basal bodies in
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silvered ciliates. Further, I added a chapter on “Deposition and Labeling of Preparations”. All methods
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work not only with ciliates but also with many other heterotrophic and autotrophic flagellated and
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amoeboid protists. The brilliancy of the silver preparations has tempted some taxonomists to neglect
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live observation. However, many important species characteristics cannot be seen or are changed in
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silvered specimens. I thus consider all species descriptions based exclusively on silver slides as
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incomplete and of doubtful value for both α-taxonomists and ecologists.
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2
Running title: Observation and preparation of ciliates
51
INTRODUCTION
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The micrographs of various freshwater, marine and soil ciliates published by my colleagues and
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myself during the past 25 years, most based on the few methods described here and by Foissner
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(1991), have been widely accepted as being of a high standard. All methods are modifications of
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techniques which initially did not work well in our laboratory, either because they were incompletely
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described or too complicated. This paper provides detailed step-by-step protocols for the methods
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used in our laboratory. For other techniques the reader is referred to the literature cited and to the
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methodological guides by Kirby (1950), Lee et al. (1985) and Dragesco & Dragesco-Kernéis (1986).
60 61
The basic of most methods and the hunger for beautiful protist preparations, I learned from the
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literature cited, from laboratory manuals for the microscopic anatomy of plant and animal tissues (e.g.,
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Adam & Czihak, 1964; Romeis, 1968), and in the laboratories of Dr. N. Wilbert (Bonn University,
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protargol-and wet silver nitrate impregnation) and Prof. Dr. I. Dragesco (Clermont-Ferrand
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University, protargol impregnation). Most improvements were based on hard experimental work or,
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rarely, on good luck. As an example for both, I mention the use of albumized slides (luck) and of a
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chemical developer (many experiments) for the dry silver nitrate method of Klein (1926, 1943). The
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results of these modifications were so convincing that I could publish them in the “Mikrokosmos”, a
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journal for amateur microscopists, when I was just fifteen. Since that, I tried to improve the methods
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almost continuously, the last success being a simple technique for deciliation so that the often
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complex cortical and ciliary structures can be observed with outstanding clarity (Fig, 24–30).
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The photographic documentation profited greatly from the digital revolution. The images can be
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nicely improved and the problem of low focal depth can be overcome by stacking (Fig. 12–16).
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STRUCTURES REVEALED BY THE METHODS DESCRIBED AND INTERPRETATION
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OF SILVER STAINS
79 80
There is no single method which can reveal all details necessary for a good description of a ciliate.
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Likewise, no staining method is equally appropriate to all kinds of ciliates. Good descriptions usually
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demand, as is evident from Tables 1 and 2, at least live observation, silver nitrate and protargol or
3
Running title: Observation and preparation of ciliates
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silver carbonate impregnation. The brilliancy of the silver preparations has unfortunately tempted
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some taxonomists to neglect live observation. However, many important species characteristics cannot
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be seen or are changed in silvered specimens such that descriptions based exclusively on silver slides
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are incomplete and of doubtful value for α-taxonomists and ecologists. Especially the latter are usually
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not trained to correlate the silvered structures with the live appearance of the cell. Whenever it is
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possible, important and/or unusual details (e.g., species characters) should be documented not only by
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accurate drawings but also by photographs.
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Several species (e.g., in the genera Pseudoprorodon and Cyrtolophosis) persistently withstand our
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methods, their infraciliature and/or silverline pattern impregnates poorly or not at all. Further
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improvements of existing methods or new techniques should therefore be developed. Stimulating
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ideas can be found in the papers of J. Gelei (1934, 1935), G. Gelei (1939) and Horváth (1938).
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The methods described here work not only with ciliates but also with many heterotrophic and
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autotrophic flagellated and amoeboid protists.
97 98
Usually, silver impregnation is undertaken to reveal the infraciliature (= ciliary pattern) and the
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silverline pattern (= lines revealed by silver nitrate, connecting basal bodies and other cortical
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organelles such as extrusomes and the cytopyge). Extrusomes, various structures associated with the
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basal bodies of the cilia (e.g., parasomal sacs, microtubular ribbons) and several cortical fibrillar
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networks (e.g., the infraciliary lattice) are sometimes also impregnated. This may render interpretation
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of silver stains difficult. It is beyond the scope of this paper to discuss this problem in detail; in fact,
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interpretation is different for each method (and even of slight variations!) and almost for each higher
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systematic category. Thus, I must refer the reader to some key-references (Foissner, 1977, 1981; G.
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Gelei, 1939; J. Gelei, 1934; Hiller, 1991; Hiller & Bardele, 1988; Klein, 1943; Peck, 1974; Tellez et
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al., 1982; Zagon, 1970). However, the differentiation of mono- and dikinetids must be mentioned
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because this is of basic importance for successful taxonomic work.
109 110
Cilia and their basal bodies may be arranged singly (monokinetids) or in pairs (dikinetids); the basal
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bodies may be ciliated or non-ciliated (e.g., Foissner & Foissner, 1988). All silver methods described
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here reveal at least the basal bodies (cilia are usually not impregnated with silver nitrate methods!),
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but not each black or brown granule is a basal body of a cilium! Depending on the procedure used,
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other structures which have a similar size as the basal bodies are also impregnated (e.g., parasomal
4
Running title: Observation and preparation of ciliates
115
sacs, extrusomes). Thus, the arrangement and the state of the basal bodies cannot be unequivocally
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ascertained with silver methods and transmission electron microscopy is necessary (SEM is
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insufficient as it often does not reveal non-ciliated basal bodies!). The following procedure provides a
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rather reliable differentiation of mono- and dikinetids and of ciliated and non-ciliated basal bodies. (i)
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Study cells carefully in vivo, preferably with interference contrast. With some experience it is easy to
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see whether or not cilia are paired. (ii) Overstain cells with protargol and/or silver carbonate. In
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overimpregnated cells cilia are usually clearly recognizable. (iii) Non-ciliated basal bodies and/or
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parasomal sacs are often slightly smaller than ciliated basal bodies. (iv) Study the electron
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microscopic literature related to the species or group of species under investigation and try to correlate
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the stained structures with ultrastructural features.
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Insert Tables 1 and 2
126 127
METHODS
128 129
Observing living ciliates. Many physical and chemical methods have been described for retarding the
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movement of ciliates in order to observe structural details (for literature, see Foissner, 1991).
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Chemical immobilization (e.g., nickel sulphate) or physical slowing down by increasing the viscosity
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of the medium (e.g., methyl cellulose) are, in our experience, usually unsuitable. These procedures
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often change the shape of the cell or cause premortal alterations of various cell structures. The
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following simple method is therefore preferable: place about 0.5 ml of the raw sample on a slide and
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pick out (collect) the desired specimens with a micropipette (Fig. 1g–l) under a compound microscope
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equipped with a low magnification (e.g., objective 4:1, eyepiece 10×). If specimens are large enough,
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they can be collected from a Petri dish under a dissecting microscope. Working with micropipettes,
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the diameter of which must be adjusted to the size of the specimens, requires some training. The
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collected specimens are now in a very small drop of fluid. Apply small dabs of vaseline (Petroleum
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jelly) to each of the four corners of a coverslip (or on the slide; it is useful to apply the jelly by an
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ordinary syringe with a thick needle). Place this coverslip on the droplet containing the ciliates. Press
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on the vaselined corners with a mounted needle until ciliates become slightly squeezed between slide
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and coverslip (Fig. 1a–d). As the pressure is increasing, the ciliates gradually become less mobile and
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more transparent. Hence, first the location of the main cell organelles (e.g., nuclear and oral apparatus,
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contractile vacuole) and then the details (e.g., extrusomes, micronucleus) can easily be observed under
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low (× 100–400) and high (× 1000, oil immersion objective) magnifications.
5
Running title: Observation and preparation of ciliates
147 148
The shape of the cells is of course altered by this procedure. Therefore, specimens taken directly from
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the sample with a large-bore (opening ~ 1 mm) pipette must first be investigated under low
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magnification (× 100–400). Some species are too fragile to withstand handling with the micropipette
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and coverslip trapping without deterioration. Investigation with low magnification also requires some
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experience but it guarantees that undamaged cells are recorded. Video-microscopy is very useful at
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this point of investigation, especially for registrating body shape and swimming behaviour.
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A compound microscope equipped with differential interference contrast is best for observing living
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ciliates. If not available, use bright-field or phase-contrast; the latter is only useful for very flat
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species.
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Insert Fig. 1(a–l) here
158 159
Staining Procedures. Although there are numerous methods for staining ciliates, most of the older
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procedures (e.g., hematoxylin; see Kirby, 1950 for an excellent compilation of protocols) have been
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outdated by silver impregnation techniques and electron microscopy. Various silver stains are
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available, but all need some experience and are usually individually modified. However, familiarity
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with the four silver methods described below is an absolute prerequisite for studying ciliates. These
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are thus described in great detail in order to give even beginners a fair chance to obtain usable slides.
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Apart from silver impregnation, various other staining techniques are useful for taxonomic work with
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ciliates, especially the Feulgen nuclear reaction and supravital staining with methyl green-pyronin in
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order to reveal, respectively, the nuclear apparatus and the mucocysts.
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Simple, viz., molecular formulae are given for the chemicals used, since usually only these are found
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in the catalogues of the suppliers (e.g., Merck). In a laboratory manual it is thus convincing to use this
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style too, instead of the more correct constitutional or structural formulae.
173 174
Five plates of selected micrographs should show examples of excellent preparations. There are two
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ways to do this: either to use a few species and show them treated with all methods described or to
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select many species to give the reader an impression of the diversity. I decided the second way as it is
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possibly more convincing for beginners. However, two species (Colpidium colpoda, Figs 5, 6, 25, 29
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Running title: Observation and preparation of ciliates
178
and Paramecium caudatum, Figs 8, 9, 12) are shown prepared with three, and several others with two
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different methods.
180 181
I. Feulgen nuclear reaction. Descriptions of this method are to be found, e.g., in Lee et al. (1985) and
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Dragesco and Dragesco-Kernéis (1986). The Feulgen reaction reveals the nuclear apparatus very
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selectively. I use it only occasionally for α-taxonomic purposes because the nuclear apparatus usually
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stains well with protargol. As protargol often stains various small cytoplasmic inclusions, too, some
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caution is necessary, especially with multimicronucleate species. If in doubt, use the Feulgen reaction
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or an other nuclear staining method, such as that described by Larsen (1975).
187 188
II. Supravital staining with methyl green-pyronin. This simple method is an excellent technique for
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revealing the mucocysts of many ciliates (those of tetrahymenids and rather many haptorids, however,
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usually do not stain). Mucocysts are stained deeply and very selectively blue or red, and can be
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observed in various stages of swelling because the cells are not killed instantly. The nuclear apparatus
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is also stained. Examples: Fig. 10 and Figs 8, 17, 38 in Foissner (1991).
193 194
P r o c e d u r e (after Foissner, 1979)
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1.
196 197
Pick out the desired ciliates with a micropipette and place the very small drop of fluid in the centre of a slide.
2.
Add an equally sized drop of methyl green-pyronin and mix the two drops gently by swiveling the
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slide or with a needle.
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R e m a r k s : If ciliates were already mounted under the coverslip, add a small drop of dye at one
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edge of the coverslip and pass it through the preparation with a piece of filter paper placed at the
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opposite end of the coverslip.
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3.
Place a coverslip with vaselined corners on the preparation and squeeze specimens slightly.
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R e m a r k s : Observe immediately. Cells die in the stain within some seconds or minutes.
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Mucocysts stain very quickly and many can be observed at various stages of swelling. To reveal
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the nuclear apparatus, cells should be fairly strongly squashed (= flattened). The preparation is
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temporary. After 5–10 minutes the cytoplasm often becomes heavily stained and obscures other
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details.
208 209
Reagents
7
Running title: Observation and preparation of ciliates
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1 g methyl green-pyronin (Chroma-company, Küferstrasse 2, P.O. Box 1110, D-73257 Köngen,
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Germany)
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ad 100 ml distilled water and filter
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R e m a r k s : This solution is very stable and can be used for years
214 215
III. The “dry”silver nitrate methods. Because of the numerous problems with the basic dry Klein
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(1926, 1958) technique, Foissner (1976) and others (e.g., J. Gelei, 1934; Ruzicka, 1966) introduced
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some improvements. The dry methods (“dry” because cells are air-dried and not chemically fixed
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before being treated with silver nitrate) provide preliminary information on the somatic and oral
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infraciliature (= ciliary pattern) and are often best for revealing the silverline pattern (= lines revealed
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by silver nitrate and connecting basal bodies and other cortical organelles such as extrusomes and the
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cytopyge). Although the results vary highly, the method is worthwhile because it is quick and often
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produces excellent preparations, which can be well documented since the cells are flattened during
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dehydration. Only cortical structures are revealed. Examples: Fig. 8 and Figs 15, 19, 20, 23, 26, 35, 36
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in Foissner (1991).
225 226
P r o c e d u r e (after Foissner, 1976 and recent experience)
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1.
Take 5–10 clean slides and spread a very thin layer of albumen over the middle third of each with
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a finger-tip. Dry for at least 1 minute.
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R e m a r k s : The egg-albumen (remove germinal disk! do not add glycerol) must have been kept
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open in a wide-necked flask for at least 20 hours; fresh albumen is often less satisfactory. It can
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be used for 2–3 days if the flask is subsequently sealed; do not, however, stir before use but skim
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the albumen from the surface with a finger-tip. To facilitate spreading breathe on slide so that a
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film of condensation is produced on which the albumen can glide. The albumen layer must be
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very thin and uniform but should not cover the cells.
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2.
Place a drop of fluid containing the ciliates on the albumized slide, spread with a needle (do not
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touch albumen layer!), and dry the preparation at room temperature.
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R e m a r k s : Even single specimens can be placed on the albumized slide with a micropipette. If
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necessary enrich ciliates by gentle centrifugation or by leaving sample to settle for a few hours,
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after which time oxygen depletion induces many ciliates to move to the water surface. The
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amount and chemical composition of the fluid with which the ciliates are air-dried as well as
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temperature and air humidity greatly influence the results. Therefore, 5–10 slides should usually
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Running title: Observation and preparation of ciliates
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be prepared simultaneously to vary these parameters, e.g. by washing cells with different amounts
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of distilled water or fresh culture medium. Washing cells with distilled water or spreading the
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drop to a very thin film is especially recommended with saline fluids, e.g. seawater, sewage, and
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soil. Temperature and humidity are varied using an ordinary hair-dryer.
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3.
Apply some drops of silver nitrate to the dried material for about 1 minute.
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R e m a r k s : Do not touch albumen layer with the pipette. The reaction time does not influence
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the results; a minute is adequate to impregnate and fix the cells on the slide.
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4.
Wash slide for about 3 seconds with distilled water and re-dry.
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R e m a r k s : Wash gently! Apply water current from the top third of the tilted slide so that water
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runs gently over the dried material. Leave slide tilted during drying.
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5.
Pre-develop the dried slide by exposing it for 5–60 seconds to a 40–60 watt electric bulb at a
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distance of 3–10 cm.
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R e m a r k s : Time and distance influence intensity of impregnation (see also next step).
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6.
Apply a few drops of developer to the dried preparation for about 60 seconds.
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R e m a r k s : The pre-development (step 5), the composition of the developer, and the material
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itself influence impregnation intensity and quality. The best ratio of these parameters must be
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evaluated in pilot experiments. If the developer is well adjusted, the albumen around the dried
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fluid turns brown-black; if the developer is too strong, the albumen appears black (add some
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component A [see Reagents] and/or reduce pre-developing time); if the developer is too weak, the
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albumen appears brownish or yellowish (add some components B and/or C and/or increase pre-
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development time).
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7.
264 265
Pour developer off slide, rinse gently in tap water for 5–10 seconds, and immerse it in the fixative (sodium thiosulfate) for 5 minutes.
8.
Remove slide from fixative and rinse gently in tap water for 10 minutes changing the water three
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times.
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R e m a r k s : The fixative must be thoroughly removed to avoid bleaching of the impregnation.
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Preparations usually fade within a few weeks when the silver nitrate is reduced only by sunlight
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or an UV-lamp. Do not use distilled water, otherwise cells swell and eventually detach from the
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slide!
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9.
Air-dry the slide again and mount in synthetic neutral medium (e.g., Eukitt, Euparal).
9
Running title: Observation and preparation of ciliates
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R e m a r k s : Slides should be tilted during drying. Mounting medium should be of medium
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viscosity. The preparation is stable, providing good fixation and careful elimination of the
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fixative.
275 276
Reagents
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a) Silver nitrate solution (long term stability in brown flask)
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1 g silver nitrate (AgN03)
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ad 100 ml distilled water
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b) Developer (stable for about 1–3 days; must be renewed as soon as it turns dark brown or shows
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crystals; mix components in the sequence indicated)
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20 ml solution A
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1 ml solution B
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1 ml solution C
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Solution A (this is an ordinary developer for negatives; dissolve ingredients in the sequence
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indicated; stable for a year in brown bottle)
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1000 ml hot tap water (about 40 °C)
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10 g boric acid (H3B03)
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10 g borax (Na2B40–)
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5 g hydroquinone (C6H6O2)
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100 g anhydrous sodium sulfite (Na2SO3)
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2.5 g metol = methylamino-phenol-sulfate = (CH3NHC6H4OH)2 • H2SO4
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Solution B (this is a concentrated photographic paper developer; dissolve ingredients in the
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sequence indicated; stable for about 6 months when stored in fully filled cups
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100 ml distilled water
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0.4 g metol = methylamino-phenol-sulfate = (CH3NHC6H4OH)2 • H2SO4
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5.2 g anhydrous sodium sulfite (Na2SO3)
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1.2 g hydroquinone (C6H6O2)
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10.4 g sodium carbonate (Na2CO3)
300
10.4 g potassium carbonate (K2CO3)
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0.4 g potassium bromide (KBr)
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Solution C (stable for several years)
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10 g sodium hydroxide (NaOH)
10
Running title: Observation and preparation of ciliates
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ad 100 ml distilled water
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c) Fixative for impregnation (stable for years)
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50 g sodium thiosulfate (Na2S2O3 • 5 H2O)
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ad 1,000 ml distilled water
308 309
IV. The “wet” silver nitrate methods. The first wet (“wet” because cells are chemically fixed before
310
being treated with silver nitrate) method was described by Chatton and Lwoff (1930, 1936). The
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technique became well known after Corliss (1953) published the version in use in the Paris laboratory
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of Fauré-Fremiet. It works well with many kinds of limnetic and marine ciliates, especially with
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hymenostomes (e.g., Tetrahymena, Paramecium, Cyclidium), holophryids (e.g., Holophrya), most
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colpodids (e.g., Colpoda, Bresslauides), and some hypotrichs (e.g., Euplotes). Less convincing results
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are usually obtained with peritrichs (e.g., Vorticella), heterotrichs (e.g., Spirostomum, Metopus),
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oligotrichs (e.g., Halteria), and most hypotrichs (e.g., Oxytricha, Urostyla). The wet methods provide
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valuable information on the somatic and oral infraciliature as well as the silverlines, which are,
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however, often rather faintly stained. The shape of the cells is usually well preserved, which is of
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advantage to the investigation but makes photographic documentation difficult. As with the dry
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methods, only cortical structures are revealed. Examples: Figs 11, 12–16 and Figs 7, 10, 16, 28, 29 in
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Foissner (1991). Modifications have been described by, e.g., Chatton (1940), Frankel & Heckmann
322
(1968), Lynn et al. (1981) and Roberts & Causton (1988), who investigated some variables in detail.
323 324
The method described in 1991 and likely all modifications have several problems: (i) the gelatin, in
325
which the specimens are embedded, is too weak, i.e., does not become sufficiently solid, and the
326
preparations are lost or full of clouds; (ii) the gelatin becomes cloudy and/or gets sharp fissures above
327
the individual cells, even if “good” gelatin is used; (iii) the impregnation is too faint, especially on the
328
“back side”, that is the side oriented to the microscope slide; and (iv) the impregnation bleaches more
329
or less strongly within a few days or months.
330 331
Problem (i) can be solved by using “Gelatin, from Bovine Bone”, Wako company, Japan. Problem (ii)
332
occurs when the preparation becomes too warm, i.e., when the gelatin commences melting. Thus, keep
333
the preparation < 10 °C throughout the procedure. Problem (iii) is partially caused by the inability of
334
the UV-light to penetrate sufficiently the gelatin and the cells. Thus, silver reduction must be done
335
from above and below (see step 12) and chemical development should follow UV reduction. The
11
Running title: Observation and preparation of ciliates
336
fourth problem, I solved only recently (see steps 13–15). It makes the protocol rather complex but is
337
worth to be done because stable preparations can be obtained. Several slides should be prepared
338
simultaneously from the same material. If only few specimens are available, these must be handled
339
with micropipettes during steps 1–7 (difficult task!); for ample material a centrifuge may be used.
340
Until dehydration (step 19), keep all solutions cold (< 10 °C) as warming causes clouds or even
341
detaches the gelatin from the slide. The Da Fano solution is of paramount significance because it
342
decides about the strength of the impregnation. If too much is used, precipitations may develop; if too
343
less, the impregnation may become too faint. The method is not simple and requires experience. Since
344
some steps must be done quickly, it is necessary to be well organized.
345 346
Procedure
347
1.
348 349
If possible, concentrate ciliates by gentle centrifugation (the fixative is expensive) or collect individual specimens and drop them into the fixative.
2.
Put ciliates into Champy’s fluid and fix for 1–30 minutes.
350
R e m a r k s : The ratio of material to fixative should be at least 1:1, better 1:2. The fixation time
351
apparently does not influence the results greatly. We usually fix for about 10 minutes. Fixation
352
should be carried out in a fume hood since osmic vapours are highly toxic.
353
3.
Remove fixative by centrifugation or micropipette and postfix in Da Fano’s fluid for at least 5
354
minutes. Continue this replacement until the solution is the colour of Da Fano’s fluid (2–3 times
355
are usually enough).
356
R e m a r k s : Material can be stored in Da Fano’s fluid for years.
357
4.
Place a very clean, grease-free slide on a hot-plate (35–60 °C).
358
R e m a r k s : The slides must be grease-free (clean with alcohol and flame); even commercially
359
pre-cleaned slides should be cleaned with a alcohol-moist cloth.
360
5.
Place a small piece (about 2–4 mm in diameter) of gelatin in the centre of the warmed slide and
361
allow to melt.
362
R e m a r k s : Gelatin should have been stored in the refrigerator for at least one week before use.
363
Fresh gelatin may cause cloudy silver deposits.
364
6.
Quickly add an equal-sized or smaller drop of concentrated specimens to the molten gelatin and
365
remove slide from hot-plate.
366
R e m a r k s : Use as few Da Fano’s fluid as possible. Mix organisms thoroughly into the gelatin
367
using a mounted needle.
12
Running title: Observation and preparation of ciliates
368
7.
Quickly spread the drop on the slide or remove excess fluid under the dissecting microscope with
369
a warmed micropipette until ciliates remain just nicely embedded in a thin gelatin layer.
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R e m a r k s : Steps 6 and 7 must be done quickly to avoid hardening and/or desiccation of the
371
gelatin; if gelatin solidifies during the procedure return the slide to the hot-plate for a few
372
seconds. Excess fluid can be removed only if ciliates are large or very numerous. For small (
