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The use of biological stains in the analysis of late Palaeozoic pteridosperm cuticles
Review of Palaeobotany and Palynology 108 (2000) 143–150 www.elsevier.nl/locate/revpalbo
Technical note
The use of biological stains in the analysis of late Palaeozoic pteridosperm cuticles Michael Krings * Abt. Pala¨obotanik am Geologisch-Pala¨ontologischen Institut; Westfa¨lische Wilhelms-Universita¨t Mu¨nster; Hindenburgplatz 57–59; D-48143 Mu¨nster, Germany Received 18 May 1999; accepted for publication 26 July 1999
Abstract The use of biological stains in the cuticular analysis of late Palaeozoic pteridosperms based on specimens from the Stephanian Blanzy-Montceau Basin (Central France) is discussed. Bismarck Brown, Malachite Green G, Methylene Blue, Methyl Green, Neutral Red, Safranin T, and a double staining with Neutral Red and Malachite Green G, were tested. Bismarck Brown, Malachite Green G, Methylene Blue, and Neutral Red increase contrast and emphasize differences in cutinization. The double staining in Neutral Red and Malachite Green G enhances the three-dimensional morphology of complex epidermal structures. Safranin T increases contrast, emphasizes cutinization differences, and enhances the three-dimensional morphology of complex epidermal features. The colour photography of cuticles is normally not affected by the presence of stains, but some stains mask black-and-white half-tones. © 2000 Elsevier Science B.V. All rights reserved. Keywords: cuticles; epidermal anatomy; pteridosperms; staining methods
1. Introduction Cuticular analysis is a valuable method for the study of fossil pteridosperms and increasingly contributes to our knowledge of the taxonomy and autecology of many taxa (e.g. Barthel, 1962; Kerp and Barthel, 1993; Cleal and Shute, 1995; Krings and Kerp, 1997a, 1998, 1999). However, the importance of cuticles in studies of compression taxa relies largely on the ability to analyse the morphology and distribution of the epidermal features, particularly stomatal complexes and epidermal appendages (e.g. scales, trichomes) since * Fax: +49-251-8321739. E-mail address: [email protected] (M. Krings)
they play an important role in taxonomy and autecology (Barthel, 1962; Kerp, 1990; Krings, 1997). Cuticle features are especially important in comparative analyses since even minor morphological differences may be highly significant. Fossil plant cuticles are usually studied in transmitted light. Epidermal anatomy generally can be studied and photographed from macerated cuticles without any special treatment. However, problems sometimes occur when the cutinization of the epidermal anticlinal walls is especially weak, or is distorted during fossilization and/or the preparation procedure, resulting in an indistinct epidermal cell pattern ( Krings and Kerp, 1998). The analysis of the morphology of stomatal complexes and epidermal appendages and their relationship
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to the epidermis can be more problematic because the complexity of these structures (e.g. certain features of stomatal complexes) may obscure details (e.g. Krings, 1999). Moreover, a thorough analysis can be difficult because stomatal complexes and epidermal appendages may be naturally weakly cutinized and/or may not differ in colour from the surrounding cuticle. The use of stains to increase contrast and emphasize the three-dimensional morphology of complex plant structures has been important in modern plant anatomy as well as in Mesozoic and Cenozoic palaeobotany. The most commonly used stains in palaeobotany are Safranin and Methylene Blue. However, to date, the use of stains in the analysis of Palaeozoic plant cuticles is rare. Palaeozoic cuticles are often fragile and do not survive a treatment with stains when basic techniques are followed. This is mainly because in many of the original instructions, the cuticles are aggressively dehydrated (e.g. in methanol, acetone, xylene), usually causing the cuticle to become brittle and to separate into minute pieces. However, avoiding rapid dehydration often yields important results even with Palaeozoic cuticles. Methods used in the staining of cuticles of Stephanian ( Upper Carboniferous) pteridosperms from the Blanzy-Montceau Basin (Central France) are described, and some comments are offered on specific stain schedules.
2. Material and methods Materials were collected from several open-cast mines in the basin of Blanzy-Montceau, which is one of several intramontane basins of the French Massif Central. The coal-bearing strata there have been dated as late Stephanian (Langiaux, 1984). Specimens were collected by the author; others were borrowed from collections in Montceau-lesMines (collection Langiaux) and Autun (collection A. and D. Sotty). Cuticles were prepared according to instructions provided in Kerp (1990) and Kerp and Krings (1999). Pieces of rock containing plant remains were bulk-macerated in 45% hydrofluoric acid for several days. Plant remains were then picked out from the acid, gently washed in several
rinses of distilled water and subsequently macerated according to a modified standard procedure ( Krings and Kerp, 1997b) using Schulze’s reagent ( HNO with a few crystals of KClO ). The macer3 3 ated cuticles were washed in distilled water and then stained (see below). Stained cuticles were mounted in permanent glycerine-jelly microscope slides. Cuticles were photographed according to the methods described in Kerp and Krings (1999). Colour slides were made on a daylight film (e.g. Fujichrome Sensia 100 ISO) using a CB 12 blue filter, and black-and-white pictures were made on either a low-speed panchromatic (e.g. Agfapan 25 ISO) or orthochromatic (e.g. Agfaortho 25 ISO) film. The most productive method of staining fragile pteridosperm cuticles requires an absence of aggressively dehydrating chemicals as solvents for the stains, as washing liquids for the stained cuticles, and as fixers. As a suitable substitute, distilled water — as washing liquid, also (warm) glycerine — was used. None of the stained cuticles was fixed. The stains tested include Bismarck brown (Chroma, Mu¨nster, Germany), Malachite Green G (Chroma), Methylene Blue (Chroma), Methyl Green (Chroma), Neutral Red (Chroma), and Safranin T (Fluka, Deisenhofen, Germany). All stains, which are commercially sold as powders or crystals by the companies given in brackets, were used as 1% aqueous solutions. For the best results, staining solutions should always be stirred and slightly warmed to approximately 30°C immediately before use since the stains have a tendency to fall out of suspension. All cuticles were regressively stained, i.e. they were initially overstained and excessive stain was later removed. On a microscope slide, pieces of cuticle were treated in a drop of the staining solution for a few minutes; the staining reaction was observed through a stereo-microscope. If no reaction was observed after a few minutes, the slide was gently warmed on a precision heating plate to 35–40°C. If still no reaction took place, the staining result was considered negative. If a reaction took place, the cuticles were quickly removed from the staining solution in order to avoid irreversible overstaining. Using a preparation needle, cuticles were transferred into a watch-
M. Krings / Review of Palaeobotany and Palynology 108 (2000) 143–150
glass with warm distilled water (30–35°C ). However, particularly delicate cuticles can also be transferred into water by placing the entire object slide in a petri dish, which is then very carefully filled with warm water. If only a little of the excessive stain could be removed from the cuticles in water, they were also gently washed in warm glycerine. However, because of the use of distilled water and glycerine, respectively, as washing liquids, it is almost impossible to remove the excessive stain completely from the cuticles. After a gentle dehydration of the stained cuticles in pure glycerine ( Krings and Kerp, 1997b), they were mounted in permanent glycerine-jelly microscope slides. Because of this procedure, it is often impossible to avoid stains leaching from the cuticles during mounting in glycerine jelly. As a result of the lack of fixation, many stained cuticles will fade in intensity and/or become faint within approximately 2–4 months. Thus, it is important to photograph and analyse these cuticle specimens immediately!
3. Observations and comments 3.1. General observations Although all of the stains tested positive, two aspects were different. On the one hand, cuticles of a single species can react differently to various stains. On the other hand, cuticles of different species may react differently to the same stain procedure. For example, pinnule cuticles of Barthelopteris germarii, Blanzyopteris praedentata, and Lescuroptertis genuina take up all the stains used, while Pseudomariopteris paleaui cuticles failed to stain with any of the reagents. Methyl Green stained the costal fields of Barthelopteris germarii pinnules, while in Pseudomariopteris busquetii, the entire cuticle became stained. Safranin T stained the costal fields of Lescuropteris genuina pinnules and the stomata and trichome bases in the intercostal fields red but did not stain the entire intercostal fields. Pinnules of Blanzyopteris praedentata and Neuropteris cordata reacted uniformly to Safranin T. Methylene Blue stained the costal fields of pinnules of
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Barthelopteris germarii, while it entirely stained pinnule cuticles of Lescuropteris genuina, Neurodontopteris auriculata, and Pseudomariopteris busquetii. It is not entirely clear why cuticles of different pteridosperm species react differently to the same stain procedure. No general relationship between the mode of preservation of the BlanzyMontceau material (e.g. black compression, brown compression) and the reaction of the cuticles to stains is recognizable. However, the reactions to stains probably reflect differences in the composition of the cuticles themselves. In particular, thicker cuticles (e.g. upper pinnule surface cuticles of Pseudomariopteris ribeyronii, pinna axis cuticles of Neuropteris cordata) could only be stained when their inner sides were directly exposed to the stain. A plant cuticle is not a uniform layer but is heterogenous, structurally consisting of several layers each of which is characterized by its chemical composition (Holloway, 1982). These layers include (1) the cuticle proper, which is a continuous adcrustation onto the external surface of the epidermal cell walls, and (2) cutinized layers beneath the cuticle proper, which are incrustations of cuticular material into the outer periclinal and anticlinal cell walls (Esau, 1965). The behaviour of thicker pteridosperm cuticles to stains indicates that the presence of cutinized layers may be crucial for a positive staining result, and a pure cuticle proper fails to stain. However, it remains uncertain whether the presence of either the complete cutinized layers, some of their components (e.g. cell wall remains), or only spaces between the cuticular material — remaining after disintegration of cell wall material — are important. Moreover, it is unclear whether the presence/absence of the cutinized layers and their components, respectively, is of biological origin, a result of the preparation procedures, or a result of a combination of fossilization and diagenetic processes. The latter is probable because features indicating considerable modifications of the cuticle structure during fossilization and diagenesis, e.g. traces of oxidation and pyrite-holes, are often present. However, because, in modern plants, the organization of the cuticle is highly variable from species to species ( Holloway, 1982), and even within a single species, different ecological conditions during cuticle for-
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PLATE I
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mation and hardening can result in structural variations (Skoss, 1955; Napp-Zinn, 1966; Martin and Juniper, 1970), at least some of the reactions to stains are likely to be caused by natural differences in the structure of the cuticles. A major problem in staining fossil cuticles is the ability to photograph the stained cuticle features. Colour photography using a daylight film and a CB 12 blue filter ( Kerp and Krings, 1999) is usually not influenced by the presence of stains. However, documenting features using a black-andwhite film is, in some cases, difficult or even impossible because of the stains. Stains can produce light irritations that mask features and result in obscured details (e.g. Plate II, 2). These difficulties may be the result of both the intensive staining results and the incomplete removal of the staining solution from the cuticles. The use of lower stain concentrations (0.5 or 0.25%) did not reduce the stain intensity or enhance features when photographed in black-and-white film. When even lower stain concentrations ( less than 0.1%) were used, the staining reaction was normally negative. 3.2. Comments on the individual stains 3.2.1. Bismarck Brown This stain is best used after gently warming to 35–40°C. It intensifies the natural colours of the cuticles and then emphasizes the existing contrasts (Plate I, 1); however, three-dimensional epidermal features cannot be significantly enhanced.
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Bismarck Brown has the advantage that excessive amounts can be easily washed from the cuticles with warm water. In addition, slides stained with Bismarck Brown are permanent because the stain will neither fade nor become diffuse over time. It is particularly interesting that Bismarck Brown does not negatively influence features when photographed in black-and-white film, even when delicate details are examined (e.g. Krings and Kerp, 1998). Finally, colour slides of the stained cuticles are not altered by unnatural colours. 3.2.2. Malachite Green G This stain emphasizes differences in the cutinization of epidermal structures (e.g. compare Plate II, 1 with Plate II, 3), similar to Methylene Blue and Neutral Red. Excessive stain can be washed out in warm water or glycerine. The stain is not very long-lasting and will become more and more pale and diffuse with time. Neither colour photography nor black-and-white documentation is obstructed if the features are not too intensively stained. However, since Malachite Green G is highly toxic, it has to be used with caution, especially when warming is required, and hence Methylene Blue and Neutral Red, respectively, should be preferred. 3.2.3. Methylene Blue and Neutral Red These two stains are very useful when it is necessary to discern — especially smaller — differences in the cutinization of epidermal structures. Distinctive intensities of the blue(–green) and red
PLATE I Unstained and stained Stephanian pteridosperm cuticle features. 1–3. Cuticle features of Barthelopteris germarii. 1. Cuticle from a lower pinnule surface stained with Bismarck Brown; note the numerous peltate glandular trichomes (arrows); slide Rg.106; scale bar=400 mm. 2. Unstained peltate glandular trichome; slide Rg.17; scale bar=20 mm. 3. Peltate glandular trichome double stained with Neutral Red and Malachite Green G; slide Rg.MaGNr2; scale bar=20 mm. 4. Stoma from the lower side of a pinnule of Neuropteris cordata stained with Methylene Blue. Different blue to blue–green hues in the guard cells indicate variations in the thickness and cutinization of the dorsal walls; the unthickened polar areas are clearly discernible (arrows); slide Nc.Mb2; scale bar=6.25 mm. 5. Stoma from the lower side of a pinnule of Blanzyopteris praedentata, stained with Safranin T; slide Np.36Sa; scale bar=6.5 mm. 6–8. Cuticle features of Lescuropteris genuina. 6. Unstained pinnule and portion of the pinna axis; slide Lg.533; scale bar=1.5 mm. 7. Pinnule portion double stained with Neutral Red and Malachite Green G; slide Lg.753; scale bar=250 mm. 8. Detail of a pinna axis stained with Methyl Green; slide Lg.758; scale bar=200 mm.
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PLATE II
Black-and-white half-tones of unstained and stained stomata of Neuropteris cordata. 1. Unstained stoma; slide Nc.4.09; scale bar=6.25 mm. 2. Stoma double stained with Neutral Red and Malachite Green G; slide Nc.MaGNr.1; scale bar=8.5 mm. 3. Stoma stained with Malachite Green G; the arrow indicates the cutinization around the polar contact of the two guard cells; slide Nc.30MaG; scale bar=6.25 mm. 4–6. Stomata stained with Safranin T; slide Nc.30Sa; scale bars=22.5 mm (4), 6.5 mm (5), and 6.25 mm (6).
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colour, respectively, indicating different cutinizations, are easily discernible (e.g. Plate I, 4), and this is of particular interest in the analysis of very delicate and transparent cuticles or epidermal features. For example, cutinization details of the complex stomatal apparatus of Neurodontopteris auriculata could only be studied and sufficiently documented after staining the transparent cuticles with Methylene Blue ( Krings, 1999). Excessive stain can be washed out with warm water. Cuticle slides are long-lasting and do not fade. Black-andwhite photography is best accomplished on panchromatic film. 3.2.4. Methyl Green This stain does not appear to be useful for late Palaeozoic pteridosperm cuticles. The stain is intensive and does not normally improve the characterization of epidermal features, and black-andwhite photography is almost impossible. However, a particularly interesting result was seen when cuticles of Lescuropteris genuina were stained with Methyl Green. The maceration of plant remains in Schulze’s solution normally destroys internal structures, which may be preserved in the coaly layer between the cuticles of pinnules and axes. Some of these structures can be of interest for palaeoecological evaluations, e.g. the number of vascular strands in axes and/or the width of vascular strands in comparison to the width of overlying costal fields etc. Generally, in macerated cuticles of L. genuina, the patterns of the vascular strands and their width cannot be followed ( Krings and Kerp, 1997a and Plate I, 6). By staining the cuticles in Methyl Green, these features could be easily discerned. Above and below the original courses of the vascular strands, an intensive staining of the cuticle indicates the exact position of the vascular strands (Plate I, 8). However, this staining result fades within a few weeks. 3.2.5. Safranin T This is perhaps the best multipurpose stain to use with late Palaeozoic pteridosperm cuticles. Apart from its excellent qualities in emphasizing existing contrasts, it can be used in the analysis of complex three-dimensional structures, particularly
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stomatal complexes, because it enhances cell borders and varying amounts of cutinization, even the finest details (compare Plate II, 1 with Plate II, 4– 6). For example, the tiny stomatal complexes of Lescuropteris genuina were first recorded after staining with Safranin T ( Krings and Kerp, 1997a). Excessive stain can be washed out in warm water. The stain is fairly long-lasting; however, it will become increasingly pale and diffuse with time. Photography on panchromatic black-andwhite film is not obstructed if the features are not too intensively stained; the surface views of the stomatal complexes of Blanzyopteris praedentata ( Krings and Kerp, 1999) and Neuropteris cordata (Plate II, 4–6) were best documented after staining with Safranin T. Colour representation on daylight film is also acceptable, but not as effective as black-and-white photography (e.g. Plate I, 5).
3.2.6. Double staining in Neutral Red and Malachite Green G In this stain schedule, cuticles are first treated with Neutral Red for 1–4 min, then briefly rinsed in warm water, and after that immediately treated with Malachite Green for approximately 30 s. Excessive stain can be removed with warm glycerine. This procedure produces excellent results in the analysis of complex epidermal structures. The distribution patterns of blended hues of red, green, blue, and violet provide an excellent three-dimensional impression of a structure (e.g. Plate I, 3). For example, this stain resulted in the ability to resolve the complex three-dimensional structure of the peltate glandular trichomes of Barthelopteris germarii and their relationship to the epidermis (compare Plate I, 2 with Plate I, 3). Moreover, in some species (e.g. Lescuropteris genuina, Odontopteris minor f. zeilleri), the stain highlights different pinnule areas, i.e. pinnule margin, costal and intercostal fields (compare Plate I, 6 with Plate I, 7). Unfortunately, cuticles stained in this manner usually fade within a couple of months. The colour photography of stained cuticles often yields spectacular results, while significant light irregularities make black-and-white photography almost impossible (e.g. Plate II, 2).
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4. Concluding remarks
References
Experiments carried out on pteridosperms from the Blanzy-Montceau Basin (Central France) indicate that the use of stains can contribute considerably to the enhancement of features used in studies of the epidermal anatomy of late Palaeozoic pteridosperms. The use of distilled water and/or glycerine during the staining process makes it possible to stain fragile cuticles in the absence of rapidly dehydrating substances. However, using this procedure, cuticles must be photographed and studied immediately since the cuticle stain often leaches out. Moreover, black-and-white photography of cuticles can be obstructed by the presence of stain. However, with further advancements, e.g. digital photography adjustment, the latter problem could possibly be reduced. The results obtained by the staining of pteridosperm cuticles from BlanzyMontceau indicate that no single stain can generally be recommended for late Palaeozoic pteridosperm cuticles; the effect of staining may be different for various species, even from the same locality. Different stains must be tested on the cuticles under examination. However, Bismarck Brown and Safranin T appear to be the most universal stains for fossil plant materials.
Barthel, M., 1962. Epidermisuntersuchungen an einigen inkohlten Pteridospermenbla¨ttern des Oberkarbons und Perms. Geologie 11, Beiheft 33, 1–140. Cleal, C.J., Shute, C.H., 1995. A synopsis of neuropterid foliage from the Carboniferous and Lower Permian of Europe. Bull. Nat. Hist. Mus. Lond. (Geology) 51, 1–52. Esau, K., 1965. Plant Anatomy. Wiley, New York. 767 pp. Holloway, P.J., 1982. Structure and histochemistry of plant cuticular membranes: an overview. In: Cutler, D.F., Alvin, K.L., Price, C.E. ( Eds.), The Plant Cuticle. Academic Press, London, pp. 1–32. Kerp, H., 1990. The study of fossil gymnosperms by means of cuticular analysis. Palaios 5, 548–569. Kerp, H., Barthel, M., 1993. Problems of cuticular analysis of pteridosperms. Rev. Palaeobot. Palynol. 78, 1–18. Kerp, H., Krings, M., 1999. Light microscopy of fossil cuticles. In: Jones, T.P., Rowe, N.P. (Eds.), Fossil Plants and Spores: Modern Techniques. Special Publications Geological Society, London, pp. 52–56. Krings, M., 1997. Mo¨glichkeiten und Grenzen der Kutikularanalyse — das Beispiel der Samenfarne aus dem Stefan (Oberkarbon) von Blanzy-Montceau (Zentralfrankreich). Vero¨ffentlichungen des Museums fu¨r Naturkunde Chemnitz 20, 57–70. Krings, M., Kerp, H., 1997a. Cuticles of Lescuropteris genuina from the Stephanian ( Upper Carboniferous) of Central France — evidence for a climbing growth habit. Bot. J. Linn. Soc. 123, 73–89. Krings, M., Kerp, H., 1997b. An improved method for obtaining large pteridosperm cuticles. Rev. Palaeobot. Palynol. 96, 453–456. Krings, M., Kerp, H., 1998. Epidermal anatomy of Barthelopteris germarii from the Upper Carboniferous and Lower Permian of France and Germany. Am. J. Bot. 85, 553–562. Krings, M., 1999. Zum Bau der Spalto¨ffnungsapparate von Neurodontopteris auriculata (Brongniart) Potonie´, einer Pteridosperme aus dem Stefan von Blanzy-Montceau (Zentralfrankreich). Mu¨nstersche Forschungen zur Geologie und Pala¨ontologie 86, 69–78. Krings, M., Kerp, H., 1999. Morphology, growth habit, and ecology of Blanzyopteris praedentata (Gothan) nov.comb., a climbing neuropteroid seed fern from the Stephanian of Central France. Int. J. Plant Sci. 160, 603–619. Langiaux, J., 1984. Flores et faunes des formations supe´rieures du Ste´phanien de Blanzy-Montceau (Massif Central franc¸ais) — stratigraphie et pale´oe´cologie. Revue pe´riodique de ‘la Physiophile’ Socie´te´ d’e´tudes des sciences historiques de Montceau-les-Mines. Suppl. to Vol. 100. Martin, J.T., Juniper, B.E., 1970. The Cuticles of Plants. Arnold, London. 337 pp. Napp-Zinn, K., 1966. Anatomie des Blattes. I. Blattanatomie der Gymnospermen. In: Zimmermann, W., Ozenda, P., Wulff, H.D. (Eds.), Handbuch der Pflanzenanatomie. Gebr. Borntraeger, Berlin, Spezieller Teil, Band VIII, Teil 1, 369 pp. Skoss, J.D., 1955. Structure and composition of the plant cuticle in relation to environmental factors and permeability. Bot. Gazette 117, 55–72.
Acknowledgements This study was carried out at the University of Mu¨nster (Germany) in conjunction with a research project supported by the German Science Foundation (DFG grants Ke 584/2-1 and Ke 584/2-2), and was completed at the University of Kansas (Lawrence, KS, USA) with the support of the Alexander von Humboldt Foundation in the form of a Feodor Lynen Research Fellowship given to the author. The author is grateful to Dr J. Langiaux (Gourdon, France) and Dr G. Pacaud (formerly Autun, France) for making available most of the specimens studied. The author wants to thank Dr H. Kerp (Mu¨nster, Germany) for helpful discussion and Dr T.N. Taylor (Lawrence, KS, U.S.A.) for critical reading of the manuscript.
Technical note
The use of biological stains in the analysis of late Palaeozoic pteridosperm cuticles Michael Krings * Abt. Pala¨obotanik am Geologisch-Pala¨ontologischen Institut; Westfa¨lische Wilhelms-Universita¨t Mu¨nster; Hindenburgplatz 57–59; D-48143 Mu¨nster, Germany Received 18 May 1999; accepted for publication 26 July 1999
Abstract The use of biological stains in the cuticular analysis of late Palaeozoic pteridosperms based on specimens from the Stephanian Blanzy-Montceau Basin (Central France) is discussed. Bismarck Brown, Malachite Green G, Methylene Blue, Methyl Green, Neutral Red, Safranin T, and a double staining with Neutral Red and Malachite Green G, were tested. Bismarck Brown, Malachite Green G, Methylene Blue, and Neutral Red increase contrast and emphasize differences in cutinization. The double staining in Neutral Red and Malachite Green G enhances the three-dimensional morphology of complex epidermal structures. Safranin T increases contrast, emphasizes cutinization differences, and enhances the three-dimensional morphology of complex epidermal features. The colour photography of cuticles is normally not affected by the presence of stains, but some stains mask black-and-white half-tones. © 2000 Elsevier Science B.V. All rights reserved. Keywords: cuticles; epidermal anatomy; pteridosperms; staining methods
1. Introduction Cuticular analysis is a valuable method for the study of fossil pteridosperms and increasingly contributes to our knowledge of the taxonomy and autecology of many taxa (e.g. Barthel, 1962; Kerp and Barthel, 1993; Cleal and Shute, 1995; Krings and Kerp, 1997a, 1998, 1999). However, the importance of cuticles in studies of compression taxa relies largely on the ability to analyse the morphology and distribution of the epidermal features, particularly stomatal complexes and epidermal appendages (e.g. scales, trichomes) since * Fax: +49-251-8321739. E-mail address: [email protected] (M. Krings)
they play an important role in taxonomy and autecology (Barthel, 1962; Kerp, 1990; Krings, 1997). Cuticle features are especially important in comparative analyses since even minor morphological differences may be highly significant. Fossil plant cuticles are usually studied in transmitted light. Epidermal anatomy generally can be studied and photographed from macerated cuticles without any special treatment. However, problems sometimes occur when the cutinization of the epidermal anticlinal walls is especially weak, or is distorted during fossilization and/or the preparation procedure, resulting in an indistinct epidermal cell pattern ( Krings and Kerp, 1998). The analysis of the morphology of stomatal complexes and epidermal appendages and their relationship
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to the epidermis can be more problematic because the complexity of these structures (e.g. certain features of stomatal complexes) may obscure details (e.g. Krings, 1999). Moreover, a thorough analysis can be difficult because stomatal complexes and epidermal appendages may be naturally weakly cutinized and/or may not differ in colour from the surrounding cuticle. The use of stains to increase contrast and emphasize the three-dimensional morphology of complex plant structures has been important in modern plant anatomy as well as in Mesozoic and Cenozoic palaeobotany. The most commonly used stains in palaeobotany are Safranin and Methylene Blue. However, to date, the use of stains in the analysis of Palaeozoic plant cuticles is rare. Palaeozoic cuticles are often fragile and do not survive a treatment with stains when basic techniques are followed. This is mainly because in many of the original instructions, the cuticles are aggressively dehydrated (e.g. in methanol, acetone, xylene), usually causing the cuticle to become brittle and to separate into minute pieces. However, avoiding rapid dehydration often yields important results even with Palaeozoic cuticles. Methods used in the staining of cuticles of Stephanian ( Upper Carboniferous) pteridosperms from the Blanzy-Montceau Basin (Central France) are described, and some comments are offered on specific stain schedules.
2. Material and methods Materials were collected from several open-cast mines in the basin of Blanzy-Montceau, which is one of several intramontane basins of the French Massif Central. The coal-bearing strata there have been dated as late Stephanian (Langiaux, 1984). Specimens were collected by the author; others were borrowed from collections in Montceau-lesMines (collection Langiaux) and Autun (collection A. and D. Sotty). Cuticles were prepared according to instructions provided in Kerp (1990) and Kerp and Krings (1999). Pieces of rock containing plant remains were bulk-macerated in 45% hydrofluoric acid for several days. Plant remains were then picked out from the acid, gently washed in several
rinses of distilled water and subsequently macerated according to a modified standard procedure ( Krings and Kerp, 1997b) using Schulze’s reagent ( HNO with a few crystals of KClO ). The macer3 3 ated cuticles were washed in distilled water and then stained (see below). Stained cuticles were mounted in permanent glycerine-jelly microscope slides. Cuticles were photographed according to the methods described in Kerp and Krings (1999). Colour slides were made on a daylight film (e.g. Fujichrome Sensia 100 ISO) using a CB 12 blue filter, and black-and-white pictures were made on either a low-speed panchromatic (e.g. Agfapan 25 ISO) or orthochromatic (e.g. Agfaortho 25 ISO) film. The most productive method of staining fragile pteridosperm cuticles requires an absence of aggressively dehydrating chemicals as solvents for the stains, as washing liquids for the stained cuticles, and as fixers. As a suitable substitute, distilled water — as washing liquid, also (warm) glycerine — was used. None of the stained cuticles was fixed. The stains tested include Bismarck brown (Chroma, Mu¨nster, Germany), Malachite Green G (Chroma), Methylene Blue (Chroma), Methyl Green (Chroma), Neutral Red (Chroma), and Safranin T (Fluka, Deisenhofen, Germany). All stains, which are commercially sold as powders or crystals by the companies given in brackets, were used as 1% aqueous solutions. For the best results, staining solutions should always be stirred and slightly warmed to approximately 30°C immediately before use since the stains have a tendency to fall out of suspension. All cuticles were regressively stained, i.e. they were initially overstained and excessive stain was later removed. On a microscope slide, pieces of cuticle were treated in a drop of the staining solution for a few minutes; the staining reaction was observed through a stereo-microscope. If no reaction was observed after a few minutes, the slide was gently warmed on a precision heating plate to 35–40°C. If still no reaction took place, the staining result was considered negative. If a reaction took place, the cuticles were quickly removed from the staining solution in order to avoid irreversible overstaining. Using a preparation needle, cuticles were transferred into a watch-
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glass with warm distilled water (30–35°C ). However, particularly delicate cuticles can also be transferred into water by placing the entire object slide in a petri dish, which is then very carefully filled with warm water. If only a little of the excessive stain could be removed from the cuticles in water, they were also gently washed in warm glycerine. However, because of the use of distilled water and glycerine, respectively, as washing liquids, it is almost impossible to remove the excessive stain completely from the cuticles. After a gentle dehydration of the stained cuticles in pure glycerine ( Krings and Kerp, 1997b), they were mounted in permanent glycerine-jelly microscope slides. Because of this procedure, it is often impossible to avoid stains leaching from the cuticles during mounting in glycerine jelly. As a result of the lack of fixation, many stained cuticles will fade in intensity and/or become faint within approximately 2–4 months. Thus, it is important to photograph and analyse these cuticle specimens immediately!
3. Observations and comments 3.1. General observations Although all of the stains tested positive, two aspects were different. On the one hand, cuticles of a single species can react differently to various stains. On the other hand, cuticles of different species may react differently to the same stain procedure. For example, pinnule cuticles of Barthelopteris germarii, Blanzyopteris praedentata, and Lescuroptertis genuina take up all the stains used, while Pseudomariopteris paleaui cuticles failed to stain with any of the reagents. Methyl Green stained the costal fields of Barthelopteris germarii pinnules, while in Pseudomariopteris busquetii, the entire cuticle became stained. Safranin T stained the costal fields of Lescuropteris genuina pinnules and the stomata and trichome bases in the intercostal fields red but did not stain the entire intercostal fields. Pinnules of Blanzyopteris praedentata and Neuropteris cordata reacted uniformly to Safranin T. Methylene Blue stained the costal fields of pinnules of
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Barthelopteris germarii, while it entirely stained pinnule cuticles of Lescuropteris genuina, Neurodontopteris auriculata, and Pseudomariopteris busquetii. It is not entirely clear why cuticles of different pteridosperm species react differently to the same stain procedure. No general relationship between the mode of preservation of the BlanzyMontceau material (e.g. black compression, brown compression) and the reaction of the cuticles to stains is recognizable. However, the reactions to stains probably reflect differences in the composition of the cuticles themselves. In particular, thicker cuticles (e.g. upper pinnule surface cuticles of Pseudomariopteris ribeyronii, pinna axis cuticles of Neuropteris cordata) could only be stained when their inner sides were directly exposed to the stain. A plant cuticle is not a uniform layer but is heterogenous, structurally consisting of several layers each of which is characterized by its chemical composition (Holloway, 1982). These layers include (1) the cuticle proper, which is a continuous adcrustation onto the external surface of the epidermal cell walls, and (2) cutinized layers beneath the cuticle proper, which are incrustations of cuticular material into the outer periclinal and anticlinal cell walls (Esau, 1965). The behaviour of thicker pteridosperm cuticles to stains indicates that the presence of cutinized layers may be crucial for a positive staining result, and a pure cuticle proper fails to stain. However, it remains uncertain whether the presence of either the complete cutinized layers, some of their components (e.g. cell wall remains), or only spaces between the cuticular material — remaining after disintegration of cell wall material — are important. Moreover, it is unclear whether the presence/absence of the cutinized layers and their components, respectively, is of biological origin, a result of the preparation procedures, or a result of a combination of fossilization and diagenetic processes. The latter is probable because features indicating considerable modifications of the cuticle structure during fossilization and diagenesis, e.g. traces of oxidation and pyrite-holes, are often present. However, because, in modern plants, the organization of the cuticle is highly variable from species to species ( Holloway, 1982), and even within a single species, different ecological conditions during cuticle for-
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PLATE I
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mation and hardening can result in structural variations (Skoss, 1955; Napp-Zinn, 1966; Martin and Juniper, 1970), at least some of the reactions to stains are likely to be caused by natural differences in the structure of the cuticles. A major problem in staining fossil cuticles is the ability to photograph the stained cuticle features. Colour photography using a daylight film and a CB 12 blue filter ( Kerp and Krings, 1999) is usually not influenced by the presence of stains. However, documenting features using a black-andwhite film is, in some cases, difficult or even impossible because of the stains. Stains can produce light irritations that mask features and result in obscured details (e.g. Plate II, 2). These difficulties may be the result of both the intensive staining results and the incomplete removal of the staining solution from the cuticles. The use of lower stain concentrations (0.5 or 0.25%) did not reduce the stain intensity or enhance features when photographed in black-and-white film. When even lower stain concentrations ( less than 0.1%) were used, the staining reaction was normally negative. 3.2. Comments on the individual stains 3.2.1. Bismarck Brown This stain is best used after gently warming to 35–40°C. It intensifies the natural colours of the cuticles and then emphasizes the existing contrasts (Plate I, 1); however, three-dimensional epidermal features cannot be significantly enhanced.
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Bismarck Brown has the advantage that excessive amounts can be easily washed from the cuticles with warm water. In addition, slides stained with Bismarck Brown are permanent because the stain will neither fade nor become diffuse over time. It is particularly interesting that Bismarck Brown does not negatively influence features when photographed in black-and-white film, even when delicate details are examined (e.g. Krings and Kerp, 1998). Finally, colour slides of the stained cuticles are not altered by unnatural colours. 3.2.2. Malachite Green G This stain emphasizes differences in the cutinization of epidermal structures (e.g. compare Plate II, 1 with Plate II, 3), similar to Methylene Blue and Neutral Red. Excessive stain can be washed out in warm water or glycerine. The stain is not very long-lasting and will become more and more pale and diffuse with time. Neither colour photography nor black-and-white documentation is obstructed if the features are not too intensively stained. However, since Malachite Green G is highly toxic, it has to be used with caution, especially when warming is required, and hence Methylene Blue and Neutral Red, respectively, should be preferred. 3.2.3. Methylene Blue and Neutral Red These two stains are very useful when it is necessary to discern — especially smaller — differences in the cutinization of epidermal structures. Distinctive intensities of the blue(–green) and red
PLATE I Unstained and stained Stephanian pteridosperm cuticle features. 1–3. Cuticle features of Barthelopteris germarii. 1. Cuticle from a lower pinnule surface stained with Bismarck Brown; note the numerous peltate glandular trichomes (arrows); slide Rg.106; scale bar=400 mm. 2. Unstained peltate glandular trichome; slide Rg.17; scale bar=20 mm. 3. Peltate glandular trichome double stained with Neutral Red and Malachite Green G; slide Rg.MaGNr2; scale bar=20 mm. 4. Stoma from the lower side of a pinnule of Neuropteris cordata stained with Methylene Blue. Different blue to blue–green hues in the guard cells indicate variations in the thickness and cutinization of the dorsal walls; the unthickened polar areas are clearly discernible (arrows); slide Nc.Mb2; scale bar=6.25 mm. 5. Stoma from the lower side of a pinnule of Blanzyopteris praedentata, stained with Safranin T; slide Np.36Sa; scale bar=6.5 mm. 6–8. Cuticle features of Lescuropteris genuina. 6. Unstained pinnule and portion of the pinna axis; slide Lg.533; scale bar=1.5 mm. 7. Pinnule portion double stained with Neutral Red and Malachite Green G; slide Lg.753; scale bar=250 mm. 8. Detail of a pinna axis stained with Methyl Green; slide Lg.758; scale bar=200 mm.
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PLATE II
Black-and-white half-tones of unstained and stained stomata of Neuropteris cordata. 1. Unstained stoma; slide Nc.4.09; scale bar=6.25 mm. 2. Stoma double stained with Neutral Red and Malachite Green G; slide Nc.MaGNr.1; scale bar=8.5 mm. 3. Stoma stained with Malachite Green G; the arrow indicates the cutinization around the polar contact of the two guard cells; slide Nc.30MaG; scale bar=6.25 mm. 4–6. Stomata stained with Safranin T; slide Nc.30Sa; scale bars=22.5 mm (4), 6.5 mm (5), and 6.25 mm (6).
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colour, respectively, indicating different cutinizations, are easily discernible (e.g. Plate I, 4), and this is of particular interest in the analysis of very delicate and transparent cuticles or epidermal features. For example, cutinization details of the complex stomatal apparatus of Neurodontopteris auriculata could only be studied and sufficiently documented after staining the transparent cuticles with Methylene Blue ( Krings, 1999). Excessive stain can be washed out with warm water. Cuticle slides are long-lasting and do not fade. Black-andwhite photography is best accomplished on panchromatic film. 3.2.4. Methyl Green This stain does not appear to be useful for late Palaeozoic pteridosperm cuticles. The stain is intensive and does not normally improve the characterization of epidermal features, and black-andwhite photography is almost impossible. However, a particularly interesting result was seen when cuticles of Lescuropteris genuina were stained with Methyl Green. The maceration of plant remains in Schulze’s solution normally destroys internal structures, which may be preserved in the coaly layer between the cuticles of pinnules and axes. Some of these structures can be of interest for palaeoecological evaluations, e.g. the number of vascular strands in axes and/or the width of vascular strands in comparison to the width of overlying costal fields etc. Generally, in macerated cuticles of L. genuina, the patterns of the vascular strands and their width cannot be followed ( Krings and Kerp, 1997a and Plate I, 6). By staining the cuticles in Methyl Green, these features could be easily discerned. Above and below the original courses of the vascular strands, an intensive staining of the cuticle indicates the exact position of the vascular strands (Plate I, 8). However, this staining result fades within a few weeks. 3.2.5. Safranin T This is perhaps the best multipurpose stain to use with late Palaeozoic pteridosperm cuticles. Apart from its excellent qualities in emphasizing existing contrasts, it can be used in the analysis of complex three-dimensional structures, particularly
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stomatal complexes, because it enhances cell borders and varying amounts of cutinization, even the finest details (compare Plate II, 1 with Plate II, 4– 6). For example, the tiny stomatal complexes of Lescuropteris genuina were first recorded after staining with Safranin T ( Krings and Kerp, 1997a). Excessive stain can be washed out in warm water. The stain is fairly long-lasting; however, it will become increasingly pale and diffuse with time. Photography on panchromatic black-andwhite film is not obstructed if the features are not too intensively stained; the surface views of the stomatal complexes of Blanzyopteris praedentata ( Krings and Kerp, 1999) and Neuropteris cordata (Plate II, 4–6) were best documented after staining with Safranin T. Colour representation on daylight film is also acceptable, but not as effective as black-and-white photography (e.g. Plate I, 5).
3.2.6. Double staining in Neutral Red and Malachite Green G In this stain schedule, cuticles are first treated with Neutral Red for 1–4 min, then briefly rinsed in warm water, and after that immediately treated with Malachite Green for approximately 30 s. Excessive stain can be removed with warm glycerine. This procedure produces excellent results in the analysis of complex epidermal structures. The distribution patterns of blended hues of red, green, blue, and violet provide an excellent three-dimensional impression of a structure (e.g. Plate I, 3). For example, this stain resulted in the ability to resolve the complex three-dimensional structure of the peltate glandular trichomes of Barthelopteris germarii and their relationship to the epidermis (compare Plate I, 2 with Plate I, 3). Moreover, in some species (e.g. Lescuropteris genuina, Odontopteris minor f. zeilleri), the stain highlights different pinnule areas, i.e. pinnule margin, costal and intercostal fields (compare Plate I, 6 with Plate I, 7). Unfortunately, cuticles stained in this manner usually fade within a couple of months. The colour photography of stained cuticles often yields spectacular results, while significant light irregularities make black-and-white photography almost impossible (e.g. Plate II, 2).
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4. Concluding remarks
References
Experiments carried out on pteridosperms from the Blanzy-Montceau Basin (Central France) indicate that the use of stains can contribute considerably to the enhancement of features used in studies of the epidermal anatomy of late Palaeozoic pteridosperms. The use of distilled water and/or glycerine during the staining process makes it possible to stain fragile cuticles in the absence of rapidly dehydrating substances. However, using this procedure, cuticles must be photographed and studied immediately since the cuticle stain often leaches out. Moreover, black-and-white photography of cuticles can be obstructed by the presence of stain. However, with further advancements, e.g. digital photography adjustment, the latter problem could possibly be reduced. The results obtained by the staining of pteridosperm cuticles from BlanzyMontceau indicate that no single stain can generally be recommended for late Palaeozoic pteridosperm cuticles; the effect of staining may be different for various species, even from the same locality. Different stains must be tested on the cuticles under examination. However, Bismarck Brown and Safranin T appear to be the most universal stains for fossil plant materials.
Barthel, M., 1962. Epidermisuntersuchungen an einigen inkohlten Pteridospermenbla¨ttern des Oberkarbons und Perms. Geologie 11, Beiheft 33, 1–140. Cleal, C.J., Shute, C.H., 1995. A synopsis of neuropterid foliage from the Carboniferous and Lower Permian of Europe. Bull. Nat. Hist. Mus. Lond. (Geology) 51, 1–52. Esau, K., 1965. Plant Anatomy. Wiley, New York. 767 pp. Holloway, P.J., 1982. Structure and histochemistry of plant cuticular membranes: an overview. In: Cutler, D.F., Alvin, K.L., Price, C.E. ( Eds.), The Plant Cuticle. Academic Press, London, pp. 1–32. Kerp, H., 1990. The study of fossil gymnosperms by means of cuticular analysis. Palaios 5, 548–569. Kerp, H., Barthel, M., 1993. Problems of cuticular analysis of pteridosperms. Rev. Palaeobot. Palynol. 78, 1–18. Kerp, H., Krings, M., 1999. Light microscopy of fossil cuticles. In: Jones, T.P., Rowe, N.P. (Eds.), Fossil Plants and Spores: Modern Techniques. Special Publications Geological Society, London, pp. 52–56. Krings, M., 1997. Mo¨glichkeiten und Grenzen der Kutikularanalyse — das Beispiel der Samenfarne aus dem Stefan (Oberkarbon) von Blanzy-Montceau (Zentralfrankreich). Vero¨ffentlichungen des Museums fu¨r Naturkunde Chemnitz 20, 57–70. Krings, M., Kerp, H., 1997a. Cuticles of Lescuropteris genuina from the Stephanian ( Upper Carboniferous) of Central France — evidence for a climbing growth habit. Bot. J. Linn. Soc. 123, 73–89. Krings, M., Kerp, H., 1997b. An improved method for obtaining large pteridosperm cuticles. Rev. Palaeobot. Palynol. 96, 453–456. Krings, M., Kerp, H., 1998. Epidermal anatomy of Barthelopteris germarii from the Upper Carboniferous and Lower Permian of France and Germany. Am. J. Bot. 85, 553–562. Krings, M., 1999. Zum Bau der Spalto¨ffnungsapparate von Neurodontopteris auriculata (Brongniart) Potonie´, einer Pteridosperme aus dem Stefan von Blanzy-Montceau (Zentralfrankreich). Mu¨nstersche Forschungen zur Geologie und Pala¨ontologie 86, 69–78. Krings, M., Kerp, H., 1999. Morphology, growth habit, and ecology of Blanzyopteris praedentata (Gothan) nov.comb., a climbing neuropteroid seed fern from the Stephanian of Central France. Int. J. Plant Sci. 160, 603–619. Langiaux, J., 1984. Flores et faunes des formations supe´rieures du Ste´phanien de Blanzy-Montceau (Massif Central franc¸ais) — stratigraphie et pale´oe´cologie. Revue pe´riodique de ‘la Physiophile’ Socie´te´ d’e´tudes des sciences historiques de Montceau-les-Mines. Suppl. to Vol. 100. Martin, J.T., Juniper, B.E., 1970. The Cuticles of Plants. Arnold, London. 337 pp. Napp-Zinn, K., 1966. Anatomie des Blattes. I. Blattanatomie der Gymnospermen. In: Zimmermann, W., Ozenda, P., Wulff, H.D. (Eds.), Handbuch der Pflanzenanatomie. Gebr. Borntraeger, Berlin, Spezieller Teil, Band VIII, Teil 1, 369 pp. Skoss, J.D., 1955. Structure and composition of the plant cuticle in relation to environmental factors and permeability. Bot. Gazette 117, 55–72.
Acknowledgements This study was carried out at the University of Mu¨nster (Germany) in conjunction with a research project supported by the German Science Foundation (DFG grants Ke 584/2-1 and Ke 584/2-2), and was completed at the University of Kansas (Lawrence, KS, USA) with the support of the Alexander von Humboldt Foundation in the form of a Feodor Lynen Research Fellowship given to the author. The author is grateful to Dr J. Langiaux (Gourdon, France) and Dr G. Pacaud (formerly Autun, France) for making available most of the specimens studied. The author wants to thank Dr H. Kerp (Mu¨nster, Germany) for helpful discussion and Dr T.N. Taylor (Lawrence, KS, U.S.A.) for critical reading of the manuscript.