If observation of native material is not possible, for example because necessary embedding or staining kills the tissue, one has to fixate the material. Aim of fixation is to preserve the structures of interest in a close to its native form. How fixation is conducted exactly is dependent on the goal one wants to achieve Generally, the effort increases from histology to cytology to ultra-structure research: Not every structure in a cell responds equally well to a given method [4, S. 87]. Immunological methods require intact recognition sites, in situ hybridisation a loose cross-linking so that probes can still reach their complementary sequences and determination of enzymes or other molecules makes their retention in the sample necessary [4, S. 87f]. Fixation should not interfere with staining, or make the tissues brittle and difficult to section [4, S. 87].
Now that an outline of what should be achieved is set, the rest of this section will deal with how to reach these goals. First, mechanisms will be discussed, then general methods, practice, and properties of fixatives. The last part will deal with list protocols and recipes recommended for bryophytes and then general ones.
As already stated, aim of fixation is preservation of cells and cellular structures in a form close to its native state. This can be achieved through dehydration, denaturation and cross-linking.
Dehydrating agents like concentrated solvents (ethanol, propanol, acetone) deprive molecules of their hydration envelope. Without this cage of water molecules proteins and other biomolecules precipitate from the cytoplasm and coagulate [4, S. 89]. “Coagulated” tissues tend to be easy to stain and immunodetection is possible. Compared to native material structures are extensively altered [4, S. 89].
Denaturation through strong acids, heavy metals or heat leads to a loss of natural quaternary or tertiary structures. The resulting random coils form new bonds and cross-links with each other [4, S. 89]. Enzymes usually are no longer active and epitopes for immunodetection unrecognizable [4, S. 89].
Cross-linking agents like aldehydes or osmium(IV)-oxides covalently link proteins and lipids to each other. Since they are consumed in this reaction, sufficient stabilisation hinges on the amount of fixatives used. A high degree of cross-linking leads to a good preservation of structures, but enzyme activity and immunodetection are poor [4, S. 89].
Dehydration and denaturation can be achieved without the use of chemicals by entirely physical means. One can simply dry the tissues, the loss of water leads to denaturation and thus fixation. No components are lost, but non uniform drying and collapsing cells lead to a heavily altered morphology. Freeze-drying or critical-point-drying reduces the occurring damage. Generally, only thin sections can be acceptably dried [4, S. 88]. A related technique is heat fixation. The sample is rapidly heated to above 55°C, which leads to denaturation of proteins and instantaneous death of the cells. To keep structural integrity high, the heat should be applied in a controlled and even manner. Vapor bubbles wreak havoc on the cells [4, S. 88].
The best results with regard to structural integrity are achieved with cryo-fixation. The tissue is quickly cooled to low temperatures, which stops all metabolic reactions. Care must be taken to prevent formation of ice crystals [4, S. 88].
Much more frequently chemical fixation is used. Cells, tissues, whole organisms are immersed in a suitable mixture of fixatives [4, S. 88]. The fixatives start penetrating into the tissue and exert their dehydrating, denaturating and/or cross-linking actions on it. However, not all components of a mixture are equally fast. In the worst case a quick component changes the tissue in a way which prohibits penetration of slower ones. Another problem is extraction of tissue components, swelling or shrinking can produce further artifacts [4, S. 88]. Acidic fixatives destroy mitochondria, but keep chromosomes and spindle apparatus. Basic ones keep mitochondria, but destroy many other structures in the cytoplasm [4, S. 89].
Besides careful selection of sample the process improved by using mixtures of different fixatives, which advantages and disadvantages should compensate each other [4, S. 88f]. Small objects like single leaves or gametophytes or sporophytes can be difficult to treat in this conventional way. For them, methods gluing them to slides or using small pouches of filter paper will be described in more detail under Guidelines and practical methods.
Besides immersion, injection of fixatives and in situ applications by perfusion is possible, but not applicable in bryology or botany.
This section will describe general guidelines to achieve fixations of high quality, the peculiarities of certain fixatives and general practical methods with a focus on bryophytes. Protocols, step-by-step-Instructions, will be offered in the following section.
While basically every research objective needs its own customised fixation procedure, there are some general guidelines to achieve good results. The used material should be in pristine condition. It is possible to use herbarium material as later described, but the results will be worse than when using fresh material [2, § 51]. Bryophyte specimens suffer extensive damage to their ultra structures when exposed to water stress for only seconds.
To conserve as much of the structures as possible, the fixatives should penetrate the tissues very quickly. Romeis recommends the treated sections to be not thicker than 3 mm, or even 1 mm in case of electron microscopy [4, S. 100 und 101], while Schömmer sets an upper limit at 1 cm [2, § 23]. Aqueous solutions tend to penetrate faster than mixtures free of water. Detergents improve penetration [4, S. 99]. Reduced pressure, agitation, microwave ovens, higher temperatures [4, S. 99] and even sonification increase penetration velocity [4, S. 101]. Higher temperatures are a double-edged sword, however. Besides penetration, the speed of autolytic reactions increases as well, resulting in poor conservation of structures. For high resolution microscopy one should fixate at 4°C in a fridge. For histological purposes room temperature yields good enough results [4, S. 99].
According to Schömmer 24 h are enough to completely fixate a specimen [2, § 23]. Romeis recommends three to seven days for botanical material [4, S. 319]. If the preparation cannot be used immediately, it should be stored in storage solutions, not the fixatives. The specimen could become brittle and overly hard [4, S. 99].
The amount of fixative is measured relative to the volume of the specimen. For botanical samples, Romeis recommends 50ty to 100 times the specimen volume for botanical samples [4, S. 318], Schömmer is in agreement [2, § 23]. Generally, a higher specimen volume should be counteracted using a higher volume of fixative, not by a higher concentration [4, S. 99].
When formulating mixtures one has to keep in mind several things. First, the different fixatives should penetrate the specimen equally fast, so that no “crust” forms inside the object preventing penetration of a fixative [4, S. 99]. When using aldehydes, structure retention can be improved by closely mimicking the natural conditions around the target structure. This means osmolarity and pH-value of the fixative should be adjusted [4, S. 99]. Romeis recommends 400 mosm for meristematic and 800 mosm for mature plant cells. Sucrose is used to increase osmolarity, which can be checked using an osmometer or by freezing point depression [4, S. 99]. Buffers should not react with the fixative, not extract cell components, and have the right osmolarity pH-Value and buffering capacity. Usual concentrations are 0,1-0,2 mol/L, usual pH-Value 7,2-7,4 [4, S. 100]. Buffer recipes are listed in the protocols and recipes section.
In this section commonly used fixatives and mixtures are discussed regarding advantages, disadvantages and application.
Ethanol, either pure, 96% or 70%, is the most commonly used fixative in botany, as it is cheap, easy to use and can also be used to store material for some time. It does not interfere with staining [2, § 24, 4, S. 319]. It hardens specimens, sometimes to brittleness. It is recommended to let the material soak in water for several minutes (Romeis) or 24 h (Schömmer)before use [2, § 24, 4, S. 319]. A disadvantage of pure ethanol is the extensive shrinking of cells caused by it [2, § 24, 4, S. 319]. Schömmer and Romeis judge denaturated ethanol differently. Romeis claims nothing special for it, while Schömmer thinks only ethanol denaturated with petrol or phenol is suitable, while ones containing methanol, acetone or pyrimidine bases is unusable [2, § 24, 4, S. 319].
Methanol and Isopropanol are not recommended. Methanol macerates the specimens while isopropanol penetrates too slowly [2, $ 25].
Formaldehyde is available as a 30-40% aqueous solution called formol. Other mayor constituents are methanol (stabilizer, up to 10%), formic acid and formaldehyde polymers [4, S. 91]. It should be stored in tightly closed containers protected from light, since light leads to oxidation to formic acid [4, S. 91]. The formic acid interferes with silver impregnation, then the formaldehyde should be stored above 1 to 2 cm of calcium carbonate [4, S. 91]. When using mixtures formaldehyde should be added just prior for use [4, S. 91].
For use, formol is diluted to about 3-4,5% [2, $ 26]. Structure and stainability are good [4, S. 91]. Storing objects for decades is possible, but a decrease in stainability happens [4, S. 319]. Before use, the botanical material should be washed with a stream of tap water [4, S. 319]. An unpleasant property of formaldehyde are brownish precipitates sometimes formed in the specimen. They can be removed by treatment with 1-5% (w/v) ammonia in 70% ethanol. Frequent check up with a microscope is recommended [4, S. 101].
For electron microscopy used formaldehyde should be freshly prepared from paraformaldehyde. These solutions can be stored for up to 6 months at -20°C [4, S. 98]. Determination of enzymes, immune detection, in situ-hybridisation are possible, but preservation of ultra structures is poor. Only buffered solutions should be used [4, S. 98].
Glutaraldehyde is a dialdehyde which reacts with amino groups of proteins to irreversibly form covalent bridges between proteins [4, S. 96]. Since protons are released, buffered solution should be used. Conservation of ultra structure is very good, specimen can be stored 1,5% glutaraldehyde for several weeks [4, S. 96].
Mixtures of glutaraldehyde and formaldehyde are most commonly used in electron microscopy, also for bryophytes [5, S. 183].
Most gametophytes, sporophytes and protonema can be fixated by a simple immersion fixation. The specimen is immersed in the required volume of tempered fixative, plugs of gauze or cotton can be used to prevent it from settling to the ground or surfacing [4, S. 100f]. If the material is difficult to wet, detergent can be added. Romeis recommends 0,2% saponin [4, S. 319]. Schömmer recommends 24 h ( [2, § 23]), according to Romeis three days and up to a week can be necessary [4, S. 319]. As vessels polyethylene flasks or bottles or small reaction vessels with lids are possible [4, S. 318 and 101].
According to Schömmer, fixation of stems of mosses and liverworts, thalli of liverworts and leaves of Sphagnum and Polytrichum should be delayed after sectioning [2, S. 448].
For electron microscopy, sample selection is very important. Only fresh, actively growing and fully hydrated specimens should be selected. Aging cultures, material stored in polyethylene bags or rehydrated specimens should be avoided [5, S. 182]. These samples show often extensive cell damage in TEM, even when the look healthy in optical microscopy [5, S. 182]. This problem extends to specimens sectioned prior to fixation. Even the few seconds from culture vessel to fixative is enough to damage protonema severely [5, S. 182]. A counter-measurement is to flood bryophytes with water or fixative beforehand [5, S. 182]. Small gametophytes or single leaves need special methods, as does the use of dried herbarium material.
While it is recommended to use fresh material, according to Schömmer herbarium material of mosses and liverworts can be used as a substitute with good results [2, § 51]. The specimen is thoroughly soaked in water. Is retention of air bubbles a problem, vacuum should be applied. If after 24 hours the specimen didn’t reach its natural morphology, it can be gently boiled in water or lactic acid [2, § 51].
If one wants to section the specimen, embedding in celloidin is recommended by Schömmer [2, § 51]. Cuttability is improved by soaking or boiling the specimen or the celloidin block in glycerine, lactic acid or lactophenol. It can be necessary to remove cellular content by treatment with potassium hypochlorite [2, § 51]. As staining iron haematoxylin (Kernschwarz) and 2,4-diamino-azobenzene (Chrysoidin) appears to be working [2, § 51].
When working with bryophytes it can be possible that one is confronted with very small specimens. Using a stereo microscope is an incredible help, but manipulating several specimen at once is still tiresome. Several methods exists to bulk the material together to ease handling.
First, Schmelzers minimum filter pouch. As the name suggest, the material is put inside a small pouch of stable, non-defibring filter paper closed with sewing yarn. It remains inside until it is enclosed on a slide. One advantage of the method is that several samples are treated uniformly. For staining it is advisable to first ascertain staining durations on an extra specimen [2, § 64].
It is possible to glue small sections to slides using gummi arabicum (Straßburgers method) or a gelatine preparation (Adams method). Similar to this is Schmelzers celloidin method. Here, specimens are “embedded” in a small skin of celloidin, which can remain attached to a slide or peeled of rom it. Details are given in der Protocols and Recepies section [2, § 64].
[1] Frahm, J.-P. u. Müller, R.-D.: Moose unter dem Mikroskop. Archive of Bryology Special Volume (2013) 13
[2] Schömmer, F.: Kryptogamen-Praktikum. Anleitung zur mikroskopischen Untersuchung und Präparation der blütenlosen Pflanzen für Studierende und Liebhaber. Stuttgart: Kosmos Gesellschaft der Naturfreunde Franckh'sche Verlagshandlung Stuttgart 1949
[3] Smith, A. J. E.: The moss flora of Britain and Ireland. Cambridge, UK, New York: Cambridge University Press 2004
[4] Mulisch, M., Welsch, U. u. Aescht, E. (Hrsg.): Romeis mikroskopische Technik. Heidelberg: Spektrum Akademischer Verlag 2010
[5] Janice M. Glime (Hrsg.): Methods in Bryology. Nichinan, Miyazaki, Japan: The Hattori Botanical Laboratory 1988