Heidemarie Halbritter · Silvia Ulrich Friðgeir Grímsson · Martina Weber · Reinhard Zetter Michael Hesse · Ralf Buchner · Matthias Svojtka Andrea Frosch-Radivo

# Illustrated Pollen Terminology

*Second Edition*

Illustrated Pollen Terminology

Heidemarie Halbritter • Silvia Ulrich Friðgeir Grímsson • Martina Weber Reinhard Zetter • Michael Hesse Ralf Buchner • Matthias Svojtka Andrea Frosch-Radivo

# **Illustrated Pollen Terminology**

Second Edition

Heidemarie Halbritter Division of Structural and Functional Botany Department of Botany and Biodiversity Research University of Vienna Vienna Austria Friðgeir Grímsson Department of Palaeontology University of Vienna Vienna Austria Reinhard Zetter Department of Palaeontology University of Vienna Vienna Austria Ralf Buchner Division of Structural and Functional Botany Department of Botany and Biodiversity Research University of Vienna Vienna Austria Andrea Frosch-Radivo Division of Structural and Functional Botany Department of Botany and Biodiversity Research University of Vienna Vienna Austria Silvia Ulrich Division of Structural and Functional Botany Department of Botany and Biodiversity Research University of Vienna Vienna Austria Martina Weber Division of Structural and Functional Botany Department of Botany and Biodiversity Research University of Vienna Vienna Austria Michael Hesse Division of Structural and Functional Botany Department of Botany and Biodiversity Research University of Vienna Vienna Austria Matthias Svojtka Division of Structural and Functional Botany Department of Botany and Biodiversity Research University of Vienna Vienna Austria

This book is an open access. ISBN 978-3-319-71364-9 ISBN 978-3-319-71365-6 (eBook) https://doi.org/10.1007/978-3-319-71365-6

Library of Congress Control Number: 2018946142

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### **Preface to the First Edition**

*There are more things in heaven and earth, than are dreamt of in our philosophy.* Shakespeare, Hamlet to Horatio

The principal aim in compiling this book was to provide the reader with firsthand information about the structure and outlook of the extremely manifold pollen in seed plants. This book should not be seen as a mere collection of striking and/or informative light and electron micrographs. Each of the micrographs is intended to convey a specific message related to properties and functions of the pollen grains shown. The authors hope that the book will be useful for experienced researchers as well as for beginners in palynology, but also for medicine, biochemistry, or even for lawyers and artists as an aid and guide for the evaluation and interpretation of pollen features.

Vienna, Austria Michael Hesse Heidemarie Halbritter Reinhard Zetter Martina Weber Ralf Buchner Andrea Frosch-Radivo Silvia Ulrich

### **Preface to the Second Edition**

The first edition of this book *Pollen Terminology: An illustrated handbook* was extremely successful and well received by the worldwide palynological community. As with the first edition, our main intention when compiling the second edition was to provide both scientists and the public with an easily understandable and primarily illustrative access to the hidden beautiful world of pollen and the fascinating subject of palynology. This new edition titled *Illustrated Pollen Terminology* allowed us to improve many aspects of our book and to illustrate in more detail the important concepts and various preparation techniques applied in (paleo)palynology. It is our hope that this edition will become the guidance tool for all students of palynology, as well as reference work and illustrated encyclopedia for the more advanced scientists.

Vienna, Austria Heidemarie Halbritter Silvia Ulrich Friðgeir Grímsson Martina Weber Reinhard Zetter Michael Hesse Ralf Buchner Matthias Svojtka Andrea Frosch-Radivo

### **Contents**

#### **Part I General Chapters**







#### **Abbreviations**


#### **Prefixes**



#### **Introduction**

*Illustrated Pollen Terminology* is a collection of palynological terms and wellillustrated with light and electron microscope images. The focus of this book is on the pollen of seed plants, predominantly angiosperms; therefore, it rarely explains features unique to spores or gymnosperms. A strict rationalization of terms on the basis of practical criteria has been attempted for this book. Where necessary, definitions have been reworded, newly circumscribed, or brought into focus. In addition, consistent application of EM techniques and the nowadays better understanding of pollen features have made redefinition of some terms necessary.

Since 1994, the *Glossary of Pollen and Spore Terminology*, by W. Punt, S. Blackmore, S. Nilsson, and A. L. Thomas, was the standard reference in palynology (Punt et al. 1994). In 1999 the online version by Peter Hoen appeared, with several additions. A new version published in 2007 provided informative schematic drawings containing the essentials of each term, mostly using LM observations (Punt et al. 2007). Although extremely useful for overview purposes, drawings cannot show the full range of features. This can only be achieved with various LM, SEM, and TEM micrographs, which demonstrate the stunning diversity of features as seen in this book.

This book is divided into four parts. The first part comprises the "General Chapters". The first chapter "Palynology: History and Systematic Aspects" provides a comprehensive overview of the history of palynological research and the origin and development of categories and classification systems as well as the systematic value of pollen. The following chapter "Pollen Development" explains the formation and development of a pollen grain (microsporogenesis and microgametogenesis). The third chapter "Pollen Morphology and Ultrastructure" gives a thorough overview on all aspects of pollen features, both structure and sculpture, that need to be considered when studying pollen grains. There are many features observed with microscopes that can be misleading or misinterpreted, most of them have been summarized in the consecutive chapter "Misinterpretations in Palynology." In the fifth chapter "How to Describe and Illustrate Pollen Grains" examples are given as to how to properly present palynological data. The chapter thereafter "Methods in Palynology" includes detailed and illustrated protocols of most methods and techniques used when studying recent and fossil pollen grains with LM, SEM, and TEM. The second part "Illustrated Pollen Terms" is the main part of this book, and comprises 6 chapters. All terms in this part are defined and comprehensively illustrated, and if necessary features are highlighted for easy recognition. The world's most comprehensive database on recent pollen, *PalDat* (http://www.paldat.org/), is the main source of pictures. Each term is illustrated with LM or EM pictures in order to point out the character range of a term (or, more precisely, to show the full range of a single character). In the third part, all terms are listed along with their definition in the "Glossary of Palynological Terms". Numbers following terms refer to the respective page(s) where the terms are discussed. Numbers in bold relate to illustrations in the chapters of the "Illustrated Pollen Terms." The fourth part "Annex" comprises the "Picture Copyrights" and the "Index" listing all plant names occouring in this book.

# **General Chapters**

#### **Contents**

**Palynology: History and Systematic Aspects – 3 Pollen Development – 23 Pollen Morphology and Ultrastructure – 37 Misinterpretations in Palynology – 67 How to Describe and Illustrate Pollen Grains – 85 Methods in Palynology – 97**

© Springer International Publishing AG 2018

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, https://doi.org/10.1007/978-3-319-73482-8

### **Palynology: History and Systematic Aspects**

**The History of Palynology – 4**

**Categories, Classification Systems and Systematic Value of Pollen Features – 9**

**Future Perspective – 16**

**References – 18**

3

Palynology is the science of palynomorphs, a general term for all entities found in palynological preparations (e.g., pollen, spores, cysts, diatoms). A dominating object of the palynomorph spectrum is the pollen grain. The term palynology was coined by Hyde and Williams (1955; Fig. 1). It is a combination of the Greek verb paluno (пαλύνω, "I strew or sprinkle"), palunein (пαλύνειν, "to strew or sprinkle"), the Greek noun pale (пαλƞ, in the sense of "dust, fine meal," and very close to the Latin word pollen, meaning "fine flour, dust"), and the Greek noun logos (λογος, "word, speech").

#### **The History of Palynology**

Assyrians are said to have known the principles of pollination (they practiced hand pollination of date palms), but it is unclear if they recognized the nature of pollen itself. The invention of the first microscopes and especially the compound microscope in the late sixteenth century represents the starting point of a new fascinating era. Some of the most important findings and scientists within the long tradition of light microscopy are mentioned here. For a more comprehensive overview, see Wodehouse (1935) and Ducker and Knox (1985).

Following the invention of the simple microscope by J. Janssen and Z. Janssen in 1590, the first compound microscope was developed by Hooke (1665). This was an important contribution to the study of pollen morphology. Malpighi in his "Anatomia Plantarum" was the first to describe pollen grains as having germination furrows while Grew noted in his famous work "The Anatomy of Plants" the constancy of pollen characters within the same species (Fig. 2; Grew 1682; Malpighi 1901). They are both considered the founders of pollen morphology. Camerarius described several pollination experiments and communicated the results in his letters about plant sexuality to Valentini (Camerarius 1694). He stated that male "seed dust" is necessary for seed development. Von Linné (also known before his ennoblement as Carl Nilsson Linnæus) first used the term pollen (in 1750). In the 18th and the early nineteenth centuries, there was considerable progress in pollen research and the understanding of pollination. In 1749, Gleditsch demonstrated in a spectacular experiment (Experimentum Berolinense) the central role of pollen in double fertilization. He organized the transport of an inflorescence from a male fan palm in Leipzig to a hitherto "sterile" female fan palm growing in a greenhouse in Berlin. After pollination, the female flowers produced fertile seeds for the first time in the palms lifetime (Gleditsch 1751, 1765). Placing male inflorescences within groups of female date palms for pollination was according to Theophrast already

**Fig. 1 The right word**. Excerpt from Hyde and Williams (1955). Pollen Analysis Circular no. 8, p. 6

**Fig. 2 First drawings of pollen**. Grew (1682) "The anatomy of plants"

practiced by the Assyrians and Egyptians. Kölreuter, together with Sprengel, the founder of research on flower biology, perceived the importance of insects in flower pollination and discovered for the first time that pollen plays an important role in determining the characters of the offspring (Kölreuter 1761-1766; Sprengel 1793). Kölreuter (1806, 1811) also discovered that the pollen wall is consisting of two distinct layers and made the first attempt to classify pollen based on their morphology. Sprengel recognized pores and furrows in the pollen wall and demonstrated the effects of cross pollination, dichogamy, and distinguished between entomo- and anemophily (Candolle and Sprengel 1821). Moreover, he also realized that every plant species has a characteristic pollen type (Sprengel 1804).

During the first half of the nineteenth century, some fundamental insights into pollen morphology and physiology were achieved. Purkinje made the first attempt for a palynological terminology by classifying pollen based on their morphology (Purkinje 1830). Wodehouse (1935) pointed out that "Purkinje's system of nomenclature deserved much more attention than was ever given to it by subsequent investigators. A system of this kind, had it been put into use, would have saved much confusion." Brown gave the first description of the origin and role of the pollen tube (Brown 1828, 1833). He credited Bauer as the first observer of the pollen tube's nature, of the double wall in *Asclepias* pollen, and for his minute drawings of *Asclepias* pollen. His brother Bauer, a great botanical artist, was the first to recognize compound pollen in *Acacia* and orchids. Cavolini described and illustrated the filiform pollen of sea grasses *Zostera* and *Cymodocea* (Cavolini 1792).

Göppert and Ehrenberg were the first to describe and depict fossil pollen grains and spores (Göppert 1837, 1848; Ehrenberg 1838). In 1834 von Mohl wrote his fundamental work entitled "Über den Bau und die Formen von Pollenkörner/On the structure and diversity of pollen grains," which was a major contribution to the knowledge of pollen structure and descriptive classification. von Mohl and Fritzsche recognized the principal layers of the pollen wall and published new surveys on pollen morphology (von Mohl 1835; Fritzsche 1837). The term pollenin goes back to von Grotthuss (1814), John (1814), Stolze (1816), and Fritzsche (1834). The terms "exine," "intine," and "Zwischenkörper" were established by Fritzsche and published in his book "Über den Pollen" (Figs. 3 and 4; Fritzsche 1837). He also demonstrated that apertures are predetermined in most angiosperm pollen while others are inaperturate. Zetzsche first coined the term "sporopollenin" to describe the resistant chemical substance present in the outer wall of both pollen grains and spores (Zetzsche and Huggler 1928; Zetzsche and Vicari 1931; Zetzsche et al. 1931). Campbell reported pollen of seagrasses (*Naias* and *Zannichellia*) to be thin walled, without exospore (exine), and two-celled. Moreover, Campbell described the mitotic division of the generative cell into two (sperm) cells (Campbell 1897). Hofmeister and Strasburger provided ground-breaking insights into the development and internal structure of pollen and fertilization (involves the fusion of a single sperm nucleus and the egg nucleus) and investigated the bi- and tricellular pollen condition of many angiosperms (Strasburger 1884; Hofmeister 1849). The role of the second sperm nucleus in the pollen tube remained unexplained until double fertilization was discovered by Guignard (1891, 1899); Nawaschin (1898). Nägeli studied the ontogeny of pollen grains within anthers and was the first to recognize the callose wall (Nägeli 1842). Schacht described differences in exine patterning, exine thickness, and apertures covered by an operculum. He also used cytochemical staining techniques to detect pollen reserves. He was also the first to cut sections of embedded pollen with razor blades for anatomical studies (Schacht 1856/59). Strasburger described the basic concepts of pollen wall development already in 1889, but major break-through in pollen wall ontogeny was achieved much later by Heslop-Harrison (1975). The first successful classification of orchidaceous plants based on pollen features was made by Lindley (1836). Later, Fischer recognized the potential of pollen morphology in aiding the phylogenetic position of angiosperms (Fischer 1890).

Paleopalynology was established at the end of the nineteenth century, when P. Reinsch published the first photomicrographs of fossil pollen and spores from Russian coals (Reinsch 1884). He also described methods for the extraction of palynomorphs from coal samples with concentrated potassium hydroxide (KOH) and hydrofluoric acid (HF). Von Post published the first pollen diagram (profile) using exclusively arboreal pollen (von Post 1916). Already before and especially after the Second World War, Schopf as well as Potonié published their impressive publications devoted to fossil spores and pollen (e.g., Potonié 1956; Schopf 1957, 1964). Schopf established the systematic study of palynomorphs, while Potonié was one of the first who recognized the stratigraphic value of paleopalynology, applying his "turmal classification" system (Potonié 1934; see also "The Treme System and the NPC-Classification" below). The rise of stratigraphic palynology started shortly before 1950 and played a prominent role in petroleum explorations during the second half of the twentieth century (Manten 1966).

**Fig. 3 Detailed drawings of pollen**. Fritzsche (1837) "Über den Pollen"

The key role of palynology in stratigraphy depends upon the fact that the natural biopolymer sporopollenin in the spore/pollen walls is extremely resistant; thus, pollen/spores are often abundantly preserved in sedimentary rocks.

The twentieth century up to ca 1960 was dominated by the skillful use of the LM, with many new findings; for example, the LO-analysis, a method for analyzing patterns of exine organization by light microscopy, focusing at different levels distinct

**Fig. 4 First detailed drawings of pollen**. Fritzsche (1837) "Über den Pollen"

features appear bright (*L* = Lux) or dark (*O* = Obscuritas). Textbooks by Wodehouse, Erdtman, or Fægri and Iversen summarized the knowledge of palynology from that time, but are still in good use (Erdtman 1943, 1952, 1957, 1969; Fægri and Iversen 1950, 1989; Wodehouse 1935). During this time palynology also became more diverse and applied in numerous fields among others: aeropalynology, biostratigraphy, copropalynology, cryopalynology, forensic palynology, iatropalynology, melissopalynology, paleopalynology, archeology, paleoclimatology, and palynotaxonomy.

Electron Microscopy with its two most important instrument types, the Transmission Electron Microscope (TEM) and the Scanning Electron Microscope (SEM), facilitated major breakthroughs in palynology. The TEM revealed new and stunning insights into pollen wall development and stratification. This prompted authors to publish new descriptions and create new terms. As pointed out by Knox: "The terminology applied to the pollen wall is daunting, especially as it has been developed from early light microscopy work, and then transposed to the images seen in the transmission and scanning electron microscopes" (Knox 1984, p. 204).

One of the first reports on the ultrastructure of recent pollen using TEM were published by H. Fernandez-Moran and A. O. Dahl (1952), and by K. Mühlethaler (1953). The first reports on the ultrastructure of fossil pollen were published by Ehrlich and Hall (1959; Pettitt and Chaloner (1964). During the 1950s and early 1960s considerable progress in TEM preparation methods (from fixation to microtome sectioning and staining) took place. EM-based information on ornamentation details of pollen grains was rare up to the mid-1960s. Only TEM-based casts or replica methods were available, all of them with limited resolution and depth of focus (e.g., the single-stage carbon replica technique; Mühlethaler 1955; Bradley 1958; Rowley and Flynn 1966). The time-consuming and laborious TEM replica procedures were an obstacle to extensive surveys of pollen morphology and later replaced by SEM (Harley and Ferguson 1990). The introduction of SEM in palynology in the mid of the 1960s was a key innovation in the study of the fine relief (sculpture) of pollen and spore surfaces. Advantages of SEM include the relatively simple and rapid preparation methods and the supreme depth of focus. SEM was considered from the very first moment as the quantum leap in EM (Hay and Sandberg 1967). The first SEM micrographs of pollen grains were published by Thornhill et al. (1965) and Erdtman and Dunbar (1966). Since then palynologists have been provided with a plethora of beautiful micrographs. Like Blackmore noted "The scanning electron microscope has provided a greater impetus to palynology than any other technical development during the history of the subject." Blackmore (1992). The LM with basic and advanced equipment, such as the fluorescent super-resolution microscopy, is overcoming the Abbe limit of LM resolution (especially STED microscopy, Hell 2009). The super-resolution LM and the two main types of EM form an expedient combination of imaging techniques. The LM remains the "workhorse method" (Traverse 2007; see the compendia by Reille 1992, 1995, 1998), but is limited regarding various morphological and structural features. Therefore, the role of SEM as an essential part in illustrating exine sculpture and ornamentation cannot be overrated (Harley and Ferguson 1990). The TEM still plays an important role, for example, in elucidating the complex steps of exine formation and development (e.g., Blackmore et al. 2007, 2010; Gabarayeva and Grigorjeva 2010; Gabarayeva et al. 2010).

The first and especially the second half of the twentieth century saw palynology at its peak, combining light microscopy with electron microscopy techniques. In addition to the above-mentioned scientists, other great palynologists have also promoted our science toward its present multifaceted appearance. These include among others: B. Albert, H.-J. Beug, G. El Ghazali, F. Firbas, M. Harley, J. Jansonius, W. Klaus, G. O. W. Kremp, B. Lugardon, S. Nadot, A. Maurizio, J. Muller, S. Nilsson, J. R. Rowley, J. J. Skvarla, H. Straka, G. Thanikaimoni, R. H. Tschudy, M. van Campo, T. van der Hammen, and A. Le Thomas.

#### **Categories, Classification Systems and Systematic Value of Pollen Features**

F or the scientist, categories are essential for classifying natural characters in their diversity, defining their range and placing them in a systematic order. In addition to the theoretical concept, categorization always depends on the manner in which a feature is perceived: i.e. on the **visibility** of a feature, and/or their specific value. Categorization also greatly depends on the technical equipment and method(s) used, as well as on the **subjective interpretation** of character(s) (see "Methods in Palynology"). Thus, categorization of features is difficult to standardize. An example is the category **pollen size**: there is not just a natural size variation within a single anther/flower/taxon, dimensions may also vary depending on the preparation method(s) used, and the observer's evaluation. Moreover, sometimes the size of a pollen grain is just at the boundary between two adjacent pollen size categories (for size categories: see "Pollen Morphology and Ultrastructure").

When describing and categorizing pollen, two basic groupings are known from the literature: pollen type and pollen class. **Pollen type** is a general term categorizing pollen grains by a distinct combination of characters and is used in connection with systematics, affiliating the pollen type with a distinct

**Fig. 5 Pollen type vs pollen class**. **A**-**B**. *Polygonum aviculare*, Polygonaceae; all *Polygonum* pollen sharing the combined features observed here belong to the *Polygonum* aviculare type. This pollen can also be included in the pollen class "tricolporate". **C**-**D**. *Leontodon saxatilis*, Asteraceae; all Asteraceae pollen sharing the combined features observed here (lophate, tricolporate, echinate) belong to the *Leontodon* type, characteristic for the "Liguliflorae" group within the Asteraceae. This pollen can also be included in the pollen class "tricolporate"

taxon/a (e.g., *Polygonum aviculare* type/*Leontodon* type, Fig. 5). The term "pollen type" is sometimes (colloquially) misused: for example, *Croton* type, which is a distinct feature of ornamentation and is correctly termed *Croton* pattern.

**Pollen class** is an artificial grouping of pollen grains that share a single or more, distinctive characters (see "Illustrated Pollen Terms"). Pollen classes can refer to pollen units (e.g., polyads, tetrads), to shape (e.g., saccate, polygonal, heteropolar, arcus), to aperture type and location (e.g., inaperturate, sulcate, ulcerate, colpate, colporate, porate, synaperturate, spiraperturate), or to an extremely distinctive ornamentation character (e.g., lophate, clypeate). These classes can be useful in identification keys as they have a good diagnostic, although mostly no systematic, value. In general, a pollen grain may belong to more than one pollen class; in such cases, the more significant feature should be ranked first (e.g., *Pistia*: plicate-inaperturate, *Hemigraphis*: plicate-colporate, *Typha*: tetradsulcerate, *Rhododendron*: tetrads-colporate).

Many terms in palynology were coined at a time when only LM observations were available. Mainly for historical reasons, inconsequent nomenclatural applications, enumerations of synonyms, and even differing definitions have been found for one and the same term. During the twentieth century, questions of terminology became more and more problematic. The main reasons were the increasing numbers of publications in palynology, dealing with sometimes insufficiently described or "uncommon" pollen features, and simultaneously the advent of manifold applied fields of palynology. For various

**Fig. 6 Nomenclature in Paleopalynology**. **A**. *Oculopollis* sp., fossil, Upper Cretaceous, Hungary, polar view. **B**. *Oculopollis* sp., fossil, Upper Cretaceous, Hungary, polar view. **C**. *Trudopollis* sp., fossil, Upper Cretaceous, Hungary, polar view. **D**. *Trudopollis* sp., fossil, Upper Cretaceous, Hungary, polar view

reasons, nearly all authors used their own terminology. Nonetheless, in the 1950s attempts were made to restrict the wording and to state the definitions of terms more precisely (Erdtman 1947; Erdtman and Vishnu-Mittre 1956). A limited list of pollen morphological terms and definitions was published as early as 1950 by Iversen and Troels-Smith. Later, Kremp (1968), in his famous encyclopedia, provided a monumental enumeration of all known terms. Reitsma (1970) took the first resolute step to overcome the problem of synonyms in palynological terminology, though unfortunately not taking into account the variation range of palynological features. Fægri and Iversen (1989, 4th ed.) restricted their glossary of terms exclusively used in their book. Moore et al. (1991, 2nd ed.) provided a glossary of selected terms used in their pollen and spore keys. Standardization came with the glossary by Punt et al. (1994, 2007). The main advance of their concise and comprehensive terminology is the consistent use of drawings and the critical comments on terms.

A complex category issue in (Paleo-) Palynology is the nomenclature question. In Paleopalynology, for morphotaxa often form-generic names are used. The nomenclature of form-genera is either artificial when the relationship is not known at all (e.g., *Oculopollis* and *Trudopollis* from the Normapolles group, Fig. 6), or semi-biological, when reference to an extant taxon is suspected but not proven (e.g., *Liliacidites*). However, if reference to extant taxa is certain, then a biological nomenclature is possible (e.g., *Quercus* sp.).

#### **The Turmal System**

A quite different classification and nomenclature is **Potonié's turmal system**. This is an artificial, informal, neutral suprageneric classification scheme for fossil (especially Carboniferous or Permian) pollen and spores. It is subdivided into a hierarchy of progressively finer units (ranks): anteturma, turma, subturma, infraturma, subinfraturma, and corresponds mostly to morphological features (for details see Traverse 2007).

#### **The Treme System and the NPC-Classification**

The **"-treme system**" of aperture configuration as an alternative or an addition to the traditional nomenclature was introduced by Erdtman and Straka (1961). The suffix -treme is derived from trema (pl. tremata) and is synonymous with aperture. In combination with prefixes such as cata-, ana-, zono-, and panto-, the position of germination sites in relation to pollen polarity can be designated. Catatreme indicates the proximal, anatreme the distal, zonotreme the equatorial, and pantotreme the global position of apertures. Other prefixes such as mono-, di-, tri-, tetra- indicate the number of apertures irrespective of their position.

The **NPC-classification** by Erdtman and Straka (1961), resting upon the -treme system, is a morphological system for classifying pollen and spores. This system is based on the aperture features: their number (N), position (P), and character (C). Their NPC-system for spore/pollen classification was used as a diagnostic tool in systematics. As an example, the three apertures (N3) of pollen grains, having a zonotreme position (P4) and being colporate (C5) have the NPC-formula 345 (Fig. 7; Erdtman and Straka 1961). Taxa with the same general NPCformula are grouped together, those showing a different formula, separately. This system does not work in, e.g., heteroaperturate or inaperturate (formula 000) pollen, or pollen tetrads. Unfortunately, the NPC-system ignores other pollen characters including shape and ornamentation that are indispensable for a complete description.

#### **Systematic Value of Pollen Features**

One of the main research interests in palynology focuses on the taxon-specific patterns of the pollen wall, how they developed and evolved. Moreover, pollen can provide phylogenetic evidence important to plant systematics (Hesse and Blackmore 2013). The reconstruction of phylogenies has continuously developed. The advances in modern phylogenetic approaches are resulting in constant changes in plant systematics, even whole genomes are being used together with multiple DNA analyses for a better insight into relationships (Stuessy and Funk 2013). Critically evaluated pollen features may be a useful tool for systematics with a significant diagnostic value, supporting or contradicting the results of molecular studies ("The palynological compass" sensu Blackmore 2000; Hesse and Blackmore 2013). Palynological features are very valuable, especially in delimiting taxa (Ulrich et al. 2012). Regarding multiple-gene tree studies with conflicting results, pollen data combined with other morphological evidence (e.g., floral characters) have more recently become an important indicator of which tree may be the best representative (Stuessy and Funk 2013; Ulrich et al. 2012, 2013). Furthermore, pollen morphological studies proved to be indispensable for the understanding of evolutionary

**Fig. 7 NPC-classification of pollen**. **A-B.** *Androsace chamaejasme*, Primulaceae, a tricolporate pollen with the formula N3P4C5

processes and systematics. For taxonomic studies, pollen features that have value for the lower and higher taxonomic levels should be obtained by a combined study using LM, SEM, and TEM (Stuessy 1979).

Alternation of generations is a unique feature of plants that occurs in green algae, mosses, ferns, gymnosperms, and angiosperms. Pollen grains develop in anthers as the result of meiosis and mitoses (two in angiosperms, three to five in gymnosperms) and represent an extra generation, the highly reduced male gametophyte. Therefore, pollen grains are not simply small parts of a plant like leaves or seeds; they are the complete (hidden) haploid counterpart to the more dominant plant, which represents the diploid generation (Kessler and Harley 2004). During dispersal, pollen grains are completely separated from the parent plant and perfectly adapted for their role — the transfer of male genetic material — and are able to resist hostile environmental stresses on their way to the female flower parts. Usually, pollen does not suffer to the same extent from the various and harsh selective pressures to which the diploid plant is subjected. Because selective pressures (e.g., temperature, precipitation) upon pollen characters are predominantly absent or low, compared to those on the diploid plant, pollen features may remain constant for millions of years, meaning pollen features can be conservative and of taxonomic value (Wodehouse 1928, 1935; Hao et al. 2001; Grímsson et al. 2014, 2016, 2017a, b). Therefore, identical and rare conditions in fossil vs recent pollen probably belong to only one group and were not invented independently in distant groups (e.g., fossil *Spinizonocolpites* pollen and recent *Nypa* pollen, Arecaceae; Zetter and Hofmann 2001; Gee 2001). Selective pressures might concern especially the pollen aperture number, but also the pollen sculpture and the mode of pollination ecology (Furness and Rudall 2004). Pollen features are, if used for a systematic purpose, at least as important as any other morphological character of the diploid generation. For this reason pollen morphology claims a crucial role in, e.g., systematics and palynostratigraphy, for example in elucidating the early history of angiosperms. Angiosperm pollen from the Early Cretaceous are usually sulcate (typical for basal angiosperms) with a columellate infratectum (which is restricted to angiosperms). The first appearance of dispersed tricolpate pollen, typical for eudicots, is not known before the latest Barremian, is rare in the Aptian of Southern Laurasia and Northern Gondwana, but is ubiquitous in the Albian of both provinces. Tricolporate pollen appears first in the late Albian, and triporate pollen in the middle Cenomanian (Doyle and Endress 2010; Friis et al. 2011; Doyle 2012). For a detailed overview of structural pollen diversification and of the stratigraphic appearance of major angiosperm pollen types during the Cretaceous, see Friis et al. (2011) and Mendes et al. (2014).

Palynological data may be helpful at all levels of systematics, especially in angiosperms (c.f. Stuessy 2009). When pollen of a taxon (representing family/ ies or genus/era) is characteristic and similar among species they are termed **stenopalynous** (Fig. 8), and occur, for example, in Poaceae, Lamiaceae, Asclepiadaceae, Brassicaceae, Asteroideae, and Cichorioideae. On the contrary, **eurypalynous** (Fig. 9) taxa are heterogeneous and pollen can vary among others in size, aperture, and in exine stratification. Examples for eurypalynous groups are Acanthaceae (Sarawichit 2012) and Araceae (Harley and Baker 2001; Ulrich et al. 2017).

At the highest taxonomic level (e.g., angiosperms vs gymnosperms, dicots vs monocots), a columellate exine condition occurs exclusively in angiosperms. A lamellate endexine is typical for gymnosperms, whereas the angiosperm endexine is usually not lamellate, except in immature stages (*Orobanche hederae*). But in very few cases there is a continuously lamellate endexine present, like in *Ambrosia* (Furness and Rudall 1999a, b, Weber and Ulrich 2010). In inaperturate pollen of Araceae the endexine is exceptionally thick and spongy, which may be a functional benefit and of systematic value. A strong phylogenetic signal comes from the aperture arrangement: the "tricolpate" condition is a synapomorphy for eudicots, tricolporate pollen occurs only in core eudicots, while sulcate pollen is a plesiomorphic condition in basal angiosperms (Nadot et al. 2006). Palynologists have long wondered about the two fundamental evolutionary shifts occurring at the base of the eudicot clade, both in aperture position (from distal to equatorial) and number (from one to three or more). Most probably, these changes in pollen morphology have a systematic and simultaneously a functional background. The shift from a distal, single aperture to equatorially or globally situated apertures, increases the number of possible germination sites (Furness and Rudall 2004). Pollen morphology does not support sharp delimitation between dicots and monocots, as dicotyledonous pollen characters also occur in some monocots and conversely. In early-diverging angiosperms the formation of pollen features appears to be more plastic than in dicots (especially in eudicots). Manifold combinations of pollen features are typical for basal angiosperms and even for the most basal eudicots, the Ranunculales. All of them are more or less eurypalynous. In contrast, late-divergent eudicots are often stenopalynous and

**Fig. 8 Stenopalynous taxa (family level)**. Pollen of different Poaceae all look very similar, for example in *Alopecurus* (**A**), *Cutandia* (**B**), *Dactylis* (**C**), *Fargesia* (**D**), *Poa* (**E**), *Sesleria* (**F**) the pollen is spherical, ulcerate with nano-sized sculpture elements

PALYNOLOGY: HISTORY AND SYSTEMATIC ASPECTS

**Fig. 9 Eurypalynous taxa (family level)**. **A**-**F**. Pollen of different Araceae genera look very different. **A**. *Ambrosina* pollen is plicate and inaperturate. **B**. *Dracunculus* pollen is verrucate and inaperturate. **C**. *Pinellia* pollen is echinate and inaperturate. **D**. *Cyrtosperma* pollen is reticulate and ulcerate. **E**. *Anthurium* pollen is reticulate-microechinate and diporate. **F**. *Monstera* pollen is psilate, with ring-like aperture

appear somewhat "poor" regarding the diversity of pollen features (Hesse et al. 2000). In general, the richness and variation of morphological features in pollen decreases in eudicots (Furness and Rudall 1999a). In Alismatales, many pollen features are adaptive and related to their aquatic/semiaquatic habitat, e.g., thin-walled, inaperturate pollen have evolved iteratively, even filamentous pollen is not rare (Furness and Banks 2010).

Fine example for adaptive and simultaneously systematic values is the ring-like aperture found especially in monocots, while only few occur in dicots. A ring-like aperture was probably the best way to a target-oriented harmomegathic movement, to contract or expand a large area adapted for pollen tube formation. This type of aperture might be relict of early angiosperms, before the advent of the "eudicot-tricolpates".

Examples for diagnostic features at lower taxonomic levels (family) are saccate pollen, typical for Pinaceae and Podocarpaceae. A small papilla is characteristic for Taxodioideae pollen (see "Illustrated Pollen Terms"). Another example for a strong phylogenetic signal comes from an aroid subfamily, the aperigoniate Aroideae (Araceae). They are characterized by several synapomorphies: inaperturate pollen, often with an outermost non-sporopollenin layer (exine absent) and a thick spongy endexine. The absence of callose in pollen development is the reason for this uncommon wall structure, that differs from all other currently known angiosperms (Anger and Weber 2006; Hesse 2006a, b).

At the lowest taxonomic level (genus, species) a combination of distinct morphological and structural features usually refers to a particular genus or species. Even very inconspicuous features can represent an example of systematic value, like the *Pinus* subgenus *Strobus* (Haploxylon) type and the *Pinus* subgenus *Pinus* (Diploxylon) type (see "Pollen Morphology and Ultrastructure"). Another example is the large genus *Amorphophallus* (Araceae), showing high diversity in ornamentation (e.g., Ulrich et al. 2017). As a result of the harmomegathic effect, the shape of pollen may change, which is enabled by the elasticity of the exine and infoldings of the apertures. The aperture type and arrangement may lead to characteristic infoldings. Therefore, the shape of pollen in dry state can be typical for a family or genus (e.g., Halbritter and Hesse 2004). For example, tricolporate pollen of the genus *Chaenarrhinum* (Plantaginaceae) is heteropolar. The heteropolarity is only apparent in dry condition. Also, tricolpate pollen of Lamiaceae is highly characteristic in dry condition: it is prolate, extremely flattened, and with apertures arranged in a very distinct manner (Fig. 10; see also "harmomegathic effect" in "Pollen Morphology and Ultrastructure").

#### **Future Perspective**

Nowadays, palynology serves as an indispensable tool for various applied sciences such as systematics (Doyle and Endress 2010; Dransfield et al. 2008), melissopalynology (Jones and Bryant 1996), and forensics (e.g., Mildenhall et al. 2006; Bryant 2013; Weber and Ulrich 2016), but should also stand alone as a basic field in science. In general, compared to the sporophyte the male gametophyte in seed plants is poorly investigated. From ca. 260.000 to 422.000 plant species (e.g., Thorne 2002; Govaerts 2003; Scotland and Wortley 2003; *The Plant List* currently accepts 350.699 species) only about 10% have been studied with respect to pollen grain morphology, and regarding pollen ultrastructure it is even much less. Therefore, it is important to continue classical and more advanced palynological studies.

Despite the long tradition of palynology and its application in many fields, it should be considered why it is important and where it is heading in the near future. In the twenty-first century, no matter what role palynology will play, being a basic field of science or more probably a bundle of applied fields, a vital issue will be the increase of our knowledge of pollen grains and in this context the enhancement of pollen terminology. Online pollen databases (efficient in data storage, data transmitting and dissemination) will get more and more important for the exchange of pollen and spore information (for example, *PalDat;* Weber and Ulrich 2017). Journals are nowadays published simultaneously in print as well as in electronic format, both have manifold advantages and disadvantages. Nevertheless, illustrated monographs, like this one, will retain their role of detailed information and long-living documentation.

**Fig. 10 Characteristic shape of pollen in dry condition**. **A**-**B**. *Lamium maculatum,* Lamiaceae, pollen in hydrated and dry condition. **C**-**D**. *Microrrhinum minus,* Plantaginaceae, pollen in hydrated and dry condition. **E**-**F**. *Scutellaria baicalensis,* Lamiaceae, pollen in hydrated and dry condition

#### **References**


Potonié R (1956) Synopsis der Gattungen der Sporae dispersae, I. Teil: Sporites. Beih Geol Jahrb 23: 1–103


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### **Pollen Development**

**Microsporogenesis and Microgametogenesis – 24**

**References – 35**

23

#### **Microsporogenesis and Microgametogenesis**

Pollen is source and transport unit for the male gametes (or their progenitor cell). The unicellular pollen grain represents the microspore of seed plants, the multicellular pollen grain the male gametophytic generation. The development of a pollen grain includes **microsporogenesis** and **microgametogenesis** (Figs. 1 and 2, Gomez et al. 2015; Keijzer and Willemse 1988). Microsporogenesis starts with the differentiation of microspore mother cells (MMC) respectively **pollen mother cells** (PMC). These diploid cells become enclosed by a thick **callose** wall and undergo meiosis, forming a tetrad of four haploid **microspores**, each encased in another callose wall insulating them from each other and from the surrounding diploid tapetal cells (Figs. 1 C-E, and 2). Cytokinesis following meiotic nuclear divisions is accompanied by the formation of cleavage planes determined by the configuration and orientation of the meiotic spindle axes. In the case of **successive cytokinesis**, planes are formed after the first and second meiotic divisions leading to the formation of various microspore tetrad types (see "Pollen Morphology and Ultrastructure"). During **simultaneous cytokinesis** the cleavage planes are formed simultaneously after the second meiotic division and microspores become arranged in a **tetrahedral tetrad** (Furness and Rudall 1999, 2001).

Pollen wall formation starts while the microspores are arranged in tetrads, encased by callose. The first step starts with the deposition of **primexine**, a fibrillar polysaccharidic material, on the surface of the microspores. The primexine forms a template where sporopollenin precursors and subsequently **sporopollenin** are deposited, building the final pollen wall (Fig. 1 E). Apertures are formed where the endoplasmic reticulum has prevented the deposition of primexine.

During pollen formation and maturation the **tapetum** plays an important role, usually forming a single layer of cells circumscribing the loculus. Tapetal cells are specialized and have a short lifespan. They finally lose their cellular organization and are reabsorbed. Two types of tapetum are known: the **secretory** (or glandular or parietal) and the **amoeboid** (or periplasmodial). In the secretory type (e.g., in Apiaceae) the tapetal cells remain stationary until they finish their physiological functions. In the amoeboid type (e.g., in Araceae) cells lose their individuality at an early developmental stage by degeneration of the cell walls (Furness and Rudall 1999, 2001). The protoplasts then fuse and intrude into the locule where they enclose the pollen grains (Fig. 3). The tapetum plays an important role during several stages of pollen development (Pacini 1997). Its main function is the nourishment of the microspores, but it also synthesizes enzymes (e.g., callase), exine precursors, pollen coatings, forms Ubisch bodies (orbicules) and viscin threads (both equivalents to the ektexine). The most striking material produced by the tapetum is **pollenkitt** (and **tryphine** in Brassicaceae), a sticky, heterogeneous material composed of neutral lipids, flavonoids, carotenoids, proteins and polysaccharides. Pollenkitt serves numerous functions: keeping pollen grains together during transport, protecting pollen (from water loss, ultraviolet radiation, hydrolysis and exocellular enzymes), and maintaining sporophytic proteins inside exine cavities.

Microgametogenesis (Fig. 1 G-K) in angiosperms includes first and second pollen mitosis, leading to the formation of the male gametes, the **sperm cells** (Mccormick 1993; Cresti et al. 1992). Microgametogenesis starts with formation of a central vacuole within the uninucleate microspore, pushing the nucleus towards the pollen wall. As long as the nucleus is in a central position within the cytoplasm, the cell is called a **microspore** (Fig. 1 F). With the dislocation of the microspore nucleus the cell becomes the young **pollen grain** (Fig. 1 G).

The **first pollen mitosis** is followed by an asymmetric cell division, leading to the formation of a smaller generative cell and a larger **vegetative cell** with a **vegetative nucleus** (Figs. 4 and 5). Subsequently, the generative cell detaches from the pollen wall and is finally located within the cytoplasm of the vegetative cell (Fig. 1 I). The **generative cell**, sparse in organelles, becomes **spindle-shaped** and the shape of the generative nucleus changes correspondingly (Figs. 6 and 7).

The **second pollen mitosis** includes a symmetric cell division, and divides the generative cell into two **sperm cells** (Figs. 8 and 9), the final stage of gametophytic development (Fig. 1 J-K). Angiosperm pollen is either **two-celled** (75%) or **three-celled** (25% of investigated taxa) at the time of anthesis (Brewbaker 1967; Edlund et al. 2004; Williams et al. 2014). In the latter case the second pollen mitosis takes place in the **pollen tube** (Fig. 1 K), after **germination** of the pollen grain on a stigma or on a corresponding structure (Figs. 10 and 11, Edlund et al. 2004, Mascarenhas 1993). In some families, genera with three- as well as two-celled/nuclear pollen grains occur (e.g., Araceae, Brewbaker 1967).

Microgametogenesis in gymnosperms includes several mitotic divisions. Normally, pollen grains of conifers, cycads and allies are multicelled at anthesis, and comprise prothallial cell(s), a large tube cell and a small antheridial cell. The tube cell becomes a pollen tube; the antheridial cell undergoes division into the stalk cell and the spermatogenous cell, the latter finally dividing into the male gametes (sperm cells or spermatozoids).

**Fig. 1 Pollen development in angiosperms. A**. Schematic illustration of an angiosperm flower. **B**. Cross section of anther. **C**. Pollen mother cell (PMC) encased in callose (diploid nucleus dark red). **D**. Tetrad of four haploid microspores encased in callose (haploid nucleus orange). **E**. Pollen wall formation and separation of microspores. **F**. A single free microspore with central haploid nucleus. **G**. Beginning of gametogenesis, formation of a central vacuole (white). **H**. First pollen mitosis, lens-shaped generative cell with generative nucleus attached to pollen wall. **I**. Two-celled pollen grain, generative cell detached from pollen wall. **J**. Three-celled pollen grain after second pollen mitosis, note two sperm cells with sperm nuclei. **K**. Germination can occur from either a two-celled pollen grain, followed by the formation of sperm cells, or from a three-celled pollen grain (pathways indicated by green arrows)

**A B**

**Fig. 3 Tapetum types. A**. *Hacquetia epipactis*, Apiaceae, secretory tapetum in young anther, Thiéry test. **B**. *Zantedeschia aethiopica*, Araceae, amoeboid tapetum, U+Pb

**Fig. 2 Microsporogenesis. A**. *Scrophularia nodosa*, Scrophulariaceae, tetrad tetrahedral, iodine. **B**. *Spirea* sp., Rosaceae, tetrad tetrahedral, Thiéry test. **C**. *Orobanche hederae*, Orobanchaceae, tetrad planar, potassium iodine

**Fig. 4 Variability of the vegetative nucleus in LM and TEM (cross sections). A-B**. *Dracontium asperum*, Araceae, pollen hydrated in water, vegetative cell with vegetative nucleus (asterisk) and nucleolus (arrowhead). **C**. *Galium odoratum*, Rubiaceae, vegetative cytoplasm and nucleus (asterisk) with nucleolus (arrowhead), U+Pb. **D**. *Salvia nemorosa*, Lamiaceae, vegetative nucleus (asterisk) with nucleolus (arrowhead) surrounded by cytoplasm, U+Pb. **E**. *Thymus glabrescens*, Lamiaceae, vegetative cytoplasm and nucleus (asterisk), modified Thiéry test. **F**. *Thymus glabrescens*, Lamiaceae, vegetative nucleus (asterisk) surrounded by cytoplasm, modified Thiéry test

**Fig. 5 Variability of the vegetative nucleus in TEM (cross sections). A**. *Brassica napus*, Brassicaceae, vegetative cytoplasm and nucleus (asterisk), modified Thiéry test. **B**. *Salvia verticillata*, Lamiaceae, vegetative cytoplasm and nucleus (asterisk), modified Thiéry test. **C.** *Iris pumila*, Iridaceae, vegetative cytoplasm and nucleus (asterisk), modified Thiéry test. **D.** *Consolida regalis*, Ranunculaceae, vegetative nucleus (black asterisk) and generative cell (white asterisk), modified Thiéry test. **E**. *Acinos alpinus*, Lamiaceae, vegetative nucleus (asterisk) with nucleolus (arrowhead) surrounded by cytoplasm, modified Thiéry test. **F**. *Stachys palustris*, Lamiaceae, vegetative nucleus (asterisk) with nucleolus (arrowhead) surrounded by cytoplasm, U+Pb

**Fig. 6 Generative cell and nucleus stained with acetocarmine in LM. A**. *Melampyrum nemorosum*, Orobanchaceae, spindle-shaped generative cell/nucleus (asterisk) and vegetative nucleus. **B**. *Betonica officinalis*, Lamiaceae, spindleshaped generative cell/nucleus (asterisk). **C**. *Anchomanes welwitschii*, Araceae generative cell/nucleus (asterisk). **D**. *Quercus robur*, Fagaceae, generative cell/nucleus (asterisk). **E**. *Carpinus betulus*, Betulaceae, spindle-shaped generative cell/nucleus (asterisk). **F**. *Asterostigma lividum*, Araceae, generative cell/nucleus (asterisk) and vegetative nucleus

**Fig. 7 Variability of the generative cell and nucleus in TEM (cross sections). A**. *Melampyrum nemorosum*, Orobanchaceae, pollen in overview, vegetative nucleus (black asterisk), generative cell/nucleus (white asterisk), modified Thiéry test. **B**. *Melampyrum nemorosum*, Orobanchaceae, vegetative nucleus (black asterisk), generative cell/nucleus (white asterisk), modified Thiéry test. **C**. *Betonica officinalis*, Lamiaceae, vegetative nucleus (black asterisk) and generative cell/nucleus (white asterisk) surrounded by cytoplasm, modified Thiéry test. **D**. *Ajuga reptans*, Lamiaceae, vegetative nucleus (black asterisk) and generative cell/nucleus (white asterisk) surrounded by cytoplasm, modified Thiéry test. **E**. *Acinos alpinus*, Lamiaceae, generative cell/nucleus (white asterisk) surrounded by cytoplasm, modified Thiéry test. **F**. *Stachys palustris*, Lamiaceae, generative cell/nucleus (white asterisk) with nucleolus (arrowhead) surrounded by cytoplasm, modified Thiéry test

30 GENERAL CHAPTERS

**Fig. 8 Sperm cells of different species in LM. A**. *Filarum manserichense*, Araceae, stained pollen showing two sperm cells (white asterisks) and vegetative nucleus (black asterisk), acetocarmine. **B**. *Triticum aestivum*, Poaceae, stained pollen showing two sperm cells (white asterisks) and vegetative nucleus (black asterisk), acetocarmine. **C**. *Ulmus minor*, Ulmaceae, stained pollen showing two sperm cells (white asterisks) and vegetative nucleus (black asterisk), acetocarmine. **D**. *Zea mays*, Poaceae, stained pollen showing two sperm cells, acetocarmine. **E**. *Thymus odoratissimus*, Lamiaceae, stained pollen showing two sperm cells, acetocarmine. **F**. *Amorphophallus taurostigma*, Araceae, pollen showing two sperm cells with nuclei, glycerine

**Fig. 9 Sperm cells in TEM (cross sections). A**. *Hyssopus officinalis*, Lamiaceae, vegetative cytoplasm, vegetative nucleus (black asterisk), two sperm cells/nuclei (white asterisk), modified Thiéry test. **B**. *Galium odoratum*, Rubiaceae, vegetative nucleus (black asterisk) and two sperm cells/nuclei (white asterisk) surrounded by cytoplasm, modified Thiéry test. **C**. *Smyrnium perfoliatum*, Apiaceae, vegetative nucleus (black asterisk) and two sperm cells/nuclei (white asterisk) surrounded by cytoplasm, Thiéry test. **D**. *Jasminum nudiflorum*, Oleaceae, vegetative nucleus (black asterisk) and two sperm cells/nuclei (white asterisk) surrounded by cytoplasm, Lipid-test. **E**. *Zantedeschia aetiopica*, Araceae, vegetative nucleus (black asterisk) and two sperm cells/nuclei (white asterisk) surrounded by cytoplasm; sperm cells still in contact with each other and enclosed by the vegetative nucleus, modified Thiéry test. **F**. *Melampyrum pratense*, Orobanchaceae, vegetative nucleus (black asterisk) and two sperm cells/nuclei (white asterisk) surrounded by cytoplasm, modified Thiéry test

**Fig. 10 Pollen germination and pollen tubes in SEM. A**. *Cryptanthus bromelioides*, Bromeliaceae, sulcate pollen germinating on stigma. **B**. *Prunus* sp., Rosaceae, tricolporate pollen, note germinating pollen on stigma (left side). **C**. *Oxytropis jacquinii*, Fabaceae, tricolporate pollen. **D**. *Tuberaria guttata*, Cistaceae, tricolporate pollen. **E**. *Anthurium gracile*, Araceae, inaperturate pollen. **F**. *Vanilla pompona*, Orchidaceae, porate pollen

**Fig. 11 Pollen germination and pollen tubes in LM and TEM. A-C**. *Arum cylindraceum*, Araceae, three-celled, inaperturate pollen, germination can occur anywhere on the pollen surface, staining with acetocarmine, note the two sperm nuclei (arrowhead) staining dark red with acetocarmine, pictures. **B-C** showing optical section and upper focus. **D**. *Colocasia antiquorum*, Araceae, pollen grains germinating in water. **E-F**. *Smyrnium perfoliatum*, Apiaceae, TEM sections of germinating pollen (arrowheads), Thiéry test; detail of pollen tube with sperm nucleus (**E**, asterisk)

#### **References**


**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

## **Pollen Morphology and Ultrastructure**

**Polarity and Shape – 38 Apertures – 42 Pollen Wall – 45 Harmomegathy: The Harmomegathic Effect – 57 Size – 57 Heterostyly and Pollen Dimorphism – 60 Aberrant Pollen Grains – 60 References – 63**

37

The study of pollen should encompass all structural and ornamental aspects of the grain. Pollen morphology is studied using LM and SEM and is important to visualize the general features of a pollen grain, including, e.g., symmetry, shape, size, aperture number and location, as well as ornamentation. TEM investigations are used to highlight the stratification and the uniqueness of pollen wall layers as well as cytoplasmic features. The following sections explain the most important structural and sculptural pollen features a palynologist should observe.

towards the tetrad center, to the **distal pole** of the microspore/pollen (Fig. 2). The **equatorial plane** is located at the microspore's center, perpendicular to the polar axis (Fig. 2). Therefore, the **equatorial plane** divides the microspore/pollen into a proximal and a distal half, comparable to the northern and southern hemisphere of our planet Earth.

The polarity gives rise to the polar and to the equatorial view. In dicots there is usually one polar

#### **Polarity and Shape**

Mature pollen is shed in **dispersal units**. When the post-meiotic products become separated the dispersal unit is a single pollen grain, a **monad**. Post-meiotic products also become partly separated or remain permanently united, resulting in **dyads** (a rare combination), **tetrads** or **polyads**. **Pollinaria** are dispersal units of two pollinia including a sterile, interconnecting appendage (see "Glossary of Palynological Terms").

Pollen shape and aperture location relate directly to pollen **polarity.** The polarity is determined by the spatial orientation of the microspore in the meiotic tetrad and can be examined in the **tetrad stage** (Fig. 1). The **polar axis** of each microspore/ pollen runs from the **proximal pole,** orientated

**Fig. 1 Tetrad stage**. Orientation of microspores/pollen in the tetrad; distal poles shaded green

**Fig. 2 Polar axis and equatorial plane**. **A-B.** Polar axis and equatorial plane

**Fig. 3 Polarity of pollen in dicots**. **A-B.** *Bellis perennis*, Asteraceae, polar view. **C-D.** *Bellis perennis*, Asteraceae, equatorial view

and one equatorial view (Fig. 3). In monocots, due to the mostly distal position of apertures, there are four views: distal polar, proximal polar, and two different equatorial views (Fig. 4).

**Isopolar** pollen has identical proximal and distal poles, thus the equatorial plane is a symmetry plane. In **heteropolar** pollen the proximal and distal halves differ (Fig. 5).

The various arrangements of the four microspores within **tetrads** depend on the simultaneous or successive type of cytokinesis and on the type of intersporal wall formation. The spatial arrangement of microspores after **simultaneous** cytokinesis is a **tetrahedral** (or rarely decussate) **tetrad** (Fig. 6A). This tetrad types may have systematic relevance, e.g., all species within the genus *Rhododendron* are characterized by tetrahedral tetrads. The spatial arrangement of microspores after **successive** cytokinesis leads to different morphological tetrad types, which can be differentiated into **planar** (tetragonal, linear, T-shaped) and/or **non-planar** (decussate or tetrahedral) tetrads (Fig. 6B). These morphotypes have no systematic relevance, as tetrads may vary within a genus/species, e.g., in *Typha* tetrads may be tetragonal, T-shaped and/or linear (Furness and Rudall 2001; Copenhaver 2005; see also "Illustrated Pollen Terms").

**P/E ratio** (Fig. 7) refers to the length of the polar axis (P) between the two poles compared to the equatorial diameter (E). In **isodiametric** pollen the polar axis is ± equal to the equatorial diameter. In **prolate** pollen the polar axis is longer than the equatorial diameter. In **oblate** pollen the polar axis is shorter than the equatorial diameter. **Pollen shape** refers to the 3-dimensional form of a pollen grain in relation to the P/E ratio. A pollen grain can, for

**Fig. 4 Polarity of pollen in monocots (***Allium paradoxum***, Alliaceae)**. **A-B**. Proximal polar view. **C-D.** Distal polar view. **E-F.** Equatorial view (short axis). **G-H.** Equatorial view (long axis)

**Fig. 5 Pollen symmetry**. **A.** Isopolar pollen. **B.** Heteropolar pollen

**Fig. 6 Pollen arrangements in tetrads**. **A.** Tetrahedral tetrad, *Fagus* sp., Fagaceae, fossil, Quaternary, Austria; apical view. **B.** Planar tetrad, *Typha latifolia*, Typhaceae, fossil, Quaternary, Austria

**Fig. 7 P/E ratio of pollen**. Schematic drawings of oblate (left), isodiametric (middle), and prolate (right) pollen

**Fig. 8 Aperture arrangement**. **A.** Fischer's law, apertures in pairs. **B.** Garside's law, apertures in a group of three

example, be spheroid-, cup-, boat-, cube-, tetrahedral-, triangular dipyramid-, hexafoil dipyramid-, triangular prism-, pentagonal prism-, or hexagonal prism shaped (see "Illustrated Pollen Terms").

In pollen grains with three apertures, two types of aperture arrangement occur after simultaneous cytokinesis (Fig. 8). **Fischer's law** refers to the most frequent arrangement where a pair of apertures occurs at six points in a tetrad (e.g., Ericaceae, permanent tetrads). **Garside's law** refers to the unusual arrangement of apertures where a group of three apertures occur at four points in the tetrad (probably restricted to Proteaceae, no permanent tetrads; Blackmore and Barnes 1995) (Fig. 8).

#### **Apertures**

An **aperture** is a region of the pollen wall that differs significantly from its surroundings in morphology and/or anatomy. The aperture is presumed to function as the site of germination and to play a role in harmomegathy. Pollen grains lacking apertures are called **inaperturate** (Furness 2007). The aperture definition fits both angiosperm and gymnosperm pollen, but in gymnosperms the type of aperture (e.g., leptoma; germination area) usually differs from that in angiosperms.

The polarity of the pollen grain determines the aperture terminology. A circular aperture is termed a **porus** if situated equatorially or globally; if situated distally, it is called an **ulcus.** An elongated aperture is termed a **colpus** if situated equatorially or globally; if situated distally, it is termed a **sulcus**. A combination of porus and colpus is termed a **colporus**; colpori are situated equatorially or globally.

42 GENERAL CHAPTERS

In **heteroaperturate** pollen two different types of apertures (single and/or combined) are present in a combination of colpi with colpori or pori. A circular or elliptic aperture with indistinct margins is termed a **poroid**. Additional rare combinations of ekto- and endoapertures, mostly observed in LM, include pororate and colporoidate (Fig. 9). Pollen grains that have compound apertures composed of circular ektopori and endopori are termed **pororate**. Compound apertures composed of a colpus (ektoaperture) with an indistinct endoaperture are termed **colporoidate**. When the colpus has a clear bulge in the equatorial region of a pollen grain it is termed **geniculum** (Fig. 9D).

The number of equatorial apertures (pori, colpi, colpori) is indicated by the prefixes di-, tri-, tetra-, penta- or hexa-. Writing numbers instead of prefixes is in common use, e.g., 4-porate or tetraporate, 6-colpate or hexacolpate. In this book we prefer the use of prefixes. For pollen grains with more than three apertures, positioned at the equator, the term **stephanoaperturate (stephanoporate, stephanocolpate, stephanocolporate)** is used together with the aperture number (e.g., stephano(4)porate or 4-porate, stephanoporate). Pollen grains with globally distributed apertures are termed **pantoaperturate.**

Apertures are normally covered by an exinous layer, the **aperture membrane**. The aperture membrane can be **ornamented**, e.g., covered with various exine elements, or it is **psilate** (smooth). The aperture can also be covered by an **operculum**, a distinctly delimited exine structure, covering the aperture like a lid (Halbritter and Hesse 1995; Furness and Rudall 2003).

Number, type, and position of apertures are genetically determined and usually the same within

**Fig. 9 Special aperture features observed in LM**. **A-B.** *Corylus* sp., Betulaceae, fossil, Quaternary, Austria, pororate pollen in polar and equatorial view. **C.** *Eucommia* sp., Eucommiaceae, fossil, middle Miocene, Austria, colporoidate pollen in equatorial view. **D.** *Quercus petrea*, Fagaceae, pollen with geniculum (arrowhead), optical section (left) and upper focus (right)

a species, but may also vary (e.g., *Alnus* is usually 5-porate, but number of pori can vary from 3 to 6).

A **pseudocolpus** occurs in heteroaperturate pollen and is presumed to be non-functional. Pseudocolpi mostly alternate with colpori (e.g., in Boraginaceae, Lythraceae) or are flanking each colporus (in Acanthaceae). For examples, see "Illustrated Pollen Tems." Pseudocolpi are believed to play a role in **harmomegathy**, but their effect has been poorly studied.

Pre-(prae-)pollen (Fig. 10) is characterized by proximal and sometimes additional distal apertures, and by presumed proximal germination. Pre-pollen are microspores of certain extinct basal seed plants occurring from the Late Devonian until the Cretaceous. Proximal germination is typical for spores.

Spores germinate at the **tetrad mark** (Fig. 11), the so-called **laesura** (for an extensive overview, see

**Fig. 10 Pre-pollen**. *Nuskoissporites* sp., fossil, Permian, Austria, polar view

Tryon and Lugardon 1991). The tetrad mark is situated at the proximale pole (proximal germination).

**Proximal germination** is a rare exception in seed plants (Fig. 12), e.g., *Beschorneria yuccoides* (Agavaceae) and *Annona muricata* (Annonaceae). In the two cases, this proximally situated aperture (germination area) is functionally replacing the dysfunctional sulcus (Hesse et al. 2009). In *Beschorneria*, pollen grains forming the tetrads are loosely interconnected and separate frequently. In this special case, the sulcus (distal) is not functional, whereas the proximal face, with a highly reduced exine, functions as germination site. In *Annona*, the microspores rotate within the tetrad during development and the original distally placed sulcus becomes proximally positioned (Tsou and Fu 2002).

The aperture usually acts as the (exclusive) **germination** site. In inaperturate angiosperm pollen the pollen tube can protrude at any given site. In taxoid gymnosperm pollen the exine ruptures during hydration at a specialized region, the leptoma, and is subsequently shed (Fig. 13A-B). The protoplast (enclosed by the intine) is released and a pollen tube can be formed anywhere (resembling functionally an inaperturate pollen grain). Furthermore some angiosperm taxa shed the exine before pollen tube formation, e.g., in some Annonaceae, Araceae. Within the Araceae, a shed pollen wall has been observed in several taxa, e.g., *Amorphophallus*, *Taccarum* (Ulrich et al. 2017). The outer pollen wall (composed of polysaccharide) splits immediately in water and sheds soon afterwards. Subsequently, the naked protoplast is

**Fig. 11 Tetrad mark in spores**. **A.** *Polypodium* sp., Polypodiaceae, fossil, monolete tetrad mark, middle Miocene, Austria, polar view. **B.** *Sphagnum* sp. Sphagnaceae, fossil, trilete tetrad mark, middle Miocene, Austria, polar view. **C.** Pteridaceae indet., fossil, middle Miocene, Austria, trilete tetrad mark, polar view. **D.** *Cryptogamma crispa*, Pteridaceae, trilete tetrad mark

**Fig. 12 Proximal germination**. **A.** *Beschorneria yuccoides* (Agavaceae), dry tetrad. **B.** *Beschorneria yuccoides* (Agavaceae), monad, proximal polar view, proximal face functions as germination site. **C.** *Beschorneria yuccoides* (Agavaceae), germinated tetrad, note proximal germination. **D.** *Annona muricata* (Annonaceae), mature tetrads, sulcus hidden in proximal position

floating in water and germinates about 1 hour after shedding (Fig. 13C-D).

During germination, usually a single pollen tube is formed. In some cases, instant pollen **tube-like structures** are simultaneously developed at all apertures (Fig. 14). The formation of these pollen tubelike structures, in relation with moisture, is interpreted as a pre-germinative process that takes place during dehiscence (Blackmore and Cannon 1983).

#### **Pollen Wall**

T he internal construction of the pollen wall is termed **structure**. Ornamenting elements on the pollen surface (ornamentation) are summarized under the term **sculpture** or sculpturing. However, it is not always possible to distinguish between structure and sculpture (e.g., free-standing columellae).

#### **Structure**

In general, the **pollen wall** (**sporoderm**) of seed plants is formed by two main layers: the outer **exine** and the inner **intine** (Fig. 15). The exine consists mainly of **sporopollenin**, which is an acetolysis- and decay-resistant biopolymer. The intine is mainly composed of cellulose and pectin. Commonly, the pollen wall in aperture regions is characterized by the reduction of exine structures or by a deviant exine, and a thick, often bilayered intine.

Two layers within the exine are distinguished: an inner endexine and an outer ektexine. In **tectate** pollen the ektexine usually consists of a basal **foot layer**, an **infratectum** (e.g., columellae) and a **tectum**, the **endexine** is a mainly unstructured layer (Fig. 16A–C). There are many deviations from this principal construction: layers may be thickened, variably structured or lacking. When the pollen

**Fig. 13 Exine/pollen wall shedding**. **A.** *Cephalotaxus* sp., Cephalotaxaceae, fresh pollen in water. **B.** exine (*arrowhead*) shedding prior to pollen tube formation, released protoplast (black asterisk) enclosed by a thick swelled intine (white asterisk). **C.** *Taccarum weddellianum*, Araceae, pollen wall shedding, released protoplast (black asterisk) and shed outer pollen wall (arrowhead). **D.** *Amorphophallus mangelsdorffii*, Araceae, pollen wall shedding, released protoplast (black asterisk) and shed outer pollen wall (arrowhead)

grain is lacking a tectum it is termed **atectate** (Fig. 16D–F). In apertural regions the pollen wall is generally characterized by a different exine construction.

The terms **sexine** for the outer, structured, and **nexine** for the inner, unstructured exine layer are widely used in light microscopy, but do not fully correspond to ekt- and endexine, respectively. When a cavity between the sexine and nexine is present in the interapertural area, this is termed **cavea** (Fig. 17).

#### **Sporopollenin**

John (1814) and Braconnot (1829) introduced the terms "pollenin" and "sporonin" for the resistant exine material of pollen and spores. Zetzsche et al. (1931) then combined the terms into "sporopollenin," that is the major component of the exine found in most pollen and spores, except in filiform seagrass pollen (e.g., Dobritsa et al. 2009; Jardine et al. 2015). Sporopollenin is a complex biopolymer and extremely resistant to

**Fig. 15 Pollen wall stratification**. **A.** Schematic cross section of pollen wall, pk: pollenkitt. **B.** *Ambrosia artemisiifolia*, Asteraceae, optical view showing both intine and exine; acetolyzed. **C.** *Ambrosia artemisiifolia*, Asteraceae, cross section showing both intine (yellow) and exine (blue); modified Thíery-test

**Fig. 16 Tectate vs atectate pollen wall**. **A-C.** Tectate pollen wall. **A.** 3D-model. **B.** *Saxifraga scardica*, Saxifragaceae, cross section showing pollen wall stratification: tectum with internal tectum and supratectal elements, columellate infratectum, very thin footlayer, thin compact-continuous endexine, monolayered intine (colors refer to picture **A**), *pk* pollenkitt. **C.** *Saxifraga scardica*, Saxifragaceae, exine surface in SEM, sculpture striate with nanoechinate suprasculpture (colored). **D-F.** Atectate pollen wall. **D.** 3D-model. **E.** *Iris pumila*, Iridaceae, cross section showing pollen wall stratification: tectum and infratectum lacking, compact-continuous footlayer, monolayered intine (colors refer to picture **D**). **F.** *Iris pumila*, Iridaceae, exine surface (foot layer) in SEM, sculpture verrucate (colored) and clavate (asterisk)

decay as well as to chemical and mechanical damage (e.g., Steemans et al. 2010). However, in the environment, both biotic and abiotic factors are involved in pollen decomposition. Biotic factors are, for instance, the intrusion of bacteria and fungi (e.g., Elsik 1971; Havinga 1971, 1984; Skvarla et al. 1997; Phuphumirat et al. 2011). Abiotic factors include the pH-value of the substrate (e.g., Bryant and Hall 1993),

**Fig. 17 Cavea**. *Xanthium spinosum*, Asteraceae, acetolyzed pollen in polar (left) and equatorial (right) view showing exine cavity (cavea) between sexine and nexine (arrowheads)

oxidation/reduction (e.g., Twiddle and Bunting 2010), autoxidation by UV-light and oxygen (e.g., Jardine et al. 2015), destruction due to mechanic impact, water or fire (Cushing 1967; Bryant et al. 1994; Phuphumirat et al. 2011, 2015), and rapid changes in moisture levels (Halbritter and Hesse 2004).

The preservation status and the amount of pollen and spores in sediments depends on several factors, including rapid anaerobic burial and embedding in mud or peat, absence of any microbial destruction or sapropel, and the exclusion of oxygen (Klaus 1960, 1987; Playford and Dettmann 1996; Traverse 1988, 2007).

Recent studies on the **composition of sporopollenin** suggest that it may have two different types of chemical structures, oxygenated aromatic compounds and aliphatic compounds (e.g., Wiermann et al. 2001; Dobritsa et al. 2009; Gabarayeva and Grigorjeva 2010; Gabarayeva et al. 2010; Steemans et al. 2010; Colpitts et al. 2011). Although its exact structure remains unknown, sporopollenin is believed to compose oxidative polymers of carotenoids, polyunsaturated fatty acids, and conjugated phenols (Diego–Taboada et al. 2014). Some authors are using the plural form "sporopollenins," because there is evidence for several types of sporopollenin in ferns, gymnosperms, and angiosperms (Hemsley et al. 1993; de Leeuw et al. 2006). According to Diego–Taboada et al. (2014) sporopollenin in plants share a common aliphatic core, but depending on the taxon, contain different aromatic side chains. The **chemical constitutional formula** of sporopollenin is also unknown. The **empirical formula** of sporopollenin has highly variable amounts of H- and O-numbers. A generalized formula is C90 H142 O36 (Traverse 1988; Riding and Kyffin– Hughes 2004).

The precise location of **synthesis** of sporopollenin precursors in tapetal cells and the mechanisms of secretion of sporopollenin monomers before polymerization in the microspore walls are still unclear, just as the processes involved in sporopollenin production at the cellular level (Lallemand et al. 2013). Liu and Fan (2013) reviewed the molecular regulation of sporopollenin biosynthesis, which probably includes a framework of catalytic enzyme reactions. As shown in the study by Colpitts et al. (2011), genes responsible for sporopollenin biosynthesis in *Arabidopsis* lead to the conclusion, that the pathway of sporopollenin biosynthesis seems well conserved in land plants since nearly 500 mya.

The question if sporopollenin is of sporophytic or gametophytic origin is still controversial. Probably both sources are involved. Most authors agree that sporopollenin is predominantly produced by the tapetum (Pacini and Franchi 1991; Blackmore et al. 2000; Wallace et al. 2015; Ariizumi and Toryama 2011; Quilichini et al. 2014).

The investigation of fossil pollen and spores revealed that **fossilized sporopollenin** appears chemically very different to sporopollenin found in modern plants (Fraser et al. 2011). During fossilization (coalification) and by diagenetic processes the chemical composition of sporopollenin is modified. Especially at high temperatures, above 200 °C, sporopollenin undergoes a series of chemical changes (Yule et al. 2000; Fraser et al. 2014).

Sporopollenin biochemistry appears to have remained relatively stable since at least the Middle Pennsylvanian (approx. 310 mya). Fraser et al. (2012, 2014) postulated that the structure of sporopollenin has remained constant since plants invaded land during the Middle Ordovician (470-458 mya). A recent comprehensive review on sporopollenin and other biopolymers (de Leeuw et al. 2006) suggests that there may have been multiple forms and configurations of sporopollenin over geological time.

The sporopollenin wall is regarded as a synapomorphy in land plants and allowed land dispersal during the Silurian, perhaps already during the Middle Ordovician (Rubinstein et al. 2010; Wellman 2010).

#### **Chemically Related Biomacromolecules**

Sporopollenin is not unique in pollen/spore walls. Cell walls of some algae and dinoflagellates may contain chemically related biomacromolecules, named **algaenan** and **dinosporin** (Versteegh et al. 2012; Bogus et al. 2012). Like sporopollenin these resistant biomacromolecules may also fossilize. They have been reported in, e.g., *Chlorella* (He et al. 2016), *Spirogyra* (Simons et al. 1983), and *Coleochaete* (Ueno 2009). Furthermore, "sporopollenin-like" biomacromolecules have been found in megaspores and "massulae" of water ferns (Salviniales) (van Bergen et al. 1993), as well as in fruiting bodies of cellular slime molds (Maeda 1984).

**A**

#### **The Angiosperm Pollen Wall**

In angiosperms the **ektexine** consists in general of **tectum**, **infratectum**, and **foot layer**. The outer layer, the more-or-less continuous tectum, can be covered by **supratectal elements**. The infratectum beneath is **columellate** or **granular** (a second layer of columellae may form an internal tectum). However, as, e.g., Doyle (2005) has pointed out intermediate conditions are common. Even the alveolate infratectum, that by definition is restricted to gymnosperms, can also be found in some angiosperms (see "Illustrated Pollen Terms"). The foot layer may be either continuous, discontinuous or absent. The **endexine** can be described as continuous or discontinuous, spongy or compact, overall present, in apertures only, or even completely absent. Some typical deviations of the wall thickness are termed: **arcus**, **annulus**, **tenuitas** (see "Illustrated Pollen Terms") and **costa** (a thickening of the nexine/ endexine bordering an endoaperture; Fig. 18).

#### **The Gymnosperm Pollen Wall**

The gymnosperms comprise cycads, *Ginkgo*, conifers and Gnetales. The basic stratification (ektexine, endexine, and intine) of the gymnosperm pollen wall is identical to that of angiosperms. Still, the gymnosperm pollen wall differs from that of an angiosperm by having (1) a lamellate endexine in mature pollen, and (2) an infratectum that is never columellate (Van Campo and Lugardon 1973). The infratectum is either **alveolate** or **granular**.

A special terminology applies to saccate pollen, i.e. in Pinaceae and Podocarpaceae (Fig. 19). **Saccus** is an exinous expansion forming an air sac, with an alveolate infratectum. **Corpus** is the central body of a saccate pollen grain. **Cappa** is the thick walled proximal face of the corpus. **Leptoma** in conifer pollen refers to a thinning of the pollen wall on the distal face, presumed to function as germination area. Most frequently, two sacci are present (e.g., *Abies*, *Pinus, Picea;* Pinaceae), in some taxa even three (*Dacrycarpus*, *Microstrobus*; Podocarpaceae), or only a single one (*Tsuga*; Pinaceae).

The function and evolutionary significance of saccate pollen have been subject of much confusion. The sacci of Pinaceae and Podocarpaceae are reported to play an aerodynamic role, thus being of adaptive significance for wind pollination (Schwendemann et al. 2007; Grega et al. 2013). In fact, their functional role is to float in a

**Fig. 18 Costa**. **A.** *Nyssa* sp. Nyssaceae, fossil, middle Miocene, Austria, equatorial view (costa highlighted). **B.** *Austobuxus nitidus*, Picrodendraceae, view on the thickening around the endoaperture on the inner side of the wall

liquid pollination droplet towards the ovule ("flotation hypothesis" by Leslie 2010). The flotation system is interpreted as ancestral in conifers. The absence of sacci in, e.g., Cupressaceae and Taxaceae might reflect the loss of "drop mechanism," correlated with the change of pollination mode (shift to upwards orientation of the ovules) (Doyle 2010).

In *Pinus,* pollen can be grouped into two morphotypes (Fig. 20) of systematic value (Grímsson and Zetter 2011). The *Pinus* subgenus *Strobus* (**haploxylon) type** is characterized by pollen grains with broadly attached half-spherical air sacs—in LM the leptoma shows dotted thickenings (seen as dark spots). The *Pinus* subgenus *Pinus* (**diploxylon) type** is characterized by pollen grains with narrowly attached, spherical air sacs often with nodula on nexine area—the leptoma does not show any thickenings.

**Fig. 19 Pollen terminology in saccate gymnosperm pollen**. **A-D.** *Abies* sp., Pinaceae, bisaccate pollen, fossil, Quaternary, Austria, equatorial view. **A.** Corpus highlighted. **B.** Sacci highlighted. **C.** Cappa highlighted. **D.** Leptoma highlighted

#### **Sculpture: Ornamentation**

The terms ornamentation and sculpture applies to surface features of a pollen. The term sculpture is restricted by some authors to surface features in tectate pollen grains (e.g., Praglowski 1975; Punt et al. 2007). **Sculpture elements** (areola, clava, echinus, foveola, fossula, granulum, gemma, plicae, reticulum, rugulae, striae, verruca) can be extremely variable in both size and shape. Based on size many sculpture/ornamentation elements smaller than 1 μm can be described with the prefix micro- (1–0.5 μm) or nano- (0.5–0.1 μm). Also, the boundary between two ornamentation types can be diffuse. For example, "gemmae" and "clavae" are very variable and sometimes hard to differentiate. Combinations of different sculpture/ornamentation elements are common, such as the combination reticulate and foveolate, or echinate and perforate. With a combined sculpture, the pollen ornamentation should then be described in a defined order, with the most eye-catching feature mentioned first, followed by the others. For example, *Aristolochia* pollen is verrucateperforate, as the verrucae are more prominent than the small perforations (Fig. 21). In the Caryophyllaceae, there are numerous, more-orless regularly arranged microechini and perforations. In some taxa the microechini are more prominent (microechinate-perforate), in others the perforations (perforate-microechinate) (Fig. 22). In case none of the features are eyecatching, the dominant feature might be a subjective decision of the palynologist e.g., in taxa, where two features are on a par (microechinate and perforate). A more complex example is *Sanchezia nobilis* (Acanthaceae, Fig. 23): is it plicate and reticulate? Should the rod-like elements

**Fig. 20 Pollen types in saccate Pinus pollen (fossil, middle Miocene, Austria)**. **A.** *Pinus* subgenus *Strobus* (haploxylon), polar view, thickenings (arrowhead). **B.** *Pinus* subgenus *Strobus* (haploxylon), equatorial view. **C.** *Pinus* subgenus *Pinus* (diploxylon), polar view, nodula (arrowhead). **D.** *Pinus* subgenus *Pinus* (diploxylon), equatorial view

**Fig. 21 Combined sculpture elements**. **A-B.** *Aristolochia arborea* (Aristolochiaceae), verrucate, perforate

be termed clavae or free-standing columellae? Is the aperture a porus or a colporus? *PalDat* (www. paldat.org) might provide the answers?

Sculpture/ornamentation elements are often deviating and can be distributed regularly or irregularly over the pollen surface, restricted or absent from distinct areas (polar vs equatorial, interapertural vs aperture area; Fig. 24).

**Ubisch bodies** (orbicules) are sporopollenin elements produced by the tapetum. Ubisch bodies are usually found as isolated particles lining the mature locular wall, or between pollen grains (Huysmans et al. 1998; Halbritter and Hesse 2005; Vinckier et al. 2005; Verstraete et al. 2014). They often resemble the pollen wall ornamentation. In Cupressaceae and Taxaceae, Ubisch bodies are considered part of the pollen ornamentation and are especially frequent on the leptoma of Cupressaceae (for examples, see "Illustrated Pollen Terms").

**Fig. 22 Combined sculpture elements**. **A-B.** *Stellaria media*, Caryophyllaceae, microechinate and perforate. **C-D.** *Saponaria officinalis*, Caryophyllaceae, microechinate and perforate. **E-F.** *Silene succulenta*, Caryophyllaceae, perforate and nanoechinate

**Fig. 23 Interpretation of sculpture elements**. **A-B.** *Sanchezia nobilis*, Acanthaceae, oblique equatorial view and surface detail

#### **Ornamentation in LM vs. SEM**

An accurate description of pollen ornamentation depends on the optical magnification used and particularly on the point resolution. Even the SEM at low resolution may not be sufficient to distinguish pollen grains unequivocally (see "Methods in Palynology"). Depending on the type of microscope used for pollen analysis, some pollen features may remain hidden. For LM studies, the term **scabrate** is used, describing minute sculpture elements of undefined shape and size close to the resolution limit of the LM. For example, *Juglans* pollen is scabrate in LM as well as under low magnification SEM, but is nanoechinate at high resolution SEM (Fig. 25A-B).

The descriptive terms may differ whether LM or SEM is used and should be described for both. For example, *Ulmus* pollen seen in LM is described as **verrucate**. Using low SEM magnification the ornamentation is **rugulate to verrucate** (Fig. 25C-E). High SEM magnification shows additional **granula** (≤0.1 μm).

Another example for different interpretations in LM vs SEM is the term **psilate**. Many pollen grains that appear psilate in LM show a distinct ornamentation using high SEM magnification. For example, pollen of *Allium ursinum* is psilate in LM, but is striate and perforate in SEM (Fig. 25F-G).

Terms with nano- or micro- can only be observed in SEM (see "Methods in Palynology"). For example, the term **granulate** should only be used when describing pollen ornamentation under SEM. When minute sculptural elements are observed under high resolution SEM, it is possible to distinguish real "granula" (sculpture element of different/indefinable shape, ≤ than 0.1 μm) from other nano- and/or micro-sculpture elements. For example, the allegedly granulate ornamentation of many Poaceae is in fact nanoechinate, the pointed ends of the echini are seen best in profile and not from top view (see "Illustrated Pollen Terms").

#### **Role of Pollen Ornamentation in Pollination**

Depending on the pollination mode the outer pollen wall may be either highly ornamented, often with plenty of pollen coatings (mainly pollenkitt; Pacini and Hesse 2005), or with a more or less psilate pollen surface. The pollen wall of zoophilous plants, as well as autogamous plants, is usually highly ornamented and the thick exine consists of high amounts of sporopollenin (Fægri and Iversen 1989). Pollen of anemophilous plants are known to have less ornamentation and less sporopollenin (Friedman and Barrett 2009). Usually psilate pollen in temperate and boreal zones is indicative for anemophily (Fægri and Iversen 1989), whereas in the tropics it is also indicative for zoophily (Furness and Rudall 1999). For example, in Aroideae (e.g., *Montrichardia*, *Dieffenbachia*, *Philodendron*, *Gearum*) psilate pollen usually equipped with pollenkitt is adapted for entomophily (Weber and Halbritter 2007).

#### **Functional Value of Exine Reduction**

Layers of the basic pollen wall type may vary and be partly or totally reduced (for examples, see

**Fig. 24 Pollen surface variation**. **A.** *Fallopia convolvulus,* Polygonaceae, polar view, polar area psilate to perforate and regions around apertures microechinate. **B.** *Sideritis montana,* Lamiaceae, polar view, polar and interapertural areas perforate to foveolate and regions around apertures psilate. **C-D.** *Salvia austriaca,* Lamiaceae, pollen bireticulate, except psilate polar areas (polar and equatorial view). **E-F.** *Solandra longiflora*, Solanaceae, polar area reticulate, equatorial region striato-reticulate (polar and equatorial view)

**Fig. 25 Ornamentation in LM vs SEM**. **A-B**. *Juglans* sp., Juglandaceae. **A**. Scabrate to psilate, LM. **B**. Nanoechinate, SEM. **C-E.** *Ulmus laevis*, Ulmaceae. **C**. Rugulate, LM. **D**. Rugulate to verrucate, low magnification, SEM. **E**. Verrucate, granulate, high magnification SEM. **F-G.** *Allium ursinum*, Amaryllidaceae. **F**. Psilate to scabrate, LM. **G**. Rugulateperforate, low magnification, SEM

"Pollen Wall" in "Illustrated Pollen Terms"). The sporopollenin ektexine is lacking e.g., in some genera of Monimiaceae and Lauraceae (Walker 1976), in the aquatic Ceratophyllaceae (Takahashi 1995), in many genera of Aroideae, and in the inaperturate filiform pollen of seagrasses, *Posidonia*. An absent exine is an adaptation to hydrophily and correlated with, e.g., aquatic habits, anemophily, and pollinia (Furness 2007). Interestingly, exine reduction has evolved iteratively in angiosperms, especially throughout the monocots. Orchidaceae, Asclepiadaceae, Mimosaceae, Annonaceae, and other families often produce compound pollen, where usually only the outermost pollen wall show the typical ektexine structure with tectum and columellae. Pollen grains within calymmate polyads or tetrads have extremely reduced and fragile pollen walls, that probably facilitates pollen germination (Knox and McConchie 1986). The extreme exine reduction in many orchid pollinia seems also to correlate with pollen germination (Johnson and Edwards 2000).

#### **Harmomegathy: The Harmomegathic Effect**

Pollen grains are able to absorb and release water (+ various liquids); thus, each pollen grain exists in two morphologically different conditions, **dry** and **hydrated** (Fig. 26). Harmomegathic mechanisms, e.g., infolding of the pollen wall (Rowley and Skvarla 2000), accommodate the change of the osmotic pressure in the cytoplasm during hydration or dehydration. These mechanisms are denoted as harmomegathic effect, also known as Wodehouse effect. The main purpose of the harmomegathic effect is to protect the male gametophyte against desiccation during pollen presentation and dispersal, and is often related to pollination biology.

In mature anthers, pollen is turgescent before shedding. After anther dehiscence and during pollen presentation, water loss takes place and the pollen grain becomes typically infolded. Various pollen wall features are involved in the harmomegathic effect:


The combination of these features is influencing the mode of infolding. Terms used for common morphotypes of dry pollen include: apertures sunken, boatshaped, cup-shaped, interapertural area infolded, irregularly infolded, not infolded. In addition, the pollen shape can be described with terms that might be helpful for an adequate description such as barrel-like, disk-like, or kidney-like. The mode of infolding and/or shape of pollen in dry condition may be typical for a family and/or genus and therefore of systematic relevance (see "Palynology — History and Systematic Aspects").

The harmomegathic effect is also observed in pollen taken from herbarium material, and to some degree in fossil material (Halbritter and Hesse 2004). This effect is to some degree reversible: rehydrated pollen at the stigma, or under laboratory conditions (various liquids), is again turgescent and largely recalls the shape before shedding. A second dehydration does not necessarily result in the typical dry shape but, if pollen walls are sufficiently stable, the harmomegathic effect can be induced several times in the same way. In pollen with thin walls, the susceptible internal structure may become damaged, and the harmomegathic effect may result in different and randomly shaped pollen. Infoldings of the pollen wall after acetolysis treatment are mostly not comparable with those observed in dry condition.

#### **Size**

Pollen **size** varies from less than 10 <sup>μ</sup>m to more than 100 μm (Fig. 27). To indicate pollen size the largest diameter is used (Hesse et al. 2009). The size depends on the degree of hydration and the preparation method (Reitsma 1969, see also "Methods in Palynology"). Because of this and natural variation, a range categorizing pollen size is recommended: very small (<10 μm), small (10– 25 μm), medium (26–50 μm), large (51–100 μm), and very large (>100 μm).

#### **Heterostyly and Pollen Dimorphism**

I n **heterostylous** (long-styled and short-styled) species two different pollen types occur, where pollen size and number of apertures or the ornamentation may differ. In *Linum flavum* (Linaceae) pollen of the short-styled morph is baculate, and the long-styled morph clavate (Fig. 28). In *Primula veris* (Primulaceae) the pollen of the short-styled morph is larger and has more apertures than pollen of the long-styled morph (Fig. 29A). In the tristylous species *Lythrum salicaria*

**Fig. 26 Harmomegathic effect—hydrated vs dry pollen**. **A-B.** *Cistus creticus*, Cistaceae. **A**. Spheroidal, outline circular. **B**. Prolate, outline lobate, apertures infolded. **C-D.** *Epilobium palustre*, Onagraceae, tetrad. **C**. Oblate, outline triangular. **D**. Interapertural area sunken. **E-F.** *Vriesea pabstii*, Bromeliaceae. **E**. Oblate, outline elliptic. **F**. Boat-shaped. **G-H.** *Alisma lanceolatum*, Alismataceae. **G**. Spheroidal, outline circular. **H**. Irregularly infolded

**Fig. 27 Pollen size categories**.

**Fig. 28 Pollen dimorphism — different ornamentation**. **A-D.** *Linum flavum*, Linaceae, (**A-B**) short-styled morph, baculate, (**B-C**) long-styled morph, clavate

**Fig. 29 Pollen dimorphism — different size. A.** *Primula veris***,** Primulaceae, short-styled morph (left), long-styled morph (right). **B.** *Lythrum salicaria* (Lythraceae), medium-styled morph, dimorphic pollen

**Fig. 30 Pollen dimorphism — different ornamentation**. **A-D.** *Armeria alpina*, Plumbaginaceae. **A-B**. Morph 1, reticulate. **C-D**. Morph 2, reticulate

(Lythraceae) pollen is dimorphic, with different size, ornamentation, and even color of the pollen grains (blue and yellow). Pollen of the long-styled morph is small sized (about 20 μm), short styled morph is medium sized (about 35 μm) and medium-styled morph is small to medium sized (within a single anther) (Fig. 29B). In some Plumbaginaceae, for example in distylous species of *Armeria,* pollen dimorphism (reticulate, different size of lumina and suprasculpture elements) is correlated with dimorphic stigmatic papillae, but style and stamen lengths are monomorphic (Ganders 1979; Fig. 30).

#### **Aberrant Pollen Grains**

Aberrant pollen grains are often ignored but occur regularly in small percentages in nearly all anthers and may vary from one individual to another (Pozhidaev 2000a, b; Banks et al. 2007). These aberrant pollen grains can differ from the typical pollen type of the species in shape and dimension, in number and arrangement of apertures, and in ornamentation type (Fig. 31). Reasons for the production of deviating pollen forms are genetically (polyploidy),

**Fig. 31 Aberrant pollen grains**. **A.** *Malus sieboldii*, Rosaceae, irregular aperture arrangement (usually tricolporate). **B.** *Oxalis* sp., Oxalidaceae, many aborted pollen grains, giant pollen (usually tricolpate). **C.** *Scaevola* sp., Goodeniaceae, pollen varies in size and aperture arrangement. **D.** *Scandix pecten-veneris*, Apiaceae, "double" pollen grain (usually tricolporate). **E.** *Codiaeum*-hybrid, Euphorbiaceae, ornamentation intermediate between parent plant species e.g., croton pattern and reticulate with free-standing columellae. **F.** *Codiaeum*-hybrid, Euphorbiaceae, surface detail

chemically, or environmentally induced. Such deviating, malformed pollen is frequently found in cultivated plants, ornamental plants, agricultural crops, annual plants, plants with asexual reproduction (autogamic plants, apomicts), and hybrids. Some species of apomicts, agricultural crops or cultivated plants (e.g., *Malus sieboldii*) produce only malformed pollen.

#### **References**


is a Long–Chain Fatty Acid v–Hydroxylase essential for sporopollenin synthesis in pollen of Arabidopsis. Plant Physiol 151: 574–589


**Open Access** This Chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

### **Misinterpretations in Palynology**

**Example 1: Tripartite Feature in Gymnosperms — Impression Mark – 68**

**Example 2: Tripartite Feature in Angiosperms — Triangular Tenuitas – 68**

**Example 3: Tripartite Feature in Angiosperms — Synaperture – 68**

**Example 4: Tripartite Feature in Angiosperms — Trichotomosulcus – 70**

**Example 5: Tripartite Feature in Angiosperms — Sulci vs. Colpi vs. Tenuitas – 70**

**Example 6: Tripartite Feature in Angiosperms — Triradiate Aperture – 70**

**Example 7: Apertures in Angiosperms — Planaperturate – 70**

**Example 8: Apertures in Angiosperms — Inconspicuous Pori – 72**

**Example 9: Apertures in Angiosperms — Inconspicuous Colpi – 72**

**Example 10: Apertures in Angiosperms — Hidden Apertures – 73**

**Example 11: Apertures in Angiosperms — Ring-like Apertures vs. Colpate-Operculate – 73**

**Example 12: Apertures in Angiosperms — Tenuitas vs. Poroid – 73** **Example 13: Apertures in Angiosperms — Infoldings vs. Apertures – 73**

**Example 14: Apertures in Angiosperms — Ulcerate-Operculate vs. Ring-like Aperture – 74**

**Example 15: Apertures in Angiosperms — Disulcate vs. Dicolpate – 74**

**Example 16: Apertures in Angiosperms — Zon-, Zono-, Zoni-, Zona- vs. Ring-like Aperture and Stephanoaperturate Pollen – 75**

**Example 17: Magnification Effect — Retipilate vs. Reticulum Cristatum – 76**

**Example 18: Dispersal Units — Massula vs. Polyad – 76**

**Example 19: Preparation Effect — Psilate vs. Ornamented – 77**

**Example 20: Preparation Effect — Areolate-Fossulate vs. Verrucate – 78**

**Example 21: Preparation Effect — Striate vs. Striato-Reticulate – 78**

**Example 22: Staining Methods — Absence or Presence of Endexine – 80**

67

**References – 84**

T he description of pollen ornamentation depends on three major parameters (1) the interpretations of the palynologist (which are subjective), (2) the pollen terminology applied, and (3) the magnification, resolution, and methods used.

The application of different preparation and staining methods and a combined analysis with light microscopy, scanning- and transmission electron microscopy are essential for the interpretation of pollen characters. Investigation of recent and fossil pollen material often reveals interesting features that in some cases may be misinterpreted. To demonstrate the wide range of possible misinterpretations, the following examples are given:

#### **Example 1: Tripartite Feature in Gymnosperms — Impression Mark**

Mature pollen of conifers, such as *Abies*, *Larix,* and *Pseudotsuga,* often shows proximally a Y-shaped bulge on the proximal polar side, comparable to a tetrad mark, which is called an **impression mark** (Fig. 1; Harley 1999). The mark results from the close proximity of the four pollen grains at the post-meiotic tetrad phase and is retained afterwards and is not a germination feature. Impression marks are also found in palm pollen. Note: the term tetrad mark is restricted to spores, where it is a germination feature.

#### **Example 2: Tripartite Feature in Angiosperms — Triangular Tenuitas**

S uperficially similar features in angiosperms are not comparable to those observed in gymnosperms. In recent and fossil Sapindaceae a three-armed feature (more precisely a triangle) is found. *Cardiospermum* has a narrow **triangular tenuitas** (thinning) at the proximal pole, whereas other recent and subfossil Sapindaceae show such a feature at both poles (Fig. 2).

#### **Example 3: Tripartite Feature in Angiosperms — Synaperture**

T riangular pollen as found in Myrtaceae, some Primulaceae (*Primula farinosa* or *P. denticulata*) and Loranthaceae is characterized by a tripartite feature in both polar areas (Fig. 3). These are in fact

**Fig. 1 Impression mark**. **A.** *Abies cephalonica*, Pinaceae, proximal polar view, indistinct impression mark. **B-C.** *Larix* sp., Pinaceae, fossil, middle Miocene, Austria, proximal polar view, Y-shaped impression mark in SEM (**B**) and LM (**C**)

**Fig. 2 Triangular tenuitas**. **A-B.** *Cardiospermum corindum,* Sapindaceae, tricolporate, equatorial view (**A**), proximal pole with triangular thinning area (**B**)

**Fig. 3 Synaperturate pollen**. **A-B.** *Melaleuca armillaris*, Myrthaceae, syncolporate, polar view (**A**), close-up of polar area (**B**). **C.** *Primula denticulata*, Primulaceae, syncolpate, polar view. **D.** *Primula farinosa*, Primulaceae, syncolpate, dry pollen

three colpi, extending towards and merging at the poles. The pollen is therefore synaperturate (syncolpate, syncolporate). In for example, *Primula* the colpi dissect in the polar area, leaving a triangular field at both poles.

#### **Example 4: Tripartite Feature in Angiosperms — Trichotomosulcus**

Another tripartite feature is the **trichotomosulcus** (Harley 2004), a three-armed sulcus occurring exclusively distally, as, e.g., in *Dianella*. Trichotomosulcate pollen has been discussed in relation to the evolution of the tricolpate dicot condition, but so far without success (Fig. 4).

**Fig. 4 Trichotomosulcus**. **A-B.** *Dianella tasmanica*, Phormiaceae, trichotomosulcus (**A**), dry pollen, aperture infolded (**B**)

#### **Example 5: Tripartite Feature in Angiosperms — Sulci vs. Colpi vs. Tenuitas**

T he angiosperm-like pollen of the fossil genus *Eucommiidites* is "trisulcate": a broad distal sulcus and two narrower additional "sulci" (at angles of c. 120° seen from the main sulcus; Fig. 5). This feature was erroneously interpreted as tricolpate pollen (with colpi equatorially situated).

A similar arrangement of a distal sulcus and two small additional sulci on the proximal face was described, for example, in some species of *Tulipa* (Liliaceae) and *Tinantia* (formerly *Commelinantia*, Commelinaceae), but these cases were never interpreted as equivalent to a tricolpate condition (Harley 2004) (Fig. 6). The two small additional sulci may also be interpreted as tenuitates. In some cases the three "sulci" are of similar size. The aperture condition is very similar to a tricolpate one.

#### **Example 6: Tripartite Feature in Angiosperms — Triradiate Aperture**

Another three-armed feature is the triradiate aperture in *Thesium alpinum* (Santalaceae) pollen. The heteropolar pollen is 3-aperturate, with apertures placed in the three tapered edges of a tetrahedron (Feuer 1977). Each aperture has a very inconspicuous triradiate outline, which is situated equatorially. Two of the arms point towards the neighboring tetrahedron edge and are rather short; the third, elongated arm is directed towards the rounded pole (Fig. 7).

#### **Example 7: Apertures in Angiosperms — Planaperturate**

S ometimes apertures are inconspicuous and not discernible at first sight. In pollen of *Pachira aquatica* (Malvaceae) three large, more-or-less hemispherical areas are seen equatorially, which may at first sight be interpreted as pores. However, a detailed observation reveals **planaperturate** pollen grains with three short colpi (Fig. 8).

**Fig. 5 Trisulcate pollen**. **A-C.** *Eucommiidites* sp., fossil pollen, Lower Cretaceous of U.S.A., main sulcus with membrane seen in center of pollen grain, flanked by additional narrow sulci on each side (at angles of c. 120°, **A-B**), close-up showing sulcus membrane of main sulcus (**C**). **D-F.** *Eucommiidites* sp., fossil pollen, Lower Cretaceous of U.S.A., Narrow lateral sulcus (**E**), same grain turned showing the main broad sulcus and one narrow lateral sulcus (**F**)

**Fig. 6 Trisulcate pollen**. *Tulipa kaufmanniana*, Liliaceae, trisulcate or sulcate with two tenuitates, equatorial view

**Fig. 7 Triradiate aperture**. **A-D.** *Thesium alpinum*, Santalaceae. **A.** Tricolpate, heteropolar, triradiate colpus. **B.** Polar view (flattened pole). **C.** Equatorial view. **D.** Polar view (rounded pole)

#### **Example 8: Apertures in Angiosperms — Inconspicuous Pori**

I n *Calliandra emarginata* (Mimosaceae) the monads forming a polyad are separated by narrow groove-like depressions. At low magnification the presence and localization of the apertures remain indistinct; high SEM magnification reveals that the apertures are very inconspicuous pores, situated equatorially, usually at the conjunction of three or four monads (Fig. 9 A, B).

Also, the aperture condition may be overlooked due to other eye-catching features. The clypeate pollen of *Phyllanthus* x *elongatus* (Euphorbiaceae) seems to be inaperturate. Only close-ups reveal the inconspicuous few pores between the exine shields (Fig. 9 C, D).

#### **Example 9: Apertures in Angiosperms — Inconspicuous Colpi**

T he disc-like pollen of *Oryctanthus* sp. (Loranthaceae) shows at both poles conspicuous circular depressions that are not apertures (Feuer and Kuijt 1985; Grímsson et al. 2018). The pollen is according to Grímsson et al. (2018) demi(3)colpate, with

inconspicuous slit-like colpi positioned between the polar depressions (Fig. 10). Another example are some Asteraceae pollen, where the colpi are often inconspicuous or not visible in SEM, but obvious in LM.

#### **Example 10: Apertures in Angiosperms — Hidden Apertures**

Recent and fossil triaperturate (colpate or porate) pollen of *Trapa* (Trapaceae) is distinguished by unique meridional exine ridges (crests) covering the apertures (Zetter and Ferguson 2001) (Fig. 11).

#### **Example 11: Apertures in Angiosperms — Ring-like Apertures vs. Colpate-Operculate**

T he apertures in *Passiflora* cf. *incarnata* may be interpreted as three ring-like apertures or may be interpreted as pori (or colpi) each with an operculum. In other species of *Passiflora* e.g., *P. citrina* and *P. suberosa*, the apertures are both narrower and stephanocolpate (Fig. 12).

#### **Example 12: Apertures in Angiosperms — Tenuitas vs. Poroid**

T **enuitas** is a general term for a pollen wall thinning (Kremp 1968; Harley 2004; Punt et al. 2007). It is normally found additional to apertures, e.g., in *Myosotis* (Fig. 13). A circular tenuitas can be mistaken for a **poroid**, which is a circular or elliptic aperture with an indistinct margin (see also "Illustrated Pollen Terms").

#### **Example 13: Apertures in Angiosperms — Infoldings vs. Apertures**

When pollen is infolded it can be hard to distinguish the apertures. Pollen of *Sparganium erectum* (Sparganiaceae) is in dry stage infolded, boat-shaped, and would be considered as sulcate. In fact, *Sparganium* pollen is ulcerate, the ulcus is seen clearly in the hydrated, spherical pollen stage (Fig. 14).

**A**

**B**

**Fig. 8 Planaperturate pollen**. **A-C.** *Pachira aquatica*, Malvaceae, polar view (**A**), equatorial view (**B**), oblique equatorial view (**C**)

**Fig. 9 Apertures in** *Calliandra* **and** *Phyllanthus*. **A-B.** *Calliandra tergemina*, Fabaceae, polyad, dry state (**A**). Apertures (pori) at the junction of four monads (**B**). **C-D.** *Phyllanthus x elongatus*, Euphorbiaceae, clypeate, seemingly inaperturate (**C**), Inconspicuous pores (colored) between the exine shields (**D**)

#### **Example 14: Apertures in Angiosperms — Ulcerate-Operculate vs. Ring-like Aperture**

N*ymphaea alba* (Nymphaeaceae) pollen has asymmetrical halves divided by a ring-like aperture (Fig. 15). The features of the smaller distal half may be misinterpreted as a large ulcus with a conspicuous operculum. Ultrastructural studies and germination experiments support the interpretation of a ring-like aperture (Gabarayeva and Rowley 1994; Hesse and Zetter 2005).

#### **Example 15: Apertures in Angiosperms — Disulcate vs. Dicolpate**

T he term disulcate defines two elongated apertures situated usually distally (but not directly at the distal pole), running parallel to or even in the equator (Fig. 16). If the apertures are running meridionally, pollen would be dicolpate (Halbritter and Hesse 1993). To distinguish if the pollen is disulcate or dicolpate it is important to study the pollen in tetrad arrangement to clarify the polarity and position of apertures (see Fig. 3 in "Methods in Palynology").

**Fig. 10 Apertures in** *Oryctanthus*. **A-C.** *Oryctanthus alveolatus*, Loranthaceae, acetolyzed pollen, arrowheads point to colpi, LM (**A**). two grains in polar view, SEM (**B**). close-up showing colpus (**C**). **D-E.** *Carthamus lanatus*, Asteraceae, hydrated pollen, pollen in SEM seem porate (**D**). Acetolyzed pollen, colporus (highlighted) only visible in LM (**E**)

Examples for taxa with disulcate pollen are the monocots *Tofieldia calyculata* with one sulcus distally, the other proximally, *Uvularia grandiflora*, *Eichhornia crassipes* (Hesse et al. 2009), some *Dioscorea* species (Schols et al. 2005), *Pontederia cordata* (Halbritter 2016), *Calla palustris* (Ulrich et al. 2013), and the magnoliid *Calycanthus floridus* (Huynh 1976).

#### **Example 16: Apertures in Angiosperms — Zon-, Zono-, Zoni-, Zona- vs. Ring-like Aperture and Stephanoaperturate Pollen**

T erms combining the basic prefix zon- together with its linguistic derivatives are a source of endless confusion, misunderstanding and superfluous inflation of terms. The prefix include **zon-** (in zonorate, for a ring-like endoaperture, the os, at the equator), the outdated, rarely used **zoni-** (however, with two quite different terminological applications), but especially **zona-** (indicating exclusively a ring-like feature situated anywhere) and **zono-** (indicating any feature located strictly equatorially).

Terms for ring-like (aperture) features include zona**-**aperturate, zona**-**sulculus (addressing the polarity by anazona-sulculus and catazonasulculus), zona**-**sulcus, zonate, zono**-**aperturate, and also related names (e.g., "fully zonate condition" sensu Grayum 1992). Even the misleading and contradictory **zono**-sulcus (a sulcus cannot be situated equatorially) is used instead of the correct, but phonetically confusable, **zona**-sulcus. Even the

**B C**

**Fig. 11 Apertures in** *Trapa*. **A-C.** *Trapa* sp., Trapaceae, fossil, late Miocene, Austria, equatorial view, crest broken, LM (**A**). Equatorial view, crest partly broken, colpus visible in SEM (**B**). Equatorial view, SEM (**C**)

trained palynologist may become confused. Therefore, all these terms should be avoided and we recommend the following two terms: **ring-like aperture and stephanoaperturate** (see "Illustrated Pollen Terms"). Any encircling aperture ("zonaaperturate"), irrespective of meridional or equatorial location, is simply called a **ring-like aperture**. Any case with more than three apertures at the equator ("zono-aperturate") is called **stephanoaperturate**.

#### **Example 17: Magnification Effect — Retipilate vs. Reticulum Cristatum**

T he term retipilate (reticuloid) describes a reticulum formed by pila instead of muri (Erdtman 1952). Combined investigations based on LM and SEM have revealed that the given examples *Callitriche* (Punt et al. 2007) and *Cuscuta lupuliformis* (Erdtman 1952) do not fit the definition of retipilate. In fact, the reticulum consists of muri with prominent suprasculpture elements and are without isolated pilae. Such ornamentation is termed reticulum cristatum (a special type of reticulum; muri with prominent suprasculpture elements; Fig. 17, see also "Illustrated Pollen Terms"). So far no example for retipilate sensu Erdtman (1952) is currently known.

#### **Example 18: Dispersal Units — Massula vs. Polyad**

F or a pollen dispersal unit of more than four pollen grains two terms are in use, **massula** and **polyad** (Fig. 18). The application of both terms is confusing and inconsistent in the literature. Often, the various authors employ the terms more or less interchangeably and do not provide a sharp delimitation (Walker 1971; Wagenitz 2003; Punt et al. 2007; Traverse 2007). These terms, however, are not exchangeable for historical and practical reasons (see extensive review by Teppner 2007).

The term massula was coined by Richard (1817) for parts of a pollinium in some Orchidaceae and should be used for the subunits of orchid sectile pollinia/pollinaria. Massulae within one and the same pollinium are variable and different in shape, size, and numbers of pollen grains. Unfortunately, the term massula has also been used to designate compound pollen in various other families, e.g.

**Fig. 12 Apertures in** *Passiflora*. **A-B.** *Passiflora* cf. *incarnata*, Passifloraceae; colpate, operculate aperture, polar view (**A**), equatorial view (**B**). **C.** *Passiflora citrina*, Passifloraceae, stephanocolpate, operculate, polar view. **D.** *Passiflora suberosa,* Passifloraceae, stephanocolpate, operculate, dry pollen

Fabaceae-Mimosoideae, producing dispersal units of more than four pollen grains (e.g., Wettstein 1907; Wagenitz 2003; Punt et al. 2007). For these the term polyad — coined by Iversen and Troels-Smith (1950) — should be used, denoting a symmetric dispersal unit of more than four regularly arranged and permanently united pollen grains. Polyads, currently known to occur in Fabaceae (Mimosoideae), Gentianaceae, Hippocrateaceae, Celastraceae and Annonaceae, contain a specific number of pollen grains (a multiple of four: 8, 12, 16, 24, 32, 48, 64) and show a species-specific shape.

#### **Example 19: Preparation Effect — Psilate vs. Ornamented**

Ornamentation sometimes depends on the **preparation method**. A striking example is pollen of many Aroideae (Araceae), that are ornamented (e.g., echinate, striate, verrucate) in fresh or dry condition, but become psilate following acetolysis (Fig. 19). The outer pollen wall layer and ornamentation elements are composed of polysaccharide (lack sporopollenin) and are therefore destroyed during acetolysis (Weber et al. 1999; Ulrich et al. 2017).

**Fig. 13 Tenuitas vs. poroid**. **A-B.** *Myosotis palustris*, Boraginaceae, equatorial view, heteroaperturate, alternating colpori and pseudocolpi (**A**), polar view, polar area with triangular tenuitas (**B**)

#### **Example 20: Preparation Effect — Areolate-Fossulate vs. Verrucate**

T he dehydration process with 2,2-dimethoxypropane (DMP) and critical point drying (CPD) for SEM investigations can affect the ornamentation. An example for different interpretations in relation to a varying degree of hydration is *Trichosanthes anguina* (Cucurbitaceae), where the ornamentation can reflect different degrees of hydration. The ornamentation can be described as areolate and fossulate in partially hydrated condition or verrucate and perforate in fully hydrated condition (Fig. 20).

**Fig. 14 Apertures in** *Sparganium*. **A-B.** *Sparganium erectum*, Sparganiaceae, ulcerate, equatorial view hydrated pollen (**A**), boat-shaped, dry pollen (**B**)

#### **Example 21: Preparation Effect — Striate vs. Striato-reticulate**

T he ornamentation of *Amorphophallus longituberosus* pollen in dry condition or hydrated in water is striate, but after critical point drying it becomes striate to reticulate. The striate to reticulate ornamentation of *Amorphophallus longituberosus* is a result of an expanding thin surface layer (Fig. 21 D). During rehydration, the expansion of the thin layer itself forms a reticulum (Fig. 21 C), which finally ruptures partly or completely (Ulrich et al. 2017).

**Fig. 15 Apertures in Nymphaea**. **A-C.** *Nymphaea* sp., Nymphaeaceae; ring-like aperture, polar view (**A**), Ringlike aperture, equatorial view (**B**), dry pollen, cup-shaped (**C**)

**Fig. 16 Disulcate**. **A-B.** *Calla palustris*, Araceae, polar and equatorial view

**Fig. 17 Retipilate vs. reticulum cristatum**. **A.** Drawing from Erdtman (1952). **B.** Drawings from Punt et al. (2007). **C.** *Callitriche palustris*, Plantaginaceae, acetolyzed pollen in LM. **D.** *Callitriche polymorpha*, Plantaginaceae, reticulum cristatum with small gemmae (suprasculpture) on thin muri. **E.** *Cuscuta lupuliformis*, Convolvulaceae, reticulum cristatum with nanoechini (suprasculpture)

#### **Example 22: Staining Methods — Absence or Presence of Endexine**

T he staining behavior of the endexine is very heterogeneous, even within the same plant family or the same genus (Weber and Ulrich 2010). Therefore, the endexine is often reported as absent even though the layer is actually present. In most studies on pollen ultrastructure, sections are stained with uranyl acetate and lead citrate only. To truly distinguish the presence of endexine one should/must apply potassium permanganate which stains the endexine electron dense (Fig. 22, see also "Methods in Palynology").

Misinterpretations in Palynology

**Fig. 18 Massula vs. polyad**. **A.** *Habenaria* sp., Orchidaceae, pollinium composed of numerous massulae (massula highlighted). **B.** *Orchis ustulata*, Orchidaceae, pollinium composed of numerous massulae, two massulae partly segregated (massula highlighted). **C.** *Ludisia discolor*, Orchidaceae, 2 segregated massulae. **D.** *Albizia julibrissin*, Fabaceae, polyad (monad highlighted)

**Fig. 19 Preparation effect — psilate vs. ornamented**. **A-C.** *Amorphophallus krausei*, Araceae, pollen striate in hydrated condition (**A**), psilate after acetolysis, LM (**B**) **Fig. 20 Preparation effect on ornamentation**. **A-C.** *Trichosanthes anguina*, Cucurbitaceae. **A.** Pollen at different state of hydration: fully hydrated (left), less hydrated (right). **B.** Hydrated pollen, surface detail, verrucate, perforate. **C.** Less hydrated, surface detail, areolate-fossulate

and SEM (**C**)

**Fig. 21 Preparation effect on ornamentation**. **A-D.** *Amorphophallus longituberosus*, Araceae, hydrated pollen in water with striate ornamentation, LM (**A**), dry pollen in SEM, striate (**B**), hydrated pollen in SEM, striate to reticulate (**C**), hydrated pollen in SEM, ornamentation striate with expanding thin surface layer (**D**)

**Fig. 22 Absence or presence of endexine**. **A-B.** *Thymus odoratissimus*, Lamiaceae, U + Pb staining, endexine (arrowhead) not clearly visible (**A**), potassium permanganate staining, endexine (arrowhead) clearly visible (**B**)

#### **References**


**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

84 GENERAL CHAPTERS

## **How to Describe and Illustrate Pollen Grains**

**Pollen from a Single Extant Taxon: Online Publication in** *PalDat* **– 86**

**Groups of Extant Pollen – 86**

**Fossil Pollen – 89**

**References – 95**

85

For the description of a pollen grain, a number of features are used including size, polarity and shape, aperture condition, ornamentation, and pollen wall structure. Additional and often more specialized features depend on the group of plants under study, Gymnosperms (Cycadales, Ginkgoales, Pinales, Gnetales) vs. Angiosperms (magnoliids, monocots, commelinids, eudicots). These features can only be obtained by the application of a combined analysis with LM, SEM, and TEM (Fig. 1). In order to compare and categorize pollen, a common language and understanding of technical terms is necessary.

The description and illustration of a pollen grain depends on how the material is going to be presented and if one is describing a single fossil pollen grain, pollen of a particular extant species, pollen representing several species, a whole genus, several related genera, a complete family, or even a number of families. For future work it is important to provide both LM and SEM micrographs (even TEM), including incorporated scale bars, showing each taxon and close-ups of what are considered diagnostic features of pollen. When documenting the sculpture of pollen grains in SEM it has to be made sure that the magnification is high enough to distinguish the shape and outline of sculpture elements larger than 0.1 μm in diameter. LM- and SEMdiagnosis may be different from each other, due to the methods and techniques used. The methods used to prepare pollen grains for LM, SEM, and TEM must be mentioned along with the pollen descriptions, preferably in a material and method section.

#### **Pollen from a Single Extant Taxon: Online Publication in** *PalDat*

Pollen grains from single extant species have rarely been accepted by scientific journals. There is now a new online venue *PalDat,* for publishing pollen from a single species. *PalDat* is the world's most comprehensive pollen database (www. paldat.org) and contains tools for pollen identification as well as global, free online submission and publication with review and editorial process (Weber and Ulrich 2017). *PalDat* already provides a large amount of pollen data on a variety of plant families. Each taxon entry (online publication) ideally includes a detailed description and micrographs (LM, SEM, and TEM) of the pollen, as well as images of the plant/

inflorescence/flower and information on relevant literature (Fig. 2). *PalDat* is freely accessible and following a free registration it is open for contributions from all those willing to publish their pollen descriptions and micrographs online. Registered authors may also contribute as co-authors to existing publications by submitting new images and/or new data to pollen diagnosis (with review and editorial process). All changes are recorded in the database history as links to previous versions of the publication. Each contribution is citable and accessible for all users. Registered users can download publications in pdf form. The terminology used in *PalDat* follows this book.

#### **Groups of Extant Pollen**

Many of the classical papers on pollen morphology and ultrastructure, covering a large number of extant taxa, provide only a general description of pollen types with pollen of different species lumped together. Furthermore, micrographs are showing selected taxa and usually not the same taxon photographed in both LM and SEM. This makes the data unreliable and not very useful for among others paleopalynologists that want to compare their fossil pollen grains very precisely to particular extant taxa. The decision on particular potential modern analogues of the fossil pollen grain can have major effects on the paleovegetation reconstruction and paleoecological and paleoclimate interpretations of the fossil assemblage, as well as on the paleophytogeographic signal of the taxon. It is recommended, disregardless of the description, that all species be fully illustrated by LM and SEM (and TEM when possible) and their basic and diagnostic morphological features compiled in a table so they can be easily compared (Table 1). The example shown here are Winteraceae pollen tetrads. When portraying tetrads it is useful to show their basal-, lateral-, as well as apical view, in both LM (Fig. 3) and SEM (Fig. 4). Pollen grains should be portrayed in polar and equatorial view. Illustrating pollen from different taxa together on a plate/figure with the same magnification makes it easier to realize size differences. The SEM close-ups are then used to highlight the main sculpture features or the dissimilarities of the taxa. Ideally all close-ups showing sculpture elements should have the same magnification for an easy comparison (Fig. 5).

**Fig. 1 Diagnosis worksheet**. *PalDat* worksheet with all pollen features obtained by a combined analysis using LM, SEM, and TEM. Blue dots indicate LM-, yellow dots SEM-, and red dots TEM-based analyses. *PalDat* pictures showing *Plantago maritima*

mitochondria

starch,lipids

**Fig. 2 Online publication in** *PalDat*. Screenshot showing part of the online publication of *Betula pendula* (Halbritter and Diethart 2016)


Main features of three different Winteraceae pollen tetrads

*Annotation: Measurements like exine, nexine, and sexine thickness provided in Table 1 (asterisks) are commonly used in (paleo) palynological literature. Scientists should be aware that such measurements (e.g., 0.7 or 0.9 μm) vary highly, up to 30%, depending on the methods and tools used. Therefore, the measurements should not be overrated or used for taxonomic discrimination.*

#### **Fossil Pollen**

F rom the birth of paleopalynology this branch of science has been plagued by the lack of taxonomic foundation when interpreting paleoenvironments. It is very unfortunate that numerous new "scientific" publications dealing with the subjects of paleoecology, paleovegetation, paleoclimate and various aspects of paleophytogeography still present only a list of taxa observed in LM. Some publications include LM micrographs of the most "common" taxa, but only in exceptional cases the LM micrographs are accompanied by SEM micrographs. The absence of illustrations makes it impossible for any reader to verify, or later revise, the taxonomic background and to conclude if the modern living relative or potential modern analogue of the fossil taxon is justified. Every proper scientific journal should make it a mandatory request that all pollen types are represented by at least one LM micrograph. Furthermore, all taxa that suggest some sort of different, abnormal or exceptional paleo-parameters, in an otherwise "homogeneous" assemblage, or taxa that are used to set any sort of boundaries (temperature, precipitation, biozone, time, etc.), should be illustrated using both LM and SEM (in some cases even TEM). These contrasting taxa might include a dry element in an otherwise humid assemblage, a tropical element in an otherwise temperate assemblage, or an African element in an otherwise North American-Eurasian assemblage. Even though the journal would not allow these illustrations in the printed version most of them now offer the possibility to archive online supplementary files where the pollen can be illustrated.

For those who want to produce a taxonomically valid study based on fossil material are advised to use the single-grain method when investigating fossil pollen and make sure not to sieve the sample

**Fig. 3 LM micrographs of Winteraceae pollen tetrads**. Tetrads shown in basal-(left), lateral-(middle), and apical (right) view at high focus (upper three rows) and in optical cross section (lower three rows). *Takhtajania perrieri* (first and fourth row), *Exospermum stipitatum* (second and fifth row), *Tasmannia insipida* (third and sixth row)

**Fig. 4 SEM micrographs of Winteraceae pollen tetrads**. Tetrads shown in basal view (upper row), lateral view (middle row) and apical view (lower row). *Takhtajania perrieri* (left), *Exospermum stipitatum* (middle), *Tasmannia insipida* (right)

**Fig. 5 Details of Winteraceae pollen tetrads**. SEM close-ups of *Takhtajania perrieri* (**A-B**), *Exospermum stipitatum* (**C-D**) and *Tasmannia insipida* (**E-F**), showing sculpture on distal face of pollen (**A, C, E**) and the aperture region and ulcus membrane (**B, D, F**)

during preparation (see "Methods in Palynology"). This allows the researcher to study all elements occurring within a sample using both LM and SEM and to investigate even very small and/or rare pollen grains. The small and/or rare pollen (Fig. 6) would otherwise be overlooked during the old-fashion routine LM observation, where the researcher usually counts 300–600 grains. When illustrating fossil pollen it is important to show the grain in both LM and SEM. Close-ups taken with the SEM should have magnification high enough so all sculpture elements larger than 0.1 μm become distinguishable. Sculpture and suprasculpture elements smaller than 1 μm are not observed or hard to distinguish using LM only, but will be revealed using high magnification SEM (Fig. 7). Many pollen grains that look similar or the same in LM can be distinguished using SEM. In some cases it is beneficial to turn the pollen grain once it has been photographed in SEM, re-sputter and photograph again. This applies especially to heteropolar pollen grains (Fig. 8) as well as pollen dispersed in permanent tetrads. When single pollen

**Fig. 6 Small and rare pollen, Paleocene, Western Greenland**. **A.** small fossil grains (≤10 μm in diameter) usually absent in samples after sieving. LM micrographs (left) in equatorial (upper) and polar view (lower). **B.** pollen in equatorial view, SEM. **C.** striate sculpture not seen under low magnification LM

**Fig. 7 Ornamentation LM vs. SEM, fossil, Middle Eocene, Western Greenland**. **A-C.** *Eucommia* sp. **A.** Pollen psilate in LM. **B.** Pollen in SEM, equatorial view, note sculpture. **C.** Ornamentation nanoechinate (≤0.5 μm) and granulate. **D-F.** *Ilex sp.,* **E.** LM and SEM overviews show the typical clavate sculpture known for this genus. **F.** Microrugulate suprasculpture present on the distal part of the clavae, only observed using high magnification SEM

**Fig. 8 Fossil heteropolar pollen grain, Paleocene, Western Greenland**. **A.** LM micrographs showing proximal (left) and distal (middle) poles of pollen grain and equatorial view (right). **B.** SEM overviews showing both poles of the pollen grain and the different aperture arrangements. **C-D.** SEM close-ups of proximal (**C**) and distal poles (**D**) show that the muri are much broader on the proximal pole

grains or tetrads are studied using SEM, changes in sculpture over the pollen surface are often observed, for example polar vs. equatorial region, mesocolpium vs. aperture region vs. aperture membrane (Fig. 9). Some pollen or tetrads also have Ubisch bodies or viscin threads (Hesse et al. 2000). These differences in the sculpture of fossil pollen need to be documented and it is therefore often necessary to show more than a single close-up taken with the SEM.

**Fig. 9 Fossil tetrad,** *Rhododendron* **sp., Miocene, North-east China**. **A, D.** Tetrad, overviews in LM vs. SEM. **B-C.** close-ups at same, magnification show difference in sculpture at polar region of pollen grain (**B**) vs. interapertural area (**C**). **E**. exine surface with viscin thread, SEM

#### **References**


```
PalDat – a palynological database (2000 onwards, www.
paldat.org)
```
Weber M, Ulrich S (2017) PalDat 3.0 – second revision of the database, including a free online publication tool. Grana 56: 257–262

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

### **Methods in Palynology**

**Preparation of Recent and Fossil Material for LM, SEM, and TEM – 98**

**Light Microscopy – 98**

**Scanning Electron Microscopy: Preparation of Recent Pollen – 105**

**Transmission Electron Microscopy: Pollen Wall Stratification and Ultrastructure – 105**

**Staining Methods – 114**

**Preparation of Fossil Pollen – 118**

**Preparation Method: From Rock to Palynomorphs – 119**

**The Single-Grain Method – 121**

**Recipes – 123**

**References – 127**

97

#### **Preparation of Recent and Fossil Material for LM, SEM, and TEM**

Multiple methods and techniques should be used when investigating pollen grains in order to provide comprehensive and accurate information about pollen morphology and ultrastructure (see also "Misinterpretations in Palynology"). The preparation methods used depend on the material to be studied, if the pollen grains are to be obtained from recent flower material (herbarium sheets, newly collected) or from various sedimentary rocks, sediments or soils (fossil to subfossil pollen). Recent and fossil pollen grains are easily studied using both LM and SEM, but recent pollen grains are also more often studied using TEM.

For an accurate description of any taxonomic value, it is important to study pollen grains in both LM and SEM. The LM will provide, among others, information on the endoaperture that cannot be obtained using SEM. Likewise the SEM will provide detailed information on the sculpture of the pollen grain that is not visible under the low magnification provided by the LM. For example, terms with "micro-" (like microreticulate) or "nano-" (like nanoechinate) can only be observed using SEM (Fig. 1).

*Annotation: The methods described in this section are the standard palynological techniques applied by the authors of this book and may differ in other working groups/labs around the world. All LM, SEM, and TEM micrographs in this book are produced following these standard protocols. Recipes for preparations are included at the end of this section.*

#### **Light Microscopy**

#### **Pollen Hydration Status at Dispersal**

To clarify the dehydration status of pollen grains at anthesis, pollen must be collected from newly opened anthers (Fig. 2). Fresh pollen grains are transferred immediately into a drop of pure glycerine and should be observed as soon as possible, as pollen grains expand in glycerine (within days or

**Fig. 1 LM vs. SEM**. **A-C**. *Aetanthus coriaceus,* Loranthaceae. **A**. Pollen grain looks psilate or scabrate in LM. **B**. Sculpture elements become visible under SEM. **C**. The sculpture elements are nano- to microbaculate and only identifiable using high magnification

**Fig. 2 Pollen hydration status at dispersal**. **A-C**. *Alocasia* sp., Araceae. **A**. Pollen grains fully hydrated at anthesis, binocular microscope. **B**. Pollen in glycerine, LM. **C**. Pollen hydrated in water, LM

weeks). The water content of pollen grains at the time of dispersal varies and pollen can be fully hydrated, partially hydrated, or partially dehydrated (Heslop-Harrison 1979; Nepi et al. 2001; see also **harmomegathic effect** in "Pollen Morphology and Ultrastructure").

#### **Pollen Hydrated in Water**

Fresh or dry pollen grains are hydrated in a drop of water on a glass slide and observed in LM. This should be the first step before preparing pollen for SEM to get an impression about the quality of the collected material, to make sure that the material is not degenerated or contaminated by fungi (Fig. 2). Observations on pollen hydrated in water with the LM can reveal interesting aspects. One example is *Montrichardia* (Araceae), where a drop of water triggers a massive expansion of the thick intine resulting in an explosive opening of the pollen wall (Weber and Halbritter 2007).

#### **Clarify the Pollen Polarity and Aperture Type**

To clarify the pollen polarity and the aperture type, anthers with pollen tetrads must be collected before anthesis (usually found in flower buds). Pollen tetrads can be released from the anthers in a drop of water or in glycerine. Quite often different developmental stages can be found in one anther: microspores in early and late tetrad stages (with or without callose wall), but also young microspores (before first pollen mitosis) released from the tetrad as well as mature pollen grains (Fig. 3; see also Fig. 1 in "Pollen Development"). For the investigation it might be useful to stain the material, e.g. with toluidine blue or basic fuchsin (Siegel 1967).

#### **Acetocarmine Staining: Detection of the Cellular Condition**

For the detection of the cellular condition of pollen grains, fresh pollen are put into a drop of acetocarmine and warmed on a heating plate (up to

**Fig. 3 Clarify the pollen polarity**. **A-B**. *Calla palustris,*  Araceae, tetrads in different stages as well as free microspores stained with toluidine blue (**A**) and basic fuchsin (**B**)

**Fig. 4 Clarification of the cellular condition using acetocarmine**. **A.** Binucleate pollen of *Anchomanes welwitschii*, Araceae, generative nucleus stains intensive red. **B**. Trinucleate pollen of *Amorphophallus krausei*, Araceae, sperm nuclei stain intensive red

70 °C), for a few seconds to several minutes (species dependent), and observed under the LM (Gerlach 1984). The generative nucleus in binucleate pollen grains and the sperm nuclei in trinucleate pollen stain intensively red with aceto-carmine (Fig. 4). The generative nucleus usually stains less intensive.

#### **Potassium Iodine: Detection of Starch**

For the detection of starch as reserves in the cytoplasm, pollen grains are stained with aqueous potassium iodine (Gerlach 1984). Fresh or dry pollen grains are transferred into a drop of staining solution on a glass slide. Starch present in pollen grains will stain dark brown to black (Fig. 5).

#### **Acetolysis: Visualizing Pollen Ornamentation and Aperture Number in Recent and Fossil Pollen**

**Acetolysis** (Erdtman 1960) is a standard palynological preparation technique and an indispensable method for illustrating pollen grains with the LM. Untreated or stained pollen grains will hide much of the important information for the description of a pollen grain. The acetolysis treatment should remove the cellular content and the intine, but can also destroy the aperture membrane. Moreover, it cleans pollen surfaces and colors pollen grains brown, which makes it easier to observe all details of the pollen wall.

The normal preparation procedure is a combination of two steps, chlorination and acetolyzation

**Fig. 5 Detection of starch using potassium iodine**. *Amorphophallus interruptus,* Araceae, starch (in amyloplasts) stained with potassium iodine

(Fig. 6). For **chlorination**, the sample is transferred to a test tube and covered with a layer (1.5 cm) of glacial acetic acid and a layer (ca. 3 cm) of a freshly prepared solution of saturated sodium chlorate. After adding 3 or 4 drops of concentrated HCl, the mixture is stirred with a glass rod, heated in a bath of boiling water for 3 min, centrifuged, and the liquid fraction decanted. The residue is carefully rinsed to eliminate any remaining chemicals and then finally washed in concentrated acetic acid or acetic anhydride to remove the water. For the **acetolyzation**, the sample is put into a mixture of 9 parts acetic anhydride and 1 part concentrated sulfuric acid and heated to 100 °C (at least 80 °C) for approximately 4 min (up to 10 min). The samples are ideally acetolyzed in an ultrasonic bath to avoid boiling retardation and to reduce water condensation. After the mixture has been centrifuged and the liquid fraction decanted, the residue is washed in acetic acid and 3 times with water. After washing, test tubes are turned upside down and the content dried. Glycerine is then added to the sample. For fossil pollen material both steps (chlorination and acetolyzation) are usually applied.

When **preparing recent material** (Fig. 7) it is routine to apply only the second step (acetolyzation). Traditionally, the term "acetolysis" is also used even when pollen grains have been acetolyzed only and not previously chlorinated. For acetolysis of recent pollen fresh or air dried pollen/anthers are transferred into test tubes and can be acetolyzed directly. For the analysis of soil, dust, honey, or any other samples, the material has to be washed in a beaker with about 200 ml distilled water (and

**Fig. 6 Acetolysis treatment**. Chlorination (**A**) and acetolyzation (**B**), the two steps of acetolysis

detergent, e.g., Tween) and can be sieved to remove bigger parts (leaves, branches) from the sample. In order to prevent pollen loss, it is important to use sieves with big mashes (E-D-quick sieve "260 μm"). The material is then concentrated in test tubes by centrifuging at 3000 rpm and the water decanted. The residue is washed in concentrated acetic acid to remove the remaining water and

**Fig. 7 Acetolyzation treatment of recent material**. **A**. Washing the sample in a beaker. **B**. Washing with a detergent "Tween". **C**. Sieving the sample. **D**. Decanting water from the test tube after centrifuging; the organic fraction remains at the bottom. **E**. Fresh acetolysis mixture is added to the sample in the test tube. **F**. Samples are heated in an ultrasonic bath. **G**. During acetolyzation the solution turns brown. **H**. Residue washed in acetic acid followed by water. **I**. Drying of the acetolyzed sample. **J**. Acetolyzed material in glycerine stored in cryo tubes. **K**. acetolyzed pollen from honey in LM

subsequently acetolyzed (see description "acetolyzation" above). For light microscopy one part of the acetolyzed material is transferred into glycerine. For scanning electron microscopy, acetolyzed pollen is transferred into a drop of anhydrous ethanol on a SEM stub and sputter coated with gold (see also below "Preparation of fossil material").

*Annotation: After rehydration or washing of the material (pollen/anthers) use acetic acid before and after the use of the acetolysis mixture, as it reacts intensively with water. Fresh acetolysis mixture is light yellow colored and highly reactive. Over*  *time the mixture obtains a dark brown color and becomes less reactive.*

#### **Heavy Liquid Separation**

Samples (recent and fossil) that still contain a very high mineral content after acetolysis should be treated with heavy liquid (e.g., zinc bromide solution; e.g., Eyring 1996, Traverse 2007; Fig. 8). Add ca. 2 cm of zinc bromide solution into the centrifuge tube and mix with the organic residue. Distilled

**Fig. 8 Heavy liquid separation**. **A**. Sample with high mineral content (light grey layers) after acetolysis. **B**. Mixing the sample with heavy liquid. **C**. Distilled water added without intermixing the liquids. **D**. Organic fraction (arrowhead) floating on the heavy liquid. **E**. Organic fraction (arrowhead) pipetted to a new test tube. **F**. Washing the organic fraction with water. **G**. Drying the acetolyzed sample (left) and the mineral fraction (right). **H**. Sample untreated (left) and treated with heavy liquid separation (right)

water is then carefully poured into the test tube (ca. 2 cm) and make sure that the two liquids do not intermix. After centrifuging for about 5–8 min at 3000 rpm the organic material is floating on the heavy liquid and below the distilled water. The organic material can then be transferred with a pipette into a new test tube for further washing. The inorganic parts remain at the bottom of the solution.

#### **Acetolysis the Fast Way**

A fast and easy way to prepare recent pollen grains for LM and SEM is to have a small glass bottle with a readymade acetolysis fluid (nine to one mix of 99% acetic anhydride and 95–97% sulfuric acid) at hand. Place a drop(s) of the acetolysis fluid on a glass slide. Remove anthers from the flowers and place them into the fluid on the glass slide (Fig. 9). To soften up the material let it lay in the liquid for some time and break the anther/flower material by squeezing and pressing it with the tip of a teasing needle. The slides are then heated over a candle flame for a short time to soften up the anthers, release the pollen grains from the anthers, dissolve extra organic material on pollen grain surfaces, "rehydrate" pollen grains and release their cell contents, and finally, to stain the pollen grains for LM photography. Make sure not to hold the slide over the flame for too long since it will make the pollen grains too dark. Best is to

**Fig. 9 Acetolysis the fast way**. **A**. Flower and tools needed for preparation. **B**. Brake or cut off anthers. **C**. Transfer anthers into acetolysis fluid on glass slide. **D**. To soften up the material it can lay in the fluid for some time. **E**. Carefully heat the slides over a candle light. **F**. Readymade pollen grains in the acetolysis fluid. **G**. Transfer pollen grains to fresh drops of glycerine on new glass slides and photograph in LM. **H**. Same grain photographed in SEM using the "single-grain method"

heat the slides shortly and then use the teasing needle to break down the anther material. This should be repeated until the pollen have gained the required color. Using a micromanipulator (see below) selected pollen grains are then transferred into fresh drops of glycerine on new glass slides and photographed under LM. Some pollen grains can also be transferred to SEM stubs using the technique of the "single-grain method" described below, sputter coated with gold and photographed under the SEM.

#### **Scanning Electron Microscopy: Preparation of Recent Pollen**

S EM techniques cannot substitute LM, but they can provide a great deal more information, especially about ornamentation. Samples prepared for SEM should ideally reflect the fully hydrated condition of a living pollen grain. In addition, all types of pollen coatings must be removed from the pollen surface, not to obscure details of the pollen wall.

For scanning electron microscopy dehydration and drying techniques are of great importance. The principle of critical point drying (CPD) is to avoid any damaging to the pollen due to surface tension forces occurring during transition from the liquid to the vapor phase. Due to the slow penetration time of DMP, large samples (e.g., large anthers, whole parts of flowers) should be dehydrated in a series of alcohol (70–85–96%, each about 20 min) and acetone or dehydrated in 70% ethanol (3 days) and formaldehyde dimethyl acetal (FDA, 1 day or overnight).

#### **The DMP Direct Method: Dimethoxypropane**

With the DMP direct method (Halbritter 1998) important details of hydrated pollen grains, which may be lost by conventional methods (alcohol), are well preserved without shrinkage, distortion, or dissolution (Fig. 10). The best results are obtained using acidified dimethoxypropane (DMP) for dehydration. Anthers should be collected at anthesis. Take whole or parts of anthers, or loose pollen grains and put them into a pouch made of filter paper. For analyzing pollen in hydrated condition, moisture the filter pouch with a droplet of water and wait for a few seconds before transferring them into acidified 2,2-dimethoxypropane. After 20–30 min (or up to 24 h) in DMP samples are transferred into pure acetone for a few minutes and critical-point dried in CO2 using acetone as the intermediate fluid. The CPD-pollen samples are then mounted on stubs using double-sided adhesive tape, sputter coated with gold and observed with an SEM. CPD samples can be stored, e.g., in a sealed plastic box to protect them from humidity.

This method can be used for fresh material as well as for herbarium samples (after rehydration in water). The chemical dehydration of unfixed plant material with DMP is a simple and fast method and can be applied to small samples only.

Unless stated otherwise, the pollen grains shown in this book are prepared using the DMP direct method by Halbritter (1998).

#### **Transmission Electron Microscopy: Pollen Wall Stratification and Ultrastructure**

F or TEM studies of recent and fossil pollen, more than one protocol for fixation and staining may be needed.

#### **Fixation and Embedding**

**Fixation** of samples for TEM studies (Hayat 2000) is a time-consuming process that starts with fixation on the first day (Fig. 11), followed by dehydration and infiltration on the second and third day and ends with embedding on the fourth day (Fig. 12). For prefixation, the samples (closed anthers or pollen suspension) are placed in phosphate buffered glutaraldehyde (3%). In case of large specimens (flower/anther), the relevant parts of the sample are prepared/cut within the fixation solution under a binocular microscope (placed at the fume hood to prevent toxic substances from inhalation). Samples must be free of gaseous/air-bubbles. Transfer samples into Eppendorf tubes and make holes into the lid. Place the tubes into the vacuum desiccator and evacuate from air for 10–30 min. For pre-fixation the evacuated samples are then placed for 6 h in a specimen rotator (at room temperature). After rinsing in buffer and distilled water, samples are postfixed in 2% osmium tetroxide plus 0.8% phosphate-buffered potassium ferrocyanide (2:1) for 8–12 h at 6 °C (for osmium storage see also Fig. 28). On the second day osmium tetroxide is removed and samples are washed in distilled water (3 times for 5 min each) followed by dehydration in 2,2-dimethoxypropane (3 times, for 10 min each) and finally by pure acetone (2 times for 15 min each). The infiltration process starts by adding a few drops of the embedding media (1:2) to the samples

**Fig. 10 The DMP direct method**. **A**. Pollen collected at anthesis, *Fuchsia magellanica*, Onagraceae. **B**. Filter pouches for pollen preparation; moisture filter pouches (pollen samples) with a droplet of water (asterisk) before dehydration in DMP (arrowhead). **C**. Critical point dryer (CPD) with closed chamber and upper view on open chamber (arrowhead). **D**. CPDpollen samples mounted on stubs using double-sided adhesive tape. **E**. Sputter coater. **F**. Samples sputter-coated with gold. **G**. SEM. **H**. Open chamber. **I**. Pollen in hydrated condition, SEM. **J**. Pollen in dry condition, SEM

and swirl the mixture. Repeat the procedure in 6–7 h, then let samples infiltrate overnight. This process has to be repeated on the third day. On the fourth day, acetone has to be removed before embedding the material: extract half of the acetone-resin-mixture with a pipette and wait for 2–3 h until the remaining acetone evaporates. After the fixation process the material should be stained intensive black (due to osmium), if not start from the beginning with new material.

The fixed material can now be transferred into embedding forms filled with fresh **embedding media** (Agar low-viscosity resin, see section "Recipes for TEM"). Polymerization takes place in an oven for about 12 h at 70 °C. After polymerization the specimen blocks can be stored in small plastic bags and are ready for ultrathin sectioning.

*Annotation: For fixation of pollen, the material must be centrifuged after each step and the fixation mixture/water/DMP must be extracted with a pipette.*

#### **Ultramicrotomy**

A lot of equipment and preliminary steps are involved in the ultramicrotomy process: **preparation of formvar film-coated grids, section-manipulators** 

**Fig. 11 Fixation and embedding day 1–2**. **A**. Closed anther for pre-fixation. **B**. Material in Eppendorf tube with fixation solution, make holes in lid before evaporation. **C**. Evacuation in vacuum desiccator (left) or manually in a syringe (right). **D**. Preparation/cutting of samples within the fixation solution under a binocular microscope (placed at the fume hood). **E**. Transfer of selected parts of the sample into small Eppendorf tubes with fixation solution (3% GA). **F**. Samples in specimen rotator. **G**. Post-fixation; arrowhead indicates sample with osmium solution. **H**. Post-fixation of samples (arrowhead) for 8–12 h at 6 °C (fridge in a fume hood) in Eppendorf tubes; Note: osmium solution stored in fridge (asterisk). **I**. Samples after 8–12 h: material blackened due to osmium (arrowhead). **J**. After removal of osmium, samples are dehydrated, followed by pure acetone

**Fig. 12 Fixation and embedding day 2–4**. **A**. Infiltration starts by repeatedly adding few drops of embedding media. **B**. Embedding solution (Agar low-viscosity resin) mixed using a magnetic stirrer. **C**. Final embedding into adequate embedding forms under binocular microscope. **D**. Polymerization at 70 °C in a thermostat oven (arrow). **E**. Examples of various embedding forms. **F**. Polymerized samples. **G**. Specimen blocks stored in small plastic bags

**and preparation of loops, specimen block trimming, semi-thin sectioning, making of glass knives, diamond knives,** and **ultra-thin sectioning**. Another indispensable equipment for ultramicrotomy are tweezers with an ultra fine pointed, curved, and angled precision tip.

#### **Formvar Film-Coated Grids**

Coated grids are made with a formvar solution (see "Recipes for TEM"; Fig. 13). New and cleaned glass slides are dipped with a special self-made "filming machine" into the formvar solution (minimize evaporation of the chloroform). The extraction speed of the slide influences the thickness of the formvar film: a thin

108 GENERAL CHAPTERS

film is produced by a slow, steady movement. After 1–2 min remove the glass slide steadily from the solution and dry for 2–3 min. The film can then be transferred onto a clean water surface (use distilled water in a clean staining cuvette). To loosen the film, cut the film with a scalpel along the edges of the slide and blow moist air (with a straw from your mouth) onto the film. In the same instance, dip the slide into the water at an angle of 45° to remove the film from the glass slide. When the film is floating on the water surface, don't pull out the slide, but let it slowly set into the cuvette. The quality of the film is indicated by the color: a thin film is grey to silver, whereas gold is too thick. Grids cleaned with chloroform are placed using fine pointed tweezers onto the film. To know which side of the grid is coated, always put one side (either

**Fig. 13 Making formvar film-coated grids**. **A**. Filming machine with holder for glass slide; filming solution should be protected from light. **B**. Glass slide dipped into formvar solution (under fume hood). **C-D**. Film cut along edges (arrowhead); arrows indicate cutting line on film (asterisk). **E**. Moisture film before dipping the slide under water; arrowhead indicates straw. **F**. Dipping the slide into the water at an angle of 45°; film partly floating on water (asterisk). **G**. Thin film floating on water surface (silver colored). **H**. Clean grids with chloroform. **I**. Shiny or dull side of the grid is visible under binocular. **J**. Grids on thin floating formvar film, arrow indicates space left for film extraction. **K**. Extraction of coated grids from the water surface using a parafilm-coated glass slide. **L**. Coated grids dried and stored in petri dish. **M**. Parafilm-coated slide with formvar coated grids, perforations (arrowhead) outline removed grids

shiny or dull side) of the grid down on the film. Make sure to leave enough space between grids and along one short margin to extract the film from the water surface. Use a parafilm-coated glass slide to extract the filmed grids: place the slide on free space of the film and dip with quick and steady motion at about 45° angle into the water and then pull out the slide again (Fig. 13K). Place the slide on a filter paper in a petri dish and let it dry. Formvar film-coated grids should be stored protected from light and dust-free (e.g., in the petri dish). To isolate the grids, use a needle to make perforations around the grids and remove them carefully with a forceps. Before ultra-thin sectioning, check the formvar film-coated grids for defects (e.g., holes, dust) under binocular microscope and place them with the filmed side up on a filter paper (see also Fig. 20 "Section pick up").

#### **Section Manipulators (Eyelash or Other Adequate Type of Hair)**

To separate and move semi-thin and ultra-thin sections floating on the water surface an **eyelash manipulator** is used. Usually a human eyelash (untreated) is fixed with glue or wax on a short glass pipette or wooden stick. The eyelashes should be cleaned with alcohol each time used and stored dust-free (e.g., covered with the back end of a bigger plastic pipette) (Fig. 14).

#### **Loops**

Loops are used to transfer ultra-thin sections onto formvar-coated grids (see Fig. 20 "Section pick up"). A loop should take up a droplet of water accurately and should fit exactly onto the grid. Therefore, two types of loops are produced (1) **circular loops**, that fit onto mesh-grids and (2) **oval** loops, used for slotgrids (Fig. 15).

Loops are made with wires from conventional electric cables (wires should not be too thick or thin). For making a circular loop a small piece of wire can be twisted around a circular object with appropriate diameter (e.g., screw driver). To produce an oval loop make a smaller circle and press it from two sides with a plier into an oval shape (fitting the grid slot). More ideally wrap the wire around a self-made model form fitting the grid size/slot. The wire of the loop is finally flattened with a hammer and the twisted (non-flattened) appendices fixed with glue or wax, e.g., on a short glass pipette. The loop should be cleaned before use with alcohol and stored free of dust.

**Fig. 14 Making a section manipulator**. **A**. Human eyelash. **B**. Technical equipment for making a section manipulator; arrowheads indicate eyelashes. **C**. eyelash fixed with glue on wooden stick

#### **Specimen Block Trimming**

Criteria for block trimming are: (1) a small sample size, (2) the location of the sample should be in the center of the block-face (trapezoid) and surrounded by resin, (3) the straightness of the blockface edges (parallel edges).

**Fig. 15 Making loops**. **A**. Technical equipment for making loops. **B**. Wire twisted around a circular object (model form) for mesh grids. **C**. Loops fixed with wax on glass pipettes (left), for storage loop covered with the back end of a disposable plastic pipette (right)

A specimen block must be trimmed (cut) to get small sections with a block-face of 4 mm by 4 mm in size (Fig. 16 O). A small block-face ensures good sectioning performance. Trimming is conducted with razor blades (for each block use a new razor blade). The block is fixed in a specimen holder and trimmed under a binocular microscope. The specimen block is trimmed into a pyramid with a trapezoid-shaped block-face. The tip of the pyramid should be cut away until you reach the appropriate level within the sample. A glass knife is used for initial cuts. If the specimen is rather big, the block-face can be larger for semi-thin sectioning (max. 4 mm2) to ensure that the area of interest is preserved. Such a large block must be trimmed further to reach the final required block-face for ultra-thin sectioning.

#### **Glass Knives**

**Glass knives** are generally used for semi-thin sectioning and are replaced by diamond knives for ultra-thin sectioning. Glass knives are produced with a "knife-maker" (Fig. 17). Specially produced glass strips (e.g., 6.4 × 25mm) are first cut into squares. The squares are then cut diagonally into two triangles, each with a knife edge (Fig. 17 E). The breaking line (stress line) indicates the quality of the knives. The left side of the glass knife is sharper and can also be used for ultra-thin sectioning, whereas the right side is used for semi-thin sectioning only. "**Glass knife boats**" (disposable plastic forms) are attached and sealed with hot melted dental wax (hot plate and ethanol burner) to the glass knife (see also Fig. 18). Glass knives should be stored dust-free and safe in a "glass knife box."

**Fig. 16 Specimen block trimming**. **A**. Specimen blocks of various shapes. **B**. Trimming is conducted with razor blades. **C**. Untrimmed specimen block with view on block-face (asterisk), arrowhead indicates position of specimen inside block. **D-N**. Blocks are trimmed into a pyramid with a +/− trapezoid shaped block-face and parallel edges. **O**. Final block-face with trapezoid form (white trapeze)

**Fig. 17 Making glass knives**. **A**. Knife maker. **B**. Glass stripes cut into squares. **C**. Squares are cut into two triangles. **D**. Two triangles (glass knives). **E**. Each triangle has a knife edge (arrowhead). **F**. Detail of triangle with knife edge, arrowhead indicating breaking (stress) line. **G**. Hot plate and ethanol burner for melting dental wax. **H**. Glass knife boats attached and sealed with hot wax using a spatula. **I**. Readymade glass knife. **J**. Knives stored in glass knife box

#### **Semi-Thin Sectioning**

Before selecting an area of the specimen block for ultra-thin sectioning, semi-thin sections are cut with an ultramicrotome, using a glass knife (Fig. 18). The settings for semi-thin sectioning are: section thickness between 0.5 and 2 μm (interference color purple to blue) and cutting speed 2 mm per second. Semi-thick sections are transferred with a loop into a drop of water on a glass slide. For a fast drying process put the slide on a hot plate (approx. 70 °C). While the water evaporates the sections will stretch. The dried sections are stained with toluidine blue on the glass slide, which can be sped up by placing the slide for max. 5 s on the hot plate. Carefully wash the slide with water and dry the glass slide in a filter paper block. The stained semi-thin sections are controlled with the LM to determine the quality of the fixation and to ensure that the appropriate area of the specimen is in the correct position for ultrathin sectioning.

#### **Ultra-Thin Sectioning**

**Ultra-thin sections** between 60 and 90 nm (interference color silver to pale gold) are cut using an ultramicrotome (Fig. 19). **Diamond knives** are more suitable for cutting plant material, as e.g., crystals in cells destroy the cutting edge of glass knives, generating scratches within the sections or even splitting the sections.

The knife is placed in the knife holder and the knife boat filled with distilled water. The knife should be clean, free of dust and moistened with water. The specimen block has to be placed in the specimen arm in the upper position. Then the block has to be positioned parallel to the knife-edge by rotational or lateral adjustments of block as well as the knife. By moving the block up and down in front of the knife a slit of reflected light helps to adjust the block to the knife. A narrow slit of light indicates that the block is close to the knife and a constant thickness of the slit, along the whole block-face, indicates that the block face and the knife-edge are parallel. This is the ideal position for sectioning. The settings for ultra-thin sectioning are: section thickness between 60 and 90 nm and cutting speed 1 mm per second. The section settings can be adjusted while cutting until pale gold to silver sections are produced. Sections are floating on the water surface and can be manipulated with an eye-lash. Before the ultra-thin sections can be transferred to grids, sections must be stretched to remove compressions due to cutting. For stretching a solvent (e.g., xylol, chloroform, acetone vapor) or a hot pen can be used. For the vapor method use a thin, wedge-shaped piece of filter paper moistened with a drop of solvent, hold it closely above the sections while moving it back and forth.

#### **Section Pick-Up**

The stretched sections are picked up from the water surface with a loop (Fig. 20). Depending on the size of the sections between 3 and 10 sections can be picked up at once. Center the loop above the selected sections, dip it on to the water surface, lift the sections up within a droplet of water and transfer onto a grid under a binocular microscope. Center the loop above the grid and lower it onto the grid surface. Lift up the loop and the attached grid. The water is removed slowly with a filter paper touching the first twist by the loop (Fig. 20 D). Transfer the grid with a forceps into a grid-box (sections should face the same side). Make a section protocol. Store the grid box away from light.

#### **Staining Methods**

T he application of different TEM staining techniques for one and the same sample is very important and highly recommended to avoid misinterpretations of the pollen wall structure. Therefore, sections of pollen grains are routinely stained using the several different staining methods (Figs. 21 and 22). Most staining solutions are harmful or even toxic and therefore applied under fume hood.

*Annotation: In electron microscopy there is no grey-scale terminology from white to black. Use "electron dense" for black or darkly colored structures and "electron translucent" for white to light grey colored.*

#### **Uranyl Acetate-Lead Citrate Staining: U + Pb**

Uranyl acetate-lead citrate staining is a conventional staining method (Hayat 2000; Figs. 21 and 22). Ultra-thin sections are usually collected on copper grids. Sections are stained in uranyl acetate solution (Leica Ultrastain-1) for 45 min followed by lead citrate staining (Leica Ultrastain-2) for 1–5 min at room temperature. Use of sodium hydroxide pellets for lead citrate staining prevents crystalline precipitation by absorbing moisture and carbon dioxide from the air. Sections are thoroughly washed in distilled water after each staining step (3 times for 5 min in a row of water drops).

**Fig. 18 Semi-thin sectioning**. **A**. Ultramicrotome. **B**. Glass knife positioned in the knife holder and knife boat filled with distilled water; specimen block fixed within the specimen holder. **C**. Block adjustment parallel to knife-edge by use of reflecting light. **D**. Knife should be clean, free of dust and moistened with water, asterisk indicates slightly lowered water level at knife edge for sectioning, but still moistened. **E**. Block must be close enough to knife (until slit of light almost disappears) to start sectioning. **F.** Semi-thin sections between 0.5 and 2 μm (interference color purple to blue) floating on water. **G-H**. Section pick-up with a loop (see "Section pick-up"). **I**. Transfer of sections in a drop of water on a glass slide. **J**. Slide on a hot plate (arrowhead indicates semi-thin sections). **K**. Staining sections with toluidine blue on hot plate. **L**. Rinsing the stained sections with water. **M**. Stained semi-thin sections ready for LM. **N**. Toluidine blue sections seen under LM. **O**. Final quality check before ultra-thin sectioning

**Fig. 19 Ultra-thin sectioning**. **A**. Specimen block holder with trimmed block, glass and diamond knife. **B**. Crystals in plant cells cut with glass knife, note scratches. **C**. Crystals in plant cells cut with diamond knife. **D-E**. Block adjusted parallel to knife-edge by use of reflecting light. **F-G**. Sections between 60 and 90°nm (interference color silver to pale gold). **H**. Stretched sections, note the change in size and thickness (for color change compare to picture **G**). **I**. Ultrastructure of a plant cell showing high quality fixation of several organelles in TEM

**Fig. 20 Section pick-up**. **A**. Loop for section pick-up. **B**. Loop centered above grid (arrowhead). **C**. Grid attached to loop (arrowhead). **D**. Water removed from grid, asterisk indicates wet filter paper. **E**. Dry grid placed into grid-box, sections on the left side (arrowhead). **F**. Grid-box and section protocol with color code used for different staining methods

#### **The Lipid Test for the Detection of Unsaturated Lipids: TCH + SP**

The endexine can be differentiated from the ektexine and the intine by thiocarbohydrazide-silver proteinate (TCH+SP) staining in osmium-fixed material. The endexine stains electron dense after the lipid test, indicating lipidic compounds (Fig. 22 B).

Ultra-thin sections on gold grids are treated with 0.2% TCH for 8–15 h and 1% SP for 30 min and thoroughly washed in water (3 times for 5 min in a row of water drops) (Rowley and Dahl 1977; Weber 1992).

#### **Thiéry-Test: PA + TCH + SP**

The Thiéry-test is used for the detection of neutral polysaccharides in osmium-free material (Thiéry 1967). Ultra-thin sections from osmium-free material are placed on gold grids and treated with 1% periodic acid (PA) for 45 min, 0.2% thiocarbohydrazide (TCH) for 8–15 h, and 1% silver proteinate (SP) for 30 min (Thiéry 1967). The polysaccharide intine and starch grains in amyloplasts stain electron dense (Fig. 22 C). For control samples leave out the thiocarbohydrazide step. If osmium fixed material is

**Fig. 21 Staining methods for ultra-thin sections**. **A**. Ultra-thin sections on copper or gold grids stained in a small drop of uranyl acetate on parafilm. **B**. Small drops of lead citrate on parafilm and sodium hydroxide pellets in a closed petri dish. **C**. Small drops of potassium permanganate on parafilm. **D**. Row of large water drops for washing placed on parafilm

used for the Thiéry-test, the staining time for 1% periodic acid has to be prolonged up to 60 min (instead of 30 min), to remove the osmium tetroxide from the material.

#### **Modified Thiéry-Test: PA + TCH + SP (short)**

The modified (short) Thiéry-test (Weber and Frosch 1995) is especially effective after fixation of specimens with osmium and potassium ferrocyanide and is a good method for general enhancement of contrast in the cytoplasm and the pollen wall (Fig. 22 D). Ultra-thin sections are collected on gold grids and stained with 1% periodic acid (PA) for 10 min, 0.2% thiocarbohydrazide (TCH) for 15 min, and 1% silver proteinate (SP) for 10 min (at room temperature). After all steps the sections are thoroughly washed in distilled water (3 times for 5 min in a row of water drops), and following the TCH first washed in 3% acetic acid.

#### **Potassium Permanganate: KMnO4**

Potassium permanganate staining is a simple method for the detection of the endexine. Using

118 GENERAL CHAPTERS

uranyl acetate and lead citrate, ektexine and endexine may differ in their electron opaqueness in that the endexine is higher in electron density than the ektexine, or vice versa. When the endexine is thin and less compact or discontinuous, the differentiation of the two layers may be insufficient. Typical for the endexine is its increasing thickness close to the aperture. Potassium permanganate stains the endexine electron dense, producing a distinct contrast (Weber and Ulrich 2010; Fig. 22 E). Ultra-thin sections from osmified material on copper grids are treated with 1% aqueous potassium permanganate solution for 7 min and thoroughly washed in water (3 times for 5 min in a row of water drops).

#### **Preparation of Fossil Pollen**

T here are numerous methods currently used to extract organic material, including fossil and subfossil pollen, from all different types of sediments (rocks) and soils. These methods have been summarized in detail by, e.g., Erdtman (1943), Brown (1960), Fægri and Iversen (1989), Moore et al. (1991), Wood et al. (1996), and Traverse (2007). Most of these preparation methods involve sieving of some sort and the final production of palynomorphs enclosed in

**Fig. 22 Stained pollen walls and behavior of endexine (cross-section, TEM)**. **A-C**. *Apium nodiflorum*, Apiaceae. **A**. Uranyl acetate + lead citrate (U + Pb), compactcontinuous endexine (asterisk). **B**. Lipid test (TCH+SP). compact-continuous endexine (asterisk) stains electron dense. **C**. Thiéry-Test (PA + TCH + SP), compact-continuous endexine (asterisk) stains electron translucent, intine electron dense (white asterisk). **D-E**. *Mentha aquatica*, Lamiaceae. **D**. Modified Thiéry-Test (PA + TCH + SP), thin compact-continuous endexine (asterisk) only slightly visible. **E**. Potassium permanganate (KMnO4), thin compact-continuous endexine (asterisk) electron dense

glycerine gelatine on sealed glass-slides. Majority of paleopalynological studies then focus on counting the quantity of each pollen type observed on the slides (often between 300 and 600 grains), with an unfortunate minor emphasis on pollen morphology and ultrastructure. The following preparation procedure has been used by the paleopalynology team at the University of Vienna for over 30 years and is suitable for most sedimentary rocks with minor variations. During preparation the solution is not sieved at any stage, so not to lose any small or exceptionally large palynomorphs, and the final solution is stored in glycerine suspension in small sample tubes so the palynomorphs can be studied using the so-called "single grain method." This method has been evolved to able researchers to obtain pollen characters from single fossil grains using both LM and SEM and sometimes TEM.

#### **Preparation Method: From Rock to Palynomorphs**

S edimentary rock samples (20–50 g) are washed and dried and hand ground in a mortar with a pestle (Fig. 23). Using a glass beaker the resulting powder is boiled in ≥200 ml of concentrated hydrochloride acid (HCl) for 5–10 min; this should remove all carbonates. Let the solution stand and when the residue has settled, decant most of the HCl liquid. Transfer the remainder of the solution into a copper pan or pot and add ≥150 ml of hydrofluoric acid (HF) and boil for approx. 10 min while stirring with a copper stick or spoon (or let stand in cold HF for 3–5 days, stir regularly, use acid-resistant plastic containers and tools); this should remove all the silicates. The solution is then poured slowly into a 4 L plastic beaker filled with water. After settling, the liquid is decanted and the remainder solution poured into glass beakers along with ≥200 ml of HCl and boiled again for 5–10 min; this prevents the formation of fluorite crystals. After cooling and settling decant most of the HCl and pour the remainder of the solution into two separate test tubes (glass centrifuge type). Wash the solutions 4 times with water and centrifuge and decant the liquid following each wash. Fill one large glass tube with cold water and add 1–2 teaspoons of sodium chlorate (pure crystalline powder; NaClO3). Shake this large tube and when there are crystals that cannot be dissolved in the water the solution is ready. Pour ca 1 ml of acetic acid glacial (100%, CH3COOH) and 3–4 ml of the sodium chlorate solution into the two original test tubes, then add five drops of HCl. Place the tubes in boiling water for at least 5 min and

**Fig. 23 From rock to palynomorphs**. **A**. Different types of sedimentary rocks: reddish, yellowish, and white-greyish samples usually contain few and/or badly preserved palynomorphs (back row), brown, dark-grey to blackish samples often contain well preserved pollen (front row). **B**. Sedimentary rock sample (ca 30 g) hand grounded in a mortar with a pestle. **C**. Sample boiling in ≥200 ml of HCl. **D**. Sample boiled in a copper pan with ≥150 ml of HF. **E**. The HF solution is poured slowly into a large plastic beaker filled with water. **F**. Organic material settled on the bottom of the beaker. **G**. Acetolysis, test tube in boiling water, note the stirring glass stick. **H**. Acetolyzed sample before decanting of water following the final wash

have a stirring glass stick in it at all times. The color of the sample solutions should change from dark blackish to brown or reddish. Centrifuge the test tubes and decant the liquid. Wash the residue 3–4 times with water and one last time with acetic acid glacial. Prepare a new solution in a clean and dry measuring glass-tube with 9 parts acetic anhydride (99%, (CH3CO)2O) and one part sulfuric acid (95–97%, H2SO4). Make sure to produce at least 10 ml of this solution for each original (fossil) test tube you process. Pour ca 10 ml of the new solution into each test tube. Direct tube away from your face and make sure no water comes into contact with the solution. Place the tubes again into the boiling water bath for at least 5 min. Then centrifuge and decant the liquid (again avoid contact with water). First wash the remaining residue once with acetic acid glacial, centrifuge and descant liquid, and then wash them 3–4 times with water. The remaining organic material in the test tube is finally mixed with glycerine and transferred, using pipettes, into small closable plastic test tubes. Test tubes are labelled accordingly.

#### **The Single-Grain Method**

A combined method for the investigation of fossil pollen grains was initiated by Daghlian (1982), suggesting that the same individual fossil grain should/could be observed in LM, SEM, and even TEM. This idea of how to properly investigate fossil pollen grains in a taxonomically valid way was taken further by Zetter (1989) who evolved a relatively easy method to investigate the same single fossil grain using the so-called "single-grain method," also described in Ferguson et al. (2007). To apply this method the following equipment and tools are necessary: samples prepared in the way described above, narrow glass-pipettes (see below, Fig. 24), teasing needle with an attached human nasal hair (see below, Fig. 25), an erect image compound microscope with a photographing unit, 10 and/or 20× objective lens with a minimum 10 mm working distance, glass slides, ethanol absolute, SEM stubs, sputter coater, and a functional scanning electron microscope.

#### **Making Glass-Pipettes**

It is important to have enough cheap and dispensable glass-pipettes to transfer parts of the sample from the storage tubes onto the glass slides for primary LM investigations. These pipettes are also used to make very small drops of ethanol on the surface

**Fig. 24 Making glass-pipettes**. **A**. Glass pipe held in burning gas flame starting to melt. **B**. Melting glass pulled very slowly and gently apart. **C**. Two freshly made pipettes ready for use

of the SEM stubs when transferring pollen from glycerine drops using the micromanipulator (see below

**Fig. 25 Producing a micromanipulator**. **A**. Needle pushed ca 1 cm into glue tube and turn in circles. **B**. Needle pressed onto a nasal hair. **C**. Extra glue added around the proximal part of the hair. **D**. Readymade micromanipulator

"Producing a Micromanipulator"). One possibility is to make your own pipettes (Fig. 24) by cutting down 4 mm wide glass pipes (cylinders) into ca 30 cm long units. The middle part of these is then held in/over a burning gas flame. While the glass starts to heat and melt you pull it apart from each end. The pipes will quickly give away in the middle as the glass melts. When pulled apart the glass will form two very long and narrow cones until they finally detach and one holds a perfect pipette in each hand.

#### **Producing a Micromanipulator**

The easiest way to make a really good and functional micromanipulator, that can be used to push around and pick out single pollen grains, is to attach a human nasal hair to a teasing needle (Fig. 25). Collect fresh nasal hairs from your professor or senior scientist (avoid the grey and white hairs) and lay them on a sheet of paper. Take a teasing needle and push it ca 1 cm into a glue tube while squeezing gently and turning the needle in circles. Pull out the needle and press onto one of the hairs already laying on the sheet of paper. Make sure that the distal end of the hair is facing the same way as the distal end of the needle and that it extends a few mm longer than the needle. When the hair is attached to the glue, add a little extra glue to cover the proximal part of the hair. Place the needle across the small opening of the glue tube, then press the tube gently for additional glue and at the same time turn the needle in circles while moving it back and forward.

#### **Applying the Single-Grain Method**

Use one of the self-made glass-pipettes to stir the sample and blow air through it to mix up the particles real good. Then suck up a tiny portion of the sample using the pipette and transfer onto a glass slide. When the tip of the pipette touches the glass slide drag the pipette along the middle section of the slide (left to right) to produce a long and relatively narrow glycerine strip. Using an erect image compound microscope (meaning when something is moved under the objective lens from left to right it is also seen moving in that same direction when observed through the eyepiece) place the glass slide under the special working distance 10× or 20× objective lens and move the distal end of the micromanipulator in-between the glass slide and the lens and then gently press the tip of the micromanipulator (the nasal hair) into the glycerine (Fig. 26 A-B). Using the micromanipulator grains of particular interest are brushed or pushed to the edge of the glycerine, then out of the glycerine until they are attached to the nasal hair and can be picked up and transferred to another glass slide (Fig. 26 C-H). Have a fresh drop of glycerine ready on a new glass slide. Dip the tip of the hair with the attached pollen into the glycerine drop and the pollen will automatically detach from the hair and rest in the glycerine. Because no cover slip is used this pollen can now be turned around with help from the micromanipulator and photographed in polar and equatorial views as well under different foci (high-, low focus, optical section), documenting important features such as sculpture, apertures, and thickenings or thinnings of the pollen wall (Fig. 27 A-D). After this, the pollen grain is transferred to a SEM stub to which a drop of absolute ethanol has been added to remove all traces of the glycerine from the surface of the pollen grains (Fig. 27 E-G). For this, the best way is to position the light microscope close beside a binocular stereoscope. Place a single SEM stub under the stereoscope and have a small container with fresh ethanol at your side as well as one of the glass-pipettes mentioned above. First pick out a pollen grain with the micromanipulator from the glycerine drop and slowly move over to the stereoscope. Dip the tip of the pipette into the ethanol container and it will automatically suck up a small portion of the ethanol. Press the tip of the pipette on the surface of the SEM stub to leave a tiny drop of ethanol. Then gently press the tip of the nasal hair with the attached pollen into the drop of ethanol and the pollen will be detached from the hair, float a bit in the drop and finally rest on the stub surface when the ethanol evaporates. Try to make the ethanol drops small and close to the center of the SEM stub. Up to 10 different types of grains can be placed on a single stub and additional ethanol drops can be added to clean the glycerine thoroughly off the pollen grains. The stub is then sputter coated with gold and the pollen photographed using a SEM (overviews and closeups). Pollen of particular interests can be turned. Add a drop of ethanol to the sputtered sample and flip the grain over using the micromanipulator before the ethanol evaporates (under the stereoscope). Re-sputter the sample and photograph it again using the SEM. This applies especially to any kind of heteropolar pollen/spores or tetrads of some sort.

#### **Recipes**

#### **Recipes for Light Microscopy (LM)**

#### **Acetocarmine (Staining)**

30 g acetocarmine + 2 L 45% acetic acid, 4 h boiled and filtered.

#### **Potassium Iodine (Lugol's Iodine, Detection of Starch)**

2 g potassium iodine + 1 g iodine + 100 ml distilled water

#### **Toluidine Blue (Staining)**

0.1 g Toluidine blue + 100 ml 2.5% sodium carbonate (NaCO3); durable at +4 °C

#### **Chlorination Mixture**

Acetic acid (CH3COOH) + saturated sodium chlorate (NaClO3)\* + 3–5 drops hydrochloric acid (conc. HCl)

\*Saturated sodium chlorate solution: about 10 g of NaClO3 in 10 ml distilled water (25 °C); the solution is saturated when crystals are still present.

*Annotation: solubility of sodium chlorate is depending on the temperature of water.*

#### **Acetolysis Mixture**

Acetolysis mixture: 9 parts acetic anhydride (99%) are mixed with 1 part concentrated sulfuric acid (96%).

#### **Zinc Bromide Solution (Heavy Liquid Separation for Samples with a High Mineral Content)**

250 g zinc bromide (Merck 8.18631.0250) + 25 ml 10% HCl\*, mix until all zinc bromide is solved (takes some time!), then add 100 ml distilled water.

\*10% HCl-Lösung: 27 ml H2O + 10 ml HCl (37%) = 37 ml 10% HCl

#### **Recipes for Scanning Electron Microscopy (SEM)**

#### **Dimethoxypropane (Dehydration)**

30 ml 2,2-dimethoxypropane (DMP) + 1 drop 0.2 n hydrochloric acid (HCl)

**Fig. 26 Applying the single-grain method — Part 1**. **A**. Sample on a glass slide under LM, working distance from sample to objective approx. 1 cm. **B**. Organic-rich sample and the tip of a nasal hair seen through the LM. **C**. Fossil pollen grain being brushed/pushed towards the margin of the glycerine. **D**. Fossil pollen grain pushed further away from the glycerine. **E**. Grain out of glycerine and ready to be picked up by the nasal hair. **F**. Pollen pushed a bit further. **G**. In a pushing or brushing motion the pollen is picked up from the glass slide. **H**. Single fossil pollen grain attached to tip of nasal hair

124 GENERAL CHAPTERS

**Fig. 27 Applying the single-grain method — Part 2**. **A**. Selected well-preserved pollen grains in a fresh drop of glycerine. **B**. Light microscope equipped with a photographic unit to document pollen grains and their diagnostic features. **C**. Pollen turned and photographed in equatorial view. **D**. Same pollen grain turned and photographed in polar view. **E**. Arrangement of the light microscope and stereomicroscope along with a bottle of ethanol and other tools used when transporting pollen grains from glass slides over to SEM stubs. **F**. Cleaned SEM stub under a stereomicroscope waiting for fossil pollen grains. **G**. How to hold the pipette with the ethanol (left) and the micromanipulator (right) when transferring fossil pollen grains onto SEM stubs. **H**. Photographing fossil pollen using SEM

#### **Recipes for Transmission Electron Microscopy (TEM)**

#### **3% Glutaraldehyde (Fixation)**

100 ml glutaraldehyde: 12 ml glutaraldehyde (GA, 25%) + 88 ml phosphate buffer (pH 7.2).

#### **1 % Osmium Tetroxide (Fixation)**

0.1 g osmium tetroxide (OsO4) + 10 ml distilled water. Osmium can be acquired in crystalline form within glass ampullae. The osmium crystals usually adhere inside the ampulla and can be loosened by dipping the ampulla in liquid nitrogen (in a styrofoam box). The ampulla can then be opened and the osmium crystals transferred into distilled water in a vial. Close the vial and seal it with parafilm. For faster dissolution, place the vial in an ultrasonic bath. Mix the osmium solution and pipette it into a vapor-tight bottle. Store it at 6 °C.

*Annotation: osmium is volatile and toxic, use in fume hood only; for storage, use oil with high percentage of unsaturated fatty acids (e.g. corn oil) to bind volatiles of osmium tetroxide (*Fig. 28*)*.

#### **Phosphate Buffer pH 7.2 (Fixation)**

1 phosphate buffer saline tablet (phosphate buffer saline tablets, Na2HPO4·2H2O, sodium hydrogen phosphate) + 200 ml distilled water (dispense tablet in ultrasonic bath).

#### **0.8% Potassium Ferrocyanide (Accelerator for Osmium)**

0.1 g potassium hexacyanoferrate (II) (K4Fe(CN)6·3H2O) + 12.5 ml distilled water (dispense in ultrasonic bath).

*Annotation: the fresh solution is uncolored and becomes yellow after a few days.*

#### **Agar Low Viscosity Resin Kit (Embedding)**

LV-resin (Agar Scientific): 48 g LV Resin + 8 g hardener VH1 + 44 g hardener VH2 + 2.5 ml accelerator.

*Annotation: Mix the embedding solution in a disposable plastic beaker by using a magnetic stirrer. The first two components must be mixed first before adding the remaining ingredients, then mix well again. The mixture can be used immediately for infiltration and then for embedding. Embedding solution can be stored in a freezer.*

#### **Potassium Iodine (Staining)**

3 g potassium iodide + 7 g iodine + 100 ml ethanol (92%).

**Fig. 28 Osmium storage**. **A**. Osmium solution stored at 6 °C (fridge placed in a fume hood). **B-D**. Osmium solution in a sealed bottle and stored in a plastic container, plastic container placed in glass vessel containing oil. **C**. Arrowheads showing osmium contamination from volatiles. **D**. Second glass vessel placed over the osmium containers, osmium vapor is bound to the oil and cannot escape into the atmosphere

#### **1% Potassium Permanganate (Fixation and Staining)**

1% potassium permanganate: 1 g potassium permanganate in 100 ml distilled water

#### **1% Periodic Acid (Staining)**

1 g periodic acid (PA, Firma Fluka) + 100 ml distilled water

#### **0.2% Thiocarbohydrazide (Staining)**

0.2 g thiocarbohydrazide (TCH, by *Serva*) + 100 ml 20% acetic acid (20 ml 100 % CH3COOH + 80 ml distilled water)

#### **1% Silver Proteinate (Staining)**

0.25 g silver proteinate (SP, by *Merck*) + 25 ml distilled water

#### **Uranyl Acetate (Staining)**

Prefabricated solution: "Ultrostain 1" by Leica

#### **Lead Citrate (Staining)**

Prefabricated solution: "Ultrostain 2" by Leica; used with potassium hydroxide pellets

#### **Formvar Filming Solution (Film-Coated Grids)**

2 g formvar (15/45 E) + 100 ml chloroform (pure); mix with a magnetic stirrer

#### **References**


**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# **Illustrated Pollen Terms**

This part is divided into 6 topic related chapters: "Pollen- and Dispersal Units," "Shape and Polarity," "Aperture," "Ornamentation," "Pollen Wall," and "Pollen Class." Terms are either morphologically or alphabetically grouped depending on practical use. When a term is illustrated by numerous images (one or more plates), the definition of the term occurs along with the first image. Features are often highlighted (colored) for easy recognition. Each image is accompanied by the name of the plant species illustrated, the current family name, and a short relevant description. The majority of the micrographs are SEM pictures, but include also LM and TEM. The SEM micrographs usually represent the turgescent (hydrated) state of recent pollen, but they can also be in dry condition or fossilized. The LM micrographs usually show acetolyzed pollen, but pollen grains can also be hydrated in water, glycerine, or stained with biological stains. Exceptions from the standard method (LM, SEM) are specified in the picture legend. For TEM micrographs the staining method is provided when necessary.

#### **Contents**

**Pollen– and Dispersal Units – 131 Shape and Polarity – 155 Aperture – 207 Ornamentation – 295 Pollen Wall – 379 Pollen Class – 429**

© Springer International Publishing AG 2018

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, https://doi.org/10.1007/978-3-319-73482-8

### **Pollen- and Dispersal Units**

**monad – 132**

**dyad – 134**

**pseudomonad – 135**

**tetrad – 137**

**tetrad tetrahedral – 138**

**tetrad decussate – 142**

**tetrad planar – 143**

**polyad – 145**

**massula – 147**

**pollinium – 148**

**pollinarium – 150**

131

#### **monad**

unit consisting of a single pollen grain

#### **dyad**

unit of two pollen grains

#### **pseudomonad**

unit of a permanent tetrad with three rudimentary pollen grains

#### **tetrad**

unit of four pollen grains

tetrad planar (left) and decussate (right)

#### **tetrad tetrahedral**

unit of four pollen grains in which the centers of the grains define a tetrahedron

#### **tetrad decussate**

unit of four pollen grains arranged in two pairs in two different plains

142 ILLUSTRATED POLLEN TERMS

#### **tetrad planar**

unit of four pollen grains arranged in one plane: tetragonal, T-shaped, linear

#### **polyad**

unit of more than four pollen grains (multiple of 4)


*Albizia saman,* Fabaceae polyad with 32 monads

146 ILLUSTRATED POLLEN TERMS

#### **massula**

unit of more than four pollen grains but less than the locular content of a theca *Comment*: In angiosperms only used for Orchidaceae with sectile pollinia

#### **pollinium**

unit of a more or less interconnected loculiform pollen mass *Comment*: loculi may be subdivided by septae, thus resulting in more than two pollinia

#### **pollinarium**

dispersal unit of pollinium (or pollinia) plus secretions and/or tissues that aid in the removal of the structure from the flower

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

### **Shape and Polarity**

**outline circular – 156 outline elliptic – 158 outline lobate – 160 outline triangular – 163 outline quadrangular – 166 outline polygonal – 167 P/E-ratio, oblate – 168 P/E-ratio, isodiametric – 171 P/E-ratio, prolate – 173 isopolar – 176 heteropolar – 178 shape – 181 saccus/saccate, corpus – 188 saccus, monosaccate – 189 saccus, bisaccate – 190 saccus, trisaccate – 193 infoldings, apertures sunken – 194 infoldings, boat-shaped – 196 infoldings, cup-shaped – 199 infoldings, interapertural area sunken – 201**

**infoldings, irregular – 203**

155

#### **outline circular**

outline describes the contour of pollen grains in polar and/or equatorial view

156 ILLUSTRATED POLLEN TERMS

#### **outline elliptic**

#### **outline lobate**

outline in polar view of a pollen grain with bulged interapertural areas (mainly in dry pollen grains)

162 ILLUSTRATED POLLEN TERMS

#### **outline triangular**


#### **outline quadrangular**

#### **outline polygonal**

#### **P/E-ratio, oblate**

P/E-ratio refers to the length of the polar axis between the two poles compared to the equatorial diameter oblate: pollen grain with a polar axis shorter than the equatorial diameter

170 ILLUSTRATED POLLEN TERMS

#### **P/E-ratio, isodiametric**

isodiametric: pollen grain with a polar axis equal to the equatorial diameter

#### **P/E-ratio, prolate**

prolate: pollen grain with a polar axis longer than the equatorial diameter

*Lathyrus tuberosus,* Fabaceae tricolporate, dry pollen

*Salvia sclarea,* Lamiaceae hexacolpate, dry pollen

#### **isopolar**

pollen grain with identical proximal and distal faces

176 ILLUSTRATED POLLEN TERMS

#### **heteropolar**

pollen grain with different proximal and distal faces

178 ILLUSTRATED POLLEN TERMS

tetrads

#### **shape**

3-dimensional form of a pollen grain in relation to the P/E-ratio

184 ILLUSTRATED POLLEN TERMS

*Clarkia unguiculata,* Onagraceae triangular star

*Fuchsia paniculata,* Onagraceae

lemon-shaped

186 ILLUSTRATED POLLEN TERMS

#### **saccus/saccate, corpus**

saccus: exinous expansion forming an air sac corpus: body of a saccate pollen grain

*Tsuga canadensis,* Pinaceae equatorial view

*Dacrycarpus dacrydioides,* Podocarpaceae

trisaccate, proximal polar view

#### **saccus, monosaccate**

monosaccate: pollen grain with a single saccus

#### **saccus, bisaccate**

bisaccate: pollen grain with two sacci

192 ILLUSTRATED POLLEN TERMS

#### **saccus, trisaccate**

trisaccate: pollen grain with three sacci

#### **infoldings, apertures sunken**

infoldings (dry pollen): consequence of harmomegathy in dry condition

*Moehringia muscosa,* Caryophyllaceae pantoporate

*Anemone hortensis,* Ranunculaceae spiraperturate

#### **infoldings, boat-shaped**

#### **infoldings, cup-shaped**

#### **infoldings, interapertural area sunken**

202 ILLUSTRATED POLLEN TERMS

#### **infoldings, irregular**

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

### **Aperture**

**angulaperturate – 208 annulus/annulate – 209 aperture/aperturate – 211 aperture membrane – 214 atrium, oncus – 220 brevicolpus/brevicolpate – 221 brevicolporus/brevicolporate – 222 bridge – 223 colpus/colpate, dicolpate – 225 colpus/colpate, tricolpate – 226 colpus/colpate, tetracolpate, pentacolpate – 229 colpus/colpate, hexacolpate – 230 colpus/colpate, stephanocolpate – 231 colpus/colpate, pantocolpate – 232 colporus/colporate, dicolporate – 233 colporus/colporate, tricolporate – 234 colporus/colporate, tetracolporate – 237 colporus/colporate, pentacolporate, stephanocolporate – 238 colporus/colporate, stephanocolporate – 239 ektoaperture, endoaperture, lalongate, lolongate – 240 heteroaperturate – 242 inaperturate – 244 leptoma – 247 margo – 248**

**operculum/operculate – 251 pantoaperturate, pantocolpate – 255 pantoaperturate, pantocolporate, pantoporate – 256 pantoaperturate, pantoporate – 257 papilla – 259 planaperturate – 260 pontoperculum/pontoperculate – 262 porus/porate, diporate – 263 porus/porate, triporate – 264 porus/porate, tetraporate – 266 porus/porate, pentaporate – 267 porus/porate, stephanoporate – 268 porus/porate, pantoporate – 269 poroid/poroidate – 272 pseudocolpus – 273 ring-like aperture – 274 spiral aperture/spiraperturate – 277 stephanoaperturate – 279 sulcus/sulcate – 281 sulcus/sulcate, disulcate – 287 sulcus/sulcate, trichotomosulcate – 288 sulcus/sulcate, polychotomosulcate – 289 synaperturate, syncolpate, syncolporate – 290 synaperturate, syncolporate – 291 ulcus/ulcerate – 292**

207

#### **angulaperturate**

pollen grain with an angular outline where the apertures are located at the angles

#### **annulus/annulate**

ring like wall thickening surrounding a porus or ulcus

#### **aperture/aperturate**

region of the pollen wall that differs morphologically and/or anatomically significantly from the rest of the pollen wall

cross section of aperture, apertural intine protrusion

cross section of aperture, apertural intine protrusion

#### **aperture membrane**

exine layer covering an aperture aperture membrane psilate

ILLUSTRATED POLLEN TERMS 215

brevi(3)colporate

*Roridula gorgonias,* Roridulaceae tricolpate

*Carthamus lanatus,* Asteraceae brevi(3)colporate

tricolpate

*Chenopodium hybridum,* Amaranthaceae cross section of aperture

*Aesculus carnea,* Sapindaceae cross section of aperture

#### **atrium, oncus**

atrium: space between diverging exine layers within the aperture oncus: lens-shaped body located beneath the aperture, not resistant to acetolysis

#### **brevicolpus/brevicolpate**

short colpus situated equatorially

#### **brevicolporus/brevicolporate**

short colpus in a compound aperture situated equatorially

#### **bridge**

exine connection(s) between the margins of an aperture

the term bridge is used in a more general context, e.g., for exine connections within tetrads

#### **colpus/colpate, dicolpate**

colpus: elongated aperture (length/width ratio >2) situated at the equator or globally distributed dicolpate: pollen grain with two colpi

#### **colpus/colpate, tricolpate**

tricolpate: pollen grain with three colpi

#### **colpus/colpate, tetracolpate, pentacolpate**

tetra- and pentacolpate: pollen grain with four or five colpi

#### **colpus/colpate, hexacolpate**

hexacolpate: pollen grain with six colpi

#### **colpus/colpate, stephanocolpate**

stephanocolpate: colpi situated at the equator (term usually used for six or more apertures)

#### **colpus/colpate, pantocolpate**

pantocolpate: pollen grain with colpi distributed more or less regularly over the surface

#### **colporus/colporate, dicolporate**

colporus: compound aperture composed of a colpus (ektoaperture) combined with an endoaperture of variable size and shape

dicolporate: pollen grain with two colpori

#### **colporus/colporate, tricolporate**

tricolporate: pollen grain with three colpori

#### **colporus/colporate, tetracolporate**

tetracolporate: pollen grain with 4 colpori

238 ILLUSTRATED POLLEN TERMS

#### **colporus/colporate, pentacolporate, stephanocolporate**

pentacolporate: pollen grain with five colpori stephanocolporate: colpori situated at the equator (term usually used for 6 or more apertures)

#### **colporus/colporate, stephanocolporate**

#### **ektoaperture, endoaperture, lalongate, lolongate**

ectoaperture: outer part of a compound aperture endoaperture: inner part of a compound aperture

*Dictamnus albus,* Rutaceae ektoaperture: colpus, endoaperture: lalongate porus *Scaevola aemula,* Goodeniaceae ektoaperture: colpus, endoaperture: lalongate porus

*Vitaliana primuliflora,* Primulaceae

*Vicia faba,* Fabaceae

ektoaperture: colpus, endoaperture: lalongate porus

*Rumex* sp.*,* Polygonaceae fossil, Quaternary Austria, ektoaperture: colpus, endoaperture: lolongate porus

ektoaperture: colpus, endoaperture: lalongate porus

*Phacelia campanularia,* Boraginaceae heteroaperturate pollen, ektoaperture: colpus, endoaperture lolongate porus

#### **heteroaperturate**

pollen grain with different types of apertures; only one type presumed to function as germination site

#### **inaperturate**

pollen grain without distinct apertures

#### **leptoma**

thinning of the pollen wall on the distal face in conifers, presumed to function as germination area

#### **margo**

exine area with different ornamentation bordering a colpus/colporus/sulcus

#### **operculum/operculate**

distinctly delimited exine structure covering an aperture


cross section of aperture

cross section of aperture

#### **pantoaperturate, pantocolpate**

pollen grain with apertures distributed more or less regularly over the surface

#### **pantoaperturate, pantocolporate, pantoporate**

pantocolporate, pantoporate

#### **pantoaperturate, pantoporate**

#### **papilla**

small protuberance typical for Taxodioideae pollen located distally

*Metasequoia glyptostroboides,* Cupressaceae

dry pollen

*Cunninghamia lanceolata,* Cupressaceae oblique distal polar view

#### **planaperturate**

pollen grain with an angular outline, where the apertures are situated between the angles

*Pachira aquatica,* Malvaceae tricolpate


#### **pontoperculum/pontoperculate**

elongated operculum linked to the ends of the aperture

#### **porus/porate, diporate**

porus: more or less circular aperture; pori located at the equator or regularly spread over the pollen grain diporate: pollen grains with two pori

#### **porus/porate, triporate**

triporate: pollen grain with three pori

#### **porus/porate, tetraporate**

tetraporate: pollen grain with four pori

#### **porus/porate, pentaporate**

pentaporate: pollen grain with five pori

#### **porus/porate, stephanoporate**

stephanoporate: pori situated at the equator (term usually used for six or more apertures)

#### **porus/porate, pantoporate**

pantoporate: pori distributed more or less regularly over the surface

#### **poroid/poroidate**

indistinct circular or elliptic aperture

#### **pseudocolpus**

colpus in a heteroaperturate pollen grain, presumed not to function as germination site

#### **ring-like aperture**

274 ILLUSTRATED POLLEN TERMS

circumferential aperture (situated more or less equatorially or, rarely, meridionally)

*Passiflora amethystina,* Passifloraceae debatable: 3 ring-like apertures or triporate with opercula

*Acacia dealbata,* Fabaceae close-up of polyad

aperture running meridionally *Acacia dealbata,* Fabaceae

*Pedicularis gyroflexa,* Orobanchaceae

debatable: polyad, monads with ring-like structure/thinning

#### **spiral aperture/spiraperturate**

elongated, coiled aperture

#### **stephanoaperturate**

apertures situated at the equator (term usually used for more than six apertures): stephanocolpate, stephanocolporate, stephanoporate

stephanocolpate, stephanocolporate, stephanoporate

#### **sulcus/sulcate**

elongated aperture located distally

282 ILLUSTRATED POLLEN TERMS


#### **sulcus/sulcate, disulcate**

disulcate: pollen grain with two sulci

#### **sulcus/sulcate, trichotomosulcate**

trichotomosulcus/trichotomosulcate: 3-radiate sulcus

#### **sulcus/sulcate, polychotomosulcate**

polychotomosulcus/polychotomosulcate: sulcus with more than three arms

#### **synaperturate, syncolpate, syncolporate**

synaperturate: pollen grain with anastomosing apertures syncolpate: pollen grain with anastomosing colpi

oblique equatorial view

dry pollen

#### **synaperturate, syncolporate**

syncolporate: pollen grain with anastomosing colpori

#### **ulcus/ulcerate**

more or less circular aperture located distally

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

### **Ornamentation**

**areola/areolate – 296 baculum/baculate – 298 bireticulate – 299 clava/clavate, caput – 302 clypeate – 305 croton pattern – 307 echinus/echinate – 309 fossula/fossulate – 315 foveola/foveolate – 317 free-standing columellae – 319 gemma/gemmate – 321 granulum/granulate – 323 heterobrochate – 324 homobrochate – 326 lophae/lophate, lacunae – 328 micro- – 331 nano- – 336 perforate – 342 plicae/plicate – 344 psilate – 346 reticulum/reticulate – 348 reticulum cristatum – 355 rugulae/rugulate – 358 scabrate – 360 striae/striate – 361 striato-reticulate – 366 suprasculpture – 370 supratectal element – 374 verruca/verrucate – 375**

295

#### **areola/areolate**

insular ornamentation element

areolae reticulate

*Cynodon dactylon,* Poaceae *Justicia brandegeeana,* Acanthaceae areolae in aperture area, oblique equatorial view

tetrad

#### **baculum/baculate**

rod-like, free standing element (never pointed)

#### **bireticulate**

reticulate ornamentation, where the lumina of the coarse-meshed reticulum are filled by a fine-meshed reticulum

#### **clava/clavate, caput**

clava: club-shaped element caput: distal part of clava


*Ilex* sp.*,* Aquifoliaceae equatorial view

*Ilex aquifolium,* Aquifoliaceae

*Linum flavum,* Linaceae *Aratitiyopea lopezii,* Xyridaceae clavae of different size

*Jatropha multifida,* Euphorbiaceae

*Hymenocallis littoralis,* Amaryllidaceae *Viburnum lantana,* Adoxaceae

*Geranium pratense,* Geraniaceae reticulum cristatum with clavae

304 ILLUSTRATED POLLEN TERMS

*Plumbago auriculata,* Plumbaginaceae

*Geranium sibiricum,* Geraniaceae reticulum cristatum with clavae

#### **clypeate**

pollen with exine subdivided into shields

#### **croton pattern**

special type of reticulum cristatum formed by regularly arranged suprasculpture elements on muri

*Daphne cneorum,* Thymelaeaceae *Thymelaea passerina,* Thymelaeaceae

surface detail with porus

#### **echinus/echinate**

pointed ornamentation element

310 ILLUSTRATED POLLEN TERMS

*Portulaca grandiflora,* Portulacaceae *Nuphar lutea,* Nymphaeaceae


*Petasites albus,* Asteraceae dry pollen

*Antennaria dioica,* Asteraceae

*Malva neglecta,* Malvaceae *Tanacetum corymbosum,* Asteraceae

*Lonicera fragrantissima,* Caprifoliaceae *Ipomoea purpurea,* Convolvulaceae

pantoporate

#### **fossula/fossulate**

irregular shaped groove

#### **foveola/foveolate**

tricolpate, polar view

roundish lumen more than 1 μm in diameter; distance between two adjacent lumina larger than their diameter

*Maianthemum stellatum,* Asparagaceae *Hohenbergia stellata,* Bromeliaceae

*Aechmea ampla,* Bromeliaceae tetraporate

*Aechmea ampla,* Bromeliaceae

foveolate to reticulate

#### **free-standing columellae**

columellae not covered by a tectum in semitectate pollen grains

*Viburnum opulus,* Adoxaceae *Thladiantha hookeri,* Cucurbitaceae


*Fouquieria macdougalii,* Fouquieriaceae *Ardisia elliptica,* Primulaceae

*Schotia brachypetala,* Fabaceae *Passiflora amethystina,* Passifloraceae

#### **gemma/gemmate**

globular ornamentation element

*Stenandrium dulce,* Acanthaceae gemmae of different size

*Fatsia japonica,* Araliaceae gemmate, reticulate

*Hakea kippistiana,* Proteaceae gemmate, microreticulate

gemmate, microgemmate

ILLUSTRATED POLLEN TERMS 321

322 ILLUSTRATED POLLEN TERMS

gemmate to rugulate

#### **granulum/granulate**

sculpture element of different/indefinable shape, equal or smaller than 0.1 μm in diameter (hard to outline)

#### **heterobrochate**

reticulate pollen wall with lumina of different sizes

equatorial view

*Hedera helix,* Araliaceae *Lachenalia aloides,* Asparagaceae

*Fritillaria meleagris,* Liliaceae *Anthericum ramosum,* Asparagaceae

*Billbergia seidelii,* Bromeliaceae *Limodorum abortivum,* Orchidaceae

#### **homobrochate**

reticulate pollen wall with lumina of uniform size

*Eranthemum wattii,* Acanthaceae


polar view

*Ruellia* sp.*,* Acanthaceae *Thlaspi montanum,* Brassicaceae

	- triporate, oblique view

#### **lophae/lophate, lacunae**

lophae: massive exine ridges lacunae: depressed areas surrounded by lophae

*Tragopogon orientalis,* Asteraceae polar view

*Cichorium intybus,* Asteraceae oblique view

*Tragopogon dubius,* Asteraceae equatorial view

*Prenanthes purpurea,* Asteraceae equatorial view

*Taraxacum* sp.*,* Asteraceae fossil, Quaternary, Austria, equatorial view

*Crepis biennis,* Asteraceae

polar view

#### **micro-**

prefix for small; features between 1 and 0.5 μm

*Heloniopsis kawanoi,* Melanthiaceae microechinate, granulate

*Kickxia spuria,* Plantaginaceae micro- to nanoreticulate

*Aspidistra elatior,* Asparagaceae microgemmate


*Orobanche lutea,* Orobanchaceae microgemmate, microgemmae composed of conglomerate granula

*Claytonia perfoliata,* Montiaceae microechinate, perforate

*Veronica prostrata,* Plantaginaceae striato-microreticulate

*Trillium chloropetalum,* Melanthiaceae microgemmate, microgemmae irregularly shaped

*Petrorhagia prolifera,* Caryophyllaceae microechinate, perforate

*Ptelea trifoliolata,* Rutaceae striato-microreticulate


*Tradescantia spathacea,* Commelinaceae microverrucate, perforate

*Smilax spinosa,* Smilacaceae microechinate, rugulate to microrugulate; (micro)rugulae segmented

*Quercus robur,* Fagaceae microgemmate to granulate

*Aspidistra locii,* Asparagaceae microverrucate to verrucate, microreticulate

*Clethra arborea,* Clethraceae microrugulate

*Erica pageana,* Ericaceae micro- to nanogemmate

*Anemone pratensis,* Ranunculaceae microechinate, perforate

*Drosera kansaiensis,* Droseraceae echinate, microclavate

#### **nano-**

nanogemmate

nano- to microgemmate

336 ILLUSTRATED POLLEN TERMS


*Carex filiformis,* Cyperaceae nanogemmate to nanoverrucate

*Viola calcarata,* Violaceae nanoreticulate

Cyperaceae fossil, Miocene, China, nanoverrucate

*Tilia* sp., Malvaceae fossil, Miocene, China, nanoreticulate

Rosaceae fossil, Miocene, China, nano- to microechinate

nanogemmate

*Rhaphidophora africana,* Araceae


*Arceuthobium sp.,* Santalaceae fossil, Miocene, China, nanoverrucate, echinate

*Drosera binata,* Droseraceae nanoclavate, echinate

*Heliohebe raoulii*, Plantaginaceae nanorugulate

Caprifoliaceae fossil, Miocene, China, granulate, nanoverrucate, echinate

*Alisma* sp., Alismataceae fossil, Miocene, China, nanoechinate

*Maclura pomifera*, Moraceae nanoechinate

nanoechinate, nanorugulate


#### **perforate**

pollen wall with holes less than 1 μm in diameter

*Lysimachia nemorum*, Primulaceae

#### **plicae/plicate**

coarse parallel ridges

#### **psilate**

pollen wall with smooth surface

346 ILLUSTRATED POLLEN TERMS

#### **reticulum/reticulate**

reticulum: network like pattern consisting of muri and lumina


*Luffa cylindrica*, Cucurbitaceae lumen (space enclosed by muri) colored

*Aechmea azurea*, Bromeliaceae

*Razisea citrina,* Acanthaceae equatorial view

	- *Ecballium elaterium,* Cucurbitaceae

*Ruellia brevifolia*, Acanthaceae

*Impatiens glandulifera*, Balsaminaceae

*Opuntia paraguayensis,* Cactaceae free standing columellae

*Buxus sempervirens*, Buxaceae

*Lupinus polyphyllus*, Fabaceae reticulate, perforate

*Theobroma cacao,* Malvaceae reticulate, perforate

*Tropaeolum majus*, Tropaeolaceae reticulate, perforate

*Werauhia tarmaensis*, Bromeliaceae

*Poncirus trifoliata,* Rutaceae

*Veratrum album*, Melanthiaceae

*Erythronium dens-canis*, Liliaceae reticulate with suprasculpture

*Melilotus officinalis,* Fabaceae reticulate, perforate

*Vigna speciosa*, Fabaceae polar view

*Harpochilus neesianus*, Acanthaceae reticulate, brochi with inclined columellae

*Adenia fruticosa,* Passifloraceae incomplete reticulum

*Epipactis helleborine*, Orchidaceae incomplete reticulum

*Polygala major*, Polygalaceae reticulate to foveolate

*Pisum sativum,* Fabaceae reticulate, perforate

*Billardiera heterophylla*, Pittosporaceae

reticulate to rugulate

*Aechmea allenii,* Bromeliaceae reticulate to foveolate


#### **reticulum cristatum**

special type of reticulum; muri with prominent suprasculpture

proximal polar view

*Pachysandra terminalis*, Buxaceae

*Aponogeton masoalaensis,* Aponogetonaceae

reticulum with microechini

*Mercurialis perennis*, Euphorbiaceae reticulum with microechini

*Armeria maritima*, Plumbaginaceae

#### **rugulae/rugulate**

elongated ornamentation elements irregularly arranged

*Securigera varia*, Fabaceae rugulae colored

*Peucedanum cervaria*, Apiaceae

*Sedum acre,* Crassulaceae rugulate, perforate

*Zelkova* sp., Ulmaceae fossil, Miocene, Austria, polar view

*Fagus* sp., Fagaceae fossil, Miocene, Austria, polar view

*Circaea lutetiana*, Onagraceae rugulate, perforate

*Nymphoides peltata*, Menyanthaceae

*Nicotiana tabacum,* Solanaceae rugulate, perforate

*Leucadendron discolor*, Proteaceae rugulate, perforate or microreticulate

*Myrrhis odorata*, Apiaceae

#### **scabrate**

term used for light microscopy only, describing minute sculpture elements of undefined shape and of a size close to the resolution limit of the light microscope

#### **striae/striate**

elongated ornamentation elements separated by grooves parallelly arranged

striate, perforate

#### **striato-reticulate**

ornamentation intermediate between striate and reticulate

#### **suprasculpture**

secondary sculpture elements positioned on the primary sculpture of the pollen surface

*Anthurium gracile*, Araceae nano- to microechinate

*Triglochin maritima*, Juncaginaceae

nanoechinate

*Lophophora williamsii,* Cactaceae microechinate

*Anthurium gracile*, Araceae nano- to microechinate

*Ascarina lucida*, Chloranthaceae nanogemmate

*Phillyrea angustifolia*, Oleaceae microrugulate


*Parentucellia latifolia,* Orobanchaceae microgemmate


*Linum bienne*, Linaceae micro- to nanorugulate, nanoechinate

*Betula* sp., Betulaceae fossil, Miocene, China, nanogemmate to nanoechinate

*Ulmus* sp.*,* Ulmaceae fossil, Miocene, China, granulate *Saxifraga hostii*, Saxifragaceae microgemmate

*Piper auritum*, Piperaceae microrugulate

*Tsuga* sp., Pinaceae fossil, Miocene, China, echinate to baculate

*Ceratostigma plumbaginoides*, Plumbaginaceae micro- to nanorugulate, granulate

*Akebia quinata*, Lardizabalaceae microrugulate to rugulate

*Ouratea hexasperma,* Ochnaceae nanogemmate

*Ilex* sp., Aquifoliaceae fossil, Miocene, China, microrugulate

*Polemonium caeruleum*, Polemoniaceae microgemmate

*Coffea arabica*, Rubiaceae

rugulate to microrugulate

#### **supratectal element**

sculpture element positioned on top of the tectum


*Quercus robur,* Fagaceae echini on tectum

echini on tectum

*Atriplex sagittata*, Amaranthaceae

#### **verruca/verrucate**

wart-like element broader than high

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The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

### **Pollen Wall**

**tectum/tectate – 380 eutectate – 381 semitectate – 382 atectate – 384 infratectum alveolate – 385 infratectum columellate – 386 infratectum granular – 389 infratectum absent – 390 internal tectum – 391 foot layer – 393 endexine compact-continuous – 394 endexine compact-discontinuous – 395 endexine spongy-continuous – 396 endexine lamellar-continuous – 397 endexine in aperture only – 398 endexine absent – 399 intine, ektintine, endintine – 400 primexine – 403 sexine, nexine – 404 calymmate – 405 acalymmate – 406 arcus/arcuate – 407 tenuitas (tenuitates) – 408 pollen coating, pollenkitt – 411 pollen coating, primexine matrix – 414 pollen coating, tryphine – 415 elastoviscin – 416 viscin thread – 418 Ubisch body (Ubisch bodies) – 421**

379

#### **tectum/tectate**

outer more or less continuous ektexine layer; tectum condition can be eutectate or semitectate


#### **eutectate**

pollen grain with a predominantly continuous tectum


Thiéry test

*Apium nodiflorum*, Apiaceae

#### **semitectate**

pollen grain with a discontinuous tectum

*Fraxinus excelsior*, Oleaceae U+Pb

*Odontites luteus*, Orobanchaceae modified Thiéry test

*Odontites luteus*, Orobanchaceae

*Fraxinus excelsior*, Oleaceae reticulate, heterobrochate

microreticulate

*Quesnelia lateralis*, Bromeliaceae reticulate

*Pachysandra terminalis*, Buxaceae reticulum cristatum

#### **atectate**

pollen grain lacking a tectum

#### **infratectum alveolate**

infratectum with compartments of irregular size and shape

#### **infratectum columellate**

infratectum with columellae

	- Asteraceae fractured pollen wall, acetolyzed

388 ILLUSTRATED POLLEN TERMS

#### **infratectum granular**

infratectum composed of granula, cluster of granula or elements of different size and shape

*Viola tricolor*, Violaceae

U+Pb

*Ulmus laevis*, Ulmaceae U+Pb


*Plantago lanceolata*, Plantaginaceae modified Thiéry test

#### **infratectum absent**

*Dieffenbachia humilis*, Araceae U+Pb

*Berberis vulgaris*, Berberidaceae modified Thiéry test

*Mahonia aquifolium*, Berberidaceae U+Pb

*Iris pumila*, Iridaceae modified Thiéry test

#### **internal tectum**

additional more or less continuous layer within the infratectum

*Argyranthemum* sp., Asteraceae

internal tectum colored, U+Pb

*Cyanus segetum*, Asteraceae


#### **foot layer**

inner layer of an ektexine that can be continuous, discontinuous, perforated or absent


#### **endexine compact-continuous**

distinct exine layer between ektexine and intine


#### **endexine compact-discontinuous**

#### **endexine spongy-continuous**

#### **endexine lamellar-continuous**


*Thalictrum flavum*, Ranunculaceae endexine (E), modified Thiéry test

#### **endexine in aperture only**


#### **endexine absent**

	- *Avena sativa*, Poaceae modified Thiéry test

#### **intine, ektintine, endintine**

intine: part of the pollen wall next to the cytoplasm, can be monolayered or bilayered (ektintine and endintine)


400 ILLUSTRATED POLLEN TERMS

*Apium nodiflorum*, Apiaceae ektintine (e-transparent), endintine (e-dense), Thiéry test

	- intine (polysaccharides) stain electron dense with Thiéry test

ektintine (outer layer) and endintine (inner layer) of a bilayered intine

*Aristolochia arborea*, Aristolochiaceae intine (asterisk), ektintine (e-dense), endintine (e-transparent), modified Thiéry test

*Pachypodium succulentum*, Apocynaceae channelled ektintine, modified Thiéry test

*Apoballis acuminatissima*, Araceae

bilayered intine, potassium permanganate


*Apoballis acuminatissima*, Araceae bilayered intine clearly visible with modified Thiéry test polysaccharidic layer formed during early developmental stage wherein the later exine structures are preformed


U+Pb

*Nigella arvensis*, Ranunculaceae

primexine matrix with first exine elements (arrowhead), U+Pb

#### **sexine, nexine**

terms used for light microscopy, describing the outer and inner layers of the exine

#### **calymmate**

units covered by a continuous exine envelope

#### **acalymmate**

units covered by an exine envelope which is discontinuous at the junctions between monads

#### **arcus/arcuate**

a curved wall thickening interconnecting apertures

#### **tenuitas**

general term for a thinning of the pollen wall

*Tulipa kaufmanniana*, Liliaceae sulcate with 2 tenuitates proximally, debatable trisulcate


#### **pollen coating, pollenkitt**

pollen coating consisting of sticky substances, mainly lipids

	- *Anemone ranunculoides*, Ranunculaceae modified Thiéry test

*Salix x fragilis*, Salicaceae modified Thiéry test

*Syringa vulgaris*, Oleaceae lipid test


*Scilla bifolia*, Asparagaceae lipid test

#### **pollen coating, primexine matrix**

pollen coating consisting of primexine remnants in mature pollen grains

*Apium nodiflorum*, Apiaceae primexine matrix (arrowhead), Thiéry test

*Veronica spicata*, Plantaginaceae modified Thiéry test

*Atriplex tatarica*, Amaranthaceae modified Thiéry test

*Convolvulus tricolor*, Convolvulaceae primexine matrix (arrowhead) modified Thiéry test

*Sambucus nigra*, Adoxaceae Thiéry test

*Chenopodium album*, Amaranthaceae modified Thiéry test

#### **pollen coating, tryphine**

pollen coating consisting mainly of lipids mixed with membrane remnants

*Brassica nigra*, Brassicaceae modified Thiéry test

*Brassica napus*, Brassicaceae modified Thiéry test


*Alliaria petiolata*, Brassicaceae tryphine (asterisk), endexine (E), intine (I), U+Pb

#### **elastoviscin**

highly elastic, not acetolysis resistant substance in Orchidaceae, which interconnects the subunits (monads, tetrads or massulae) of a pollinium and builds up the caudicles

#### **viscin thread**

acetolysis resistant thread arising from the exine

#### **Ubisch body (Ubisch bodies)**

polymorphic sporopollenin-element produced by the tapetum

*Chamaecyparis lawsoniana*, Cupressaceae Ubisch bodies on locular wall

*Gladiolus illyricus*, Iridaceae *Quercus robur*, Fagaceae

*Ruspolia seticalyx*, Acanthaceae Ubisch bodies forming a reticulum

*Chamaecyparis lawsoniana*, Cupressaceae Ubisch bodies attached to pollen surface

*Ruspolia seticalyx*, Acanthaceae U+Pb

*Acacia binervia*, Mimosaceae *Justicia brandegeeana*, Acanthaceae


*Actaea spicata*, Ranunculaceae U+Pb

*Urtica dioica*, Urticaceae modified Thiéry test

*Quercus robur*, Fagaceae lipid test

*Eranthis hyemalis*, Ranunculaceae U+Pb, Ubisch body (star)

*Ulmus laevis,* Ulmaceae modified Thiéry test

*Plantago major*, Plantaginaceae U+Pb

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The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

### **Pollen Class**

**pollen class – 430**

429

#### **pollen class**

artificial grouping of pollen grains that share one or more distinctive characters

lophate

lophate

ILLUSTRATED POLLEN TERMS 431

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The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# **Palynological Terms**

#### **Contents**

**Glossary of Palynological Terms – 439**

### **Glossary of Palynological Terms**

All **important terms** in palynology are listed here and explained. Terms figured in the chapters of the "Illustrated Pollen Terms" part are indicated by bold page numbers. **Non-recommended terms** are only provided with an explanatory comment. For consistency, phrases are standardized, for example, features of ornamentation are stereotypically defined as "**pollen wall with** …", and pollen wall features (or pollen shape and size) as "**pollen grain with** …".

Both the singular and the plural are given for Latin terms. The English spelling of the Latin term is added (porus, pl. pori, engl. pore) if it is preferable. Cross-references are given to terms that are **synonyms** (the preferable one is printed in bold) or that indicate the opposite condition (**antonyms**), e.g., homo- and heterobrochate. If both a Greek and a corresponding Latin form exist for a prefix, then the Greek form is used consistently: panto- (not peri-), ekto- (not ecto-), or the Greek di- (dis-), and not the Latin bi- (bis-). There are few exceptions from this rule. If the Latin form is more widely accepted, then the term is used as *nomen conservandum*, for example, bisaccate is found exclusively in the literature and not the Greek form disaccate. Sometimes two adjectival variants (-ate, -ar) are used, but in two different meanings. For example, from the noun granulum derive two adjectival forms: granular and granulate (both meaning "with granules"). These are corresponding terms used in two quite different contexts: **granular** describes a distinct type of infratectum hence a structural feature whereas **granulate** refers to an ornamentation feature—a sculpture element.

Terms not listed in the glossary belong to spores, or are considered as redundant (e.g., multiplanar tetrad), superfluous (e.g., polyplicate, because plicate pollen grains are always equipped with several to many plicae), or may be a permanent source of confusion (zon-, zona-, zoni-, zono-).

439




the study of palynomorphs found in ice










pole of a pollen grain






**structure** 45


**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

448 PALYNOLOGICAL TERMS

**Picture Copyrights – 450**

**Index – 483**

# **Picture Copyrights**

### **Part 1 General Chapters**

#### **Chapter "Palynology: History and Systematic Aspects"**

Figure 1: Hyde and Williams (1944) Figure 2: Grew (1682) Figures 3–4: Fritzsche (1837) Figure 5: *PalDat* Figure 6, all pictures: Zetter, Reinhard Figures 7–10: *PalDat*

### **Chapter "Pollen Development"**

Figure 1: Illustration by Buchner, Ralf & Halbritter, Heidemarie Figure 2: *PalDat* Figure 3: Weber, Martina Figures 4–10: *PalDat* Figure 11: Ulrich Silvia (pictures A–D), Weber, Martina (pictures E–F)

#### **Chapter "Pollen Morphology and Ultrastructure"**


Figure 15, picture A: Illustration by Buchner, Ralf & Halbritter, Heidemarie, pictures B–C: *PalDat* Figure 16, pictures A, D: Illustration by Buchner, Ralf & Halbritter, Heidemarie, pictures B, C, E, F: *PalDat* Figure 17: Weber, Martina Figure 18: Zetter, Reinhard Figures 19–20: Zetter, Reinhard Figures 21–24: *PalDat* Figure 25, A, C, F: Weber, Martina; pictures B, D, E, G: *PalDat* Figures 26–30: *PalDat*

#### **Chapter "Misinterpretations in Palynology"**

Figure 1, picture A–B: *PalDat* C: Zetter, Reinhard Figures 2–4: *PalDat* Figure 5: Zetter, Reinhard Figures 6–7: *PalDat* Figure 8, pictures A–B: Zetter, Reinhard, picuture C: *PalDat* Figure 9: *PalDat* Figure 10, pictures A–C: Grímsson, Friðgeir, D: *PalDat*, E: Weber, Martina Figure 11: Zetter, Reinhard Figures 12–14: *PalDat* Figure 15, picture A: Weber, Martina, pictures B–C: *PalDat* Figure 16: *PalDat* Figure 17, picture A. Erdtman (1952), picture B. Punt et al. 2007, picture C. Weber, Martina, pictures D–E: *PalDat* Figure 18: Svojtka, Matthias Figure 19: Ulrich, Silvia Figure 20: *PalDat* Figure 21: Ulrich, Silvia Figure 22: *PalDat*

#### **Chapter "How to Describe and Illustrate Pollen Grains"**

Figures 1–2: Weber Martina & Ulrich Silvia Figures 3–9 Table 1: Grímsson, Friðgeir & Zetter, Reinhard

#### **Chapter "Methods in Palynology"**

Figure 1: Grímsson, Friðgeir & Zetter, Reinhard Figures 2–5: Ulrich, Silvia

Figures 6–8: Weber, Martina

**Service Part** Figure 9: Grímsson, Friðgeir & Zetter, Reinhard Figure 10: Ulrich, Silvia & Halbritter, Heidemarie Figures 11–12: Weber Martina & Ulrich Silvia Figure 13, pictures A–J, L–M: Weber Martina &

Ulrich Silvia, picture K: Böhmdorfer, Gudrun Figures 14–15: Weber Martina & Ulrich Silvia

Figure 16: Weber, Martina

Figures 17–22: Weber Martina & Ulrich Silvia

Figures 23–27: Grímsson, Friðgeir & Zetter, Reinhard Figure 28: Weber, Martina & Ulrich, Silvia

#### **Part 2 Illustrated Pollen Terms**

#### **Chapter: "Pollen- and Dispersal Units"**

Picture: *Alocasia odora*, page 132: Ulrich, Silvia Picture: *Tilia tomentosa*, page 132: *PalDat* Picture: *Betula humilis*, page 132: *PalDat* Picture: *Lathyrus pratensis*, page 132: *PalDat* Picture: *Tragopogon orientalis*, page 132: Weber, Martina

© Springer International Publishing AG 2018 Picture: *Gonatopus bovinii*, page 132: Ulrich, Silvia Picture: *Haworthia herbacea*, page 133: *PalDat* Picture: *Pinus strobus*, page 133: *PalDat* Picture: *Tragopogon orientalis*, page 133: *PalDat* Picture: *Bituminaria bituminosa*, page 133: *PalDat* Picture: *Lamium galeobdolon*, page 133: *PalDat* Picture: *Dactylis glomerata*, page 133: *PalDat* Picture: *Polypleurum stylosum*, page 134: *PalDat* Picture: *Polypleurum stylosum*, page 134: *PalDat* Picture: *Zeylanidium olivaceum*, page 134: *PalDat* Picture: *Zeylanidium subulatum*, page 134: *PalDat* Picture: *Thelethylax minutiflora*, page 134: *PalDat* Picture: *Scheuchzeria palustris*, page 134: *PalDat* Picture: *Carex* sp., page 135: *PalDat* Picture: *Carex atrata*, page 135: *PalDat* Picture: *Schoenoplectus lacustris*, page 135: *PalDat* Picture: *Scirpus sylvaticus*, page 135: *PalDat* Picture: *Carex distans*, page 135: *PalDat* Picture: *Cyperus conglomeratus*, page 135: *PalDat* Picture: *Carex digitata*, page 136: Weber, Martina Picture: *Trichophorum cespitosum*, page 136: *PalDat* Picture: *Cyperus diffusus*, page 136: *PalDat* Picture: *Scirpoides holoschoenus*, page 136: *PalDat* Picture: *Eleocharis palustris*, page 136: *PalDat* Picture: *Cyperus papyrus*, page 136: *PalDat* Picture: *Erica herbacea*, page 137: *PalDat* Picture: *Chlorospatha kolbii*, page 137: *PalDat* Picture: *Epipactis helleborine*, page 137: *PalDat* Picture: *Cyprinia gracilis*, page 137: *PalDat* Picture: *Epidendrum centropetalum*, page 137: *PalDat*

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8

Picture: *Chlorospatha ceronii*, page 137: *PalDat* Picture: Ericaceae, page 138: Grímsson, Friðgeir Picture: *Epilobium hirsutum*, page 138: Weber, Martina

Picture: *Drosera scorpioides*, page 138: Weber, Martina

Picture: *Erica tetralix*, page 138: Weber, Martina Picture: *Vaccinium myrtillus*, page 138: Weber, Martina

Picture: *Andromeda polifolia*, page 138: Weber, Martina

Picture: *Rhododendron hirsutum*, page 139: *PalDat* Picture: *Victoria regia*, page 139: *PalDat*

Picture: *Luzula campestris*, page 139: *PalDat*

Picture: *Drimys granatensis*, page 139: *PalDat*

Picture: *Epilobium montanum*, page 139: *PalDat* Picture: *Oxyanthus subpunctatus*, page 139:

*PalDat*

Picture: *Epilobium parviflorum*, page 140: *PalDat* Picture: *Arbutus unedo*, page 140: *PalDat* Picture: *Chelonanthus alatus*, page 140: *PalDat* Picture: *Mimosa pudica*, page 140: *PalDat* Picture: *Ludwigia octovalvis*, page 140: *PalDat* Picture: *Agapetes macrantha*, page 140: *PalDat* Picture: *Dionaea muscipula*, page 141: *PalDat* Picture: *Juncus effusus*, page 141: *PalDat* Picture: *Moneses uniflora*, page 141: *PalDat* Picture: *Drosera binata*, page 141: *PalDat* Picture: *Vaccinium vitis-idaea*, page 141: *PalDat* Picture: *Calolisianthus pendulus*, page 141: *PalDat* Picture: *Calla palustris*, page 142: Ulrich, Silvia Picture: *Androlepis skinneri*, page 142: *PalDat* Picture: *Calluna vulgaris*, page 142: *PalDat* Picture: *Chlorospatha dodsonii*, page 142: *PalDat* Picture: *Rhodothamnus chamaecistus*, page 142: *PalDat* Picture: *Catalpa bungei*, page 142: *PalDat* Picture: *Typha latifolia*, page 143: Zetter, Reinhard Picture: *Typha latifolia*, page 143: *PalDat* Picture: *Typha latifolia*, page 143: Zetter, Reinhard Picture: *Typha latifolia*, page 143: *PalDat* Picture: *Typha latifolia*, page 143: Zetter, Reinhard

Picture: *Typha latifolia*, page 143: *PalDat* Picture: *Asimina triloba*, page 144: Weber, Martina Picture: *Calluna vulgaris*, page 144: Weber,

#### Martina

Picture: *Chlorospatha kolbii*, page 144: *PalDat* Picture: *Chlorospatha dodsonii*, page 144: *PalDat* Picture: *Cyprinia gracilis*, page 144: *PalDat* Picture: *Drosera peltata*, page 144: *PalDat* Picture: *Acacia myrtifolia*, page 145: *PalDat* Picture: *Calliandra tergemina*, page 145: *PalDat* Picture: *Pithecellobium dulce*, page 145: *PalDat* Picture: *Acacia* sp.page 145: Weber, Martina Picture: *Cymbopetalum aequale*, page 145: *PalDat*

Picture: *Chelonanthus purpurascens*, page 145: *PalDat* Picture: *Acacia* sp.page 146: *PalDat* Picture: *Acacia dealbata*, page 146: *PalDat* Picture: *Acacia karroo*, page 146: *PalDat* Picture: *Acacia karroo*, page 146: *PalDat* Picture: *Albizia julibrissin*, page 146: *PalDat* Picture: *Albizia saman*, page 146: *PalDat* Picture: *Traunsteinera globosa*, page 147: *PalDat* Picture: *Epipogium aphyllum*, page 147: *PalDat* Picture: *Herminium monorchis*, page 147: *PalDat* Picture: *Gennaria diphylla*, page 147: Svojtka, Matthias Picture: *Orchis italica*, page 147: Svojtka, Matthias Picture: *Orchis purpurea*, page 147: Svojtka, Matthias Picture: *Cephalanthera longifolia*, page 148: Svojtka, Matthias Picture: *Cephalanthera longifolia*, page 148: Svojtka, Matthias Picture: *Dendrobium farmeri*, page 148: *PalDat* Picture: *Dendrobium farmeri*, page 148: *PalDat* Picture: *Steveniella satyrioides*, page 148: *PalDat* Picture: *Steveniella satyrioides*, page 148: *PalDat* Picture: *Hammarbya paludosa*, page 149: *PalDat* Picture: *Plectrophora cultrifolia*, page 149: *PalDat* Picture: *Restrepia muscifera*, page 149: *PalDat* Picture: *Malaxis monophyllos*, page 149: *PalDat* Picture: *Stephanotis floribunda*, page 149: *PalDat* Picture: *Hoodia flava*, page 149: *PalDat* Picture: *Spiranthes spiralis*, page 150: *PalDat* Picture: *Caladenia latifolia*, page 150: *PalDat* Picture: *Ornithocephalus myrticola*, page 150: *PalDat* Picture: *Aerides multiflora*, page 150: *PalDat* Picture: *Maxillaria densa*, page 150: *PalDat* Picture: *Brassavola cucullata*, page 150: Svojtka, Matthias Picture: *Coelogyne fimbriata*, page 151: *PalDat* Picture: *Pleurothallis loranthophylla*, page 151: *PalDat* Picture: *Oncidium maizaefolium*, page 151: *PalDat* Picture: *Schoenorchis fragrans*, page 151: Svojtka, Matthias Picture: *Haraella odorata*, page 151: Svojtka, Matthias Picture: *Stanhopea oculata*, page 151: *PalDat* Picture: *Herminium monorchis*, page 152: *PalDat* Picture: *Goodyera repens*, page 152: *PalDat* Picture: *Gennaria diphylla*, page 152: *PalDat* Picture: *Steveniella satyrioides*, page 152: *PalDat* Picture: *Ludisia discolor*, page 152: *PalDat* Picture: *Anacamptis pyramidalis*, page 152: *PalDat* Picture: *Stephanotis floribunda*, page 153: *PalDat* Picture: *Ceropegia sandersonii*, page 153: *PalDat* Picture: *Hoya carnosa*, page 153: *PalDat* Picture: *Hoya multiflora*, page 153: *PalDat*

Picture: *Frerea indica*, page 153: *PalDat* Picture: *Orbeanthus hardyi*, page 153: *PalDat*

#### **Chapter: "Shape and Polarity"**

Picture: *Ligustrum* sp., page: 156: Zetter, Reinhard Picture: Chenopodiaceae, page: 156: Weber, Martina

Picture: *Hedyosmum scaberrimum*, page: 156: *PalDat*

Picture: *Anthurium ovatifolium*, page: 156: *PalDat* Picture: *Abutilon theophrasti*, page: 156: *PalDat* Picture: *Corydalis ophiocarpa*, page: 156: *PalDat* Picture: *Mayna odorata*, page: 157: *PalDat* Picture: *Saruma henryi*, page: 157: *PalDat* Picture: *Phleum pratense*, page: 157: *PalDat* Picture: *Fraxinus ornus*, page: 157: *PalDat* Picture: *Galium lucidum*, page: 157: *PalDat* Picture: *Ginkgo biloba*, page: 157: *PalDat*

Picture: *Impatiens parviflora*, page: 158: Weber, Martina

Picture: *Impatiens parviflora*, page: 158: *PalDat* Picture: *Amorphophallus interruptus*, page: 158: Ulrich, Silvia

Picture: *Allium oleraceum*, page: 158: *PalDat* Picture: *Ambrosina bassi*, page: 158: Ulrich, Silvia Picture: *Aechmea dealbata*, page: 158: *PalDat* Picture: *Salvia coccinea*, page: 159: *PalDat* Picture: *Commelina erecta*, page: 159: *PalDat* Picture: *Billbergia porteana*, page: 159: *PalDat* Picture: *Zamia loddigesii*, page: 159: *PalDat* Picture: *Galeopsis tetrahit*, page: 159: *PalDat* Picture: *Physostegia virginiana*, page: 159: *PalDat* Picture: *Acer pseudoplatanus*, page: 160: *PalDat* Picture: *Artemisia pontica*, page: 160: *PalDat* Picture: *Sanguisorba officinalis*, page: 160: *PalDat* Picture: *Orthilia secunda*, page: 160: *PalDat* Picture: *Gunnera tinctoria*, page: 160: *PalDat* Picture: *Gunnera tinctoria*, page: 160: *PalDat* Picture: *Sedum rupestre*, page: 161: *PalDat* Picture: *Viola alba*, page: 161: *PalDat* Picture: *Clematis heracleifolia*, page: 161: *PalDat* Picture: *Sanicula europaea*, page: 161: *PalDat* Picture: *Pinguicula ehlersiae*, page: 161: *PalDat* Picture: *Bellis perennis*, page: 161: *PalDat* Picture: *Artemisia* sp.page: 162: Weber, Martina Picture: *Nicotiana tabacum*, page: 162: Weber, Martina Picture: *Hypecoum imberbe*, page: 162: *PalDat* Picture: *Barringtonia asiatica*, page: 162: *PalDat* Picture: *Pelargonium punctatum*, page: 162: *PalDat* Picture: *Viola riviniana*, page: 162: *PalDat* Picture: *Lopezia racemosa*, page: 163: Weber, Martina

Picture: *Loranthus europaeus*, page: 163: *PalDat*

**Service Part** © Springer International Publishing AG 2018 Picture: *Macadamia ternifolia*, page: 163: Weber, Martina Picture: *Tropaeolum emarginatum*, page: 163: *PalDat* Picture: *Kolkwitzia amabilis*, page: 163: Weber, Martina Picture: *Acicarpha tribuloides*, page: 163: *PalDat* Picture: *Callistemon coccineus*, page: 164: *PalDat* Picture: *Hypoestes phyllostachya*, page: 164: *PalDat* Picture: *Echinops ritro*, page: 164: *PalDat* Picture: *Bupleurum rotundifolium*, page: 164: *PalDat* Picture: *Paullinia tomentosa*, page: 164: *PalDat* Picture: *Primula denticulata*, page: 164: *PalDat* Picture: *Orlaya grandiflora*, page: 165: *PalDat* Picture: *Circaea lutetiana*, page: 165: *PalDat* Picture: *Sempervivum globiferum*, page: 165: *PalDat* Picture: *Cunonia capensis*, page: 165: *PalDat* Picture: *Dipsacus fullonum*, page: 165: *PalDat* Picture: *Potentilla inclinata*, page: 165: *PalDat* Picture: *Anchusa officinalis*, page: 166: *PalDat* Picture: *Nonea pulla*, page: 166: *PalDat* Picture: *Viola tricolor*, page: 166: *PalDat* Picture: *Eremurus robustus*, page: 166: *PalDat* Picture: *Herniaria glabra*, page: 166: *PalDat* Picture: *Sideritis romana*, page: 166: *PalDat* Picture: *Viola arvensis*, page: 167: Weber, Martina Picture: *Viola arvensis*, page: 167: PalDat Picture: *Opuntia basilaris*, page: 167: *PalDat* Picture: *Talinum paniculatum*, page: 167: *PalDat* Picture: *Sarracenia alata*, page: 167: *PalDat* Picture: *Stellaria holostea*, page: 167: *PalDat* Picture: *Carya* sp., page: 168: Zetter, Reinhard Picture: *Corylus avellana*, page: 168: *PalDat* Picture: *Plectranthus esculentus*, page: 168: Weber, Martina Picture: *Salvia argentea*, page: 168: *PalDat* Picture: *Impatiens* sp., page: 168: Weber, Martina Picture: *Impatiens glandulifera*, page: 168: *PalDat* Picture: *Sarracenia alata*, page: 169: *PalDat* Picture: *Hymenocallis tubiflora*, page: 169: *PalDat* Picture: *Knautia drymeia*, page: 169: *PalDat* Picture: *Cuphea procumbens*, page: 169: *PalDat* Picture: *Hakea kippistiana*, page: 169: *PalDat* Picture: *Roridula gorgonias*, page: 169: *PalDat* Picture: *Clarkia unguiculata*, page: 170: *PalDat* Picture: *Vriesea neoglutinosa*, page: 170: *PalDat* Picture: *Acicarpha tribuloides*, page: 170: *PalDat* Picture: *Heliconia* sp., page: 170: *PalDat* Picture: *Amsonia ciliata*, page: 170: *PalDat* Picture: *Clarkia purpurea*, page: 170: *PalDat* Picture: *Parnassia palustris*, page: 171: *PalDat* Picture: *Campanula fenestrellata*, page: 171: *PalDat* Picture: *Roemeria hybrida*, page: 171: *PalDat*

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8

*PalDat* Picture: *Whitfieldia lateritia*, page: 172: *PalDat* Picture: *Whitfieldia lateritia*, page: 172: *PalDat* Picture: *Schoepfia schreberi*, page: 172: *PalDat* Picture: *Thesium arvense*, page: 172: *PalDat* Picture: *Pedicularis gyroflexa*, page: 172: *PalDat* Picture: *Basella alba*, page: 172: *PalDat* Picture: *Justicia carnea*, page: 173: Weber, Martina Picture: *Aphelandra arborea*, page: 173: *PalDat* Picture: *Aesculus* sp., page: 173: Weber, Martina Picture: *Lysimachia lichiangensis*, page: 173: *PalDat* Picture: *Colutea arborescens*, page: 173: Weber, Martina Picture: *Oxytropis jacquinii*, page: 173: *PalDat* Picture: *Crossandra flava*, page: 174: *PalDat* Picture: *Jurinea mollis*, page: 174: *PalDat* Picture: *Torilis arvensis*, page: 174: *PalDat* Picture: *Peucedanum cervaria*, page: 174: *PalDat* Picture: *Astragalus onobrychis*, page: 174: *PalDat* Picture: *Symphytum officinale*, page: 174: *PalDat* Picture: *Buglossoides purpurocaerulea*, page: 175: *PalDat* Picture: *Vitaliana primuliflora*, page: 175: *PalDat* Picture: *Platycodon grandiflorus*, page: 175: *PalDat* Picture: *Stenandrium guineense*, page: 175: *PalDat* Picture: *Lathyrus tuberosus*, page: 175: *PalDat* Picture: *Salvia sclarea*, page: 175: *PalDat* Picture: Apiaceae, page: 176: Weber, Martina Picture: *Bifora radians*, page: 176: *PalDat* Picture: *Columnea magnifica*, page: 176: *PalDat* Picture: *Asperula tinctoria*, page: 176: *PalDat* Picture: *Monotropa hypopitys*, page: 176: *PalDat* Picture: *Myosotis scorpioides*, page: 176: *PalDat* Picture: *Viburnum tinus*, page: 177: *PalDat* Picture: *Cerinthe minor*, page: 177: *PalDat* Picture: *Luffa cylindrica*, page: 177: *PalDat* Picture: *Crossandra flava*, page: 177: *PalDat* Picture: *Pulmonaria angustifolia*, page: 177: *PalDat* Picture: *Aesculus flava*, page: 177: *PalDat* Picture: *Onosma visianii*, page: 178: *PalDat* Picture: *Billbergia seidelii*, page: 178: *PalDat* Picture: *Chaenorhinum minus*, page: 178: *PalDat* Picture: *Limnanthes douglasii*, page: 178: *PalDat* Picture: *Sesleria albicans*, page: 178: *PalDat*

Picture: *Silene nutans*, page: 171: *PalDat* Picture: *Iris pumila*, page: 171: *PalDat*

Picture: *Sarcocapnos enneaphylla*, page: 171:

Picture: *Elaeagnus angustifolia*, page: 178: *PalDat* Picture: *Heliconia* sp., page: 179: *PalDat* Picture: *Quesnelia augusto-coburgii*, page: 179: *PalDat*

Picture: *Erica arborea*, page: 179: *PalDat* Picture: *Pinus strobus*, page: 179: *PalDat* Picture: *Nuphar lutea*, page: 179: *PalDat* Picture: *Sansevieria parva*, page: 179: *PalDat* Picture: *Amorphophallus yunnanensis*, page: 180: Ulrich, Silvia

Picture: *Austrobaileya scandens*, page: 180: *PalDat* Picture: *Echium italicum*, page: 180: *PalDat* Picture: *Adenanthos sericeus*, page: 180: *PalDat* Picture: *Calluna vulgaris*, page: 180: *PalDat* Picture: *Alkanna corcyrensis*, page: 180: *PalDat* Picture: *Adansonia gregorii*, page: 181: *PalDat* Picture: *Ruellia macrantha*, page: 181: *PalDat* Picture: *Cirsium oleraceum*, page: 181: *PalDat* Picture: *Phlox paniculata*, page: 181: *PalDat* Picture: *Crossandra flava*, page: 181: *PalDat* Picture: *Justicia carnea*, page: 181: *PalDat* Picture: *Basella alba*, page: 182: *PalDat* Picture: *Herniaria glabra*, page: 182: *PalDat* Picture: *Cerastium dubium*, page: 182: *PalDat* Picture: *Eremogone procera*, page: 182: *PalDat* Picture: *Paronychia polygonifolia*, page: 182:

*PalDat*

Picture: *Paronychia polygonifolia*, page: 182: *PalDat*

Picture: *Alkanna corcyrensis*, page: 183: *PalDat*

Picture: *Montrichardia arborescens*, page: 183: *PalDat*

Picture: *Heliconia rostrata*, page: 183: *PalDat* Picture: *Echium plantagineum*, page: 183: *PalDat*

Picture: *Cyperus longus*, page: 183: *PalDat*

Picture: *Ephedra foeminea*, page: 183: *PalDat*

Picture: *Brugmansia suaveolens*, page: 184: *PalDat*

Picture: *Anthyllis vulneraria*, page: 184: *PalDat*

Picture: *Corydalis cheilanthifolia*, page: 184: *PalDat*

Picture: *Schoepfia schreberi*, page: 184: *PalDat*

Picture: *Aechmea drakeana*, page: 184: *PalDat*

Picture: *Cardiospermum halicacabum*, page: 184: *PalDat*

Picture: *Whitfieldia lateritia*, page: 185: *PalDat*

Picture: *Pedicularis portenschlagii*, page: 185: *PalDat*

Picture: *Billbergia pyramidalis*, page: 185: *PalDat* Picture: *Quesnelia imbricata*, page: 185: *PalDat* Picture: *Clarkia unguiculata*, page: 185: *PalDat* Picture: *Fuchsia paniculata*, page: 185: *PalDat* Picture: *Acicarpha tribuloides*, page: 186: *PalDat* Picture: *Nicandra physalodes*, page: 186: *PalDat* Picture: *Myosotis alpestris*, page: 186: *PalDat* Picture: *Loranthus europaeus*, page: 186: *PalDat* Picture: *Sansevieria suffruticosa*, page: 186: *PalDat* Picture: *Juncus jacquinii*, page: 186: *PalDat* Picture: *Gaura lindheimeri*, page: 187: *PalDat* Picture: *Eremurus robustus*, page: 187: *PalDat* Picture: *Hyacinthoides italica*, page: 187: *PalDat* Picture: *Galanthus nivalis*, page: 187: *PalDat* Picture: *Limnanthes douglasii*, page: 187: *PalDat* Picture: *Thesium dollineri*, page: 187: *PalDat* Picture: *Picea* sp., page: 188: Grímsson, Friðgeir Picture: *Abies cephalonica*, page: 188: Weber, Martina

Picture: *Pinus cembra*, page: 188: Weber, Martina Picture: *Picea* sp., page: 188: Weber, Martina Picture: *Tsuga canadensis*, page: 188: Weber,

Martina

Picture: *Dacrycarpus dacrydioides*, page: 188: *PalDat*

Picture: *Tsuga* sp., page: 189: Zetter, Reinhard Picture: *Tsuga* sp., page: 189: Zetter, Reinhard Picture: *Tsuga canadensis*, page: 189: Weber, Martina

Picture: *Tsuga canadensis*, page: 189: *PalDat* Picture: *Tsuga* sp., page: 189: Grímsson, Friðgeir Picture: *Tsuga* sp., page: 189: Grímsson, Friðgeir Picture: *Picea* sp.page: 190: Grímsson, Friðgeir Picture: *Picea* sp.page: 190: Grímsson, Friðgeir Picture: *Picea* sp.page: 190: Grímsson, Friðgeir Picture: *Pinus* sp., page: 190: Grímsson, Friðgeir Picture: *Pinus* sp., page: 190: Grímsson, Friðgeir Picture: *Abies cephalonica*, page: 190: Weber, Martina

Picture: *Abies cephalonica*, page: 191: *PalDat* Picture: *Picea abies*, page: 191: *PalDat*

Picture: *Picea abies*, page: 191: *PalDat* Picture: *Pinus mugo*, page: 191: *PalDat*

Picture: *Abies nordmanniana*, page: 191: *PalDat*

Picture: *Picea pungens*, page: 191: *PalDat*

Picture: *Pinus heldreichii*, page: 192: *PalDat*

Picture: *Pinus nigra*, page: 192: *PalDat*

Picture: *Podocarpus* sp., page: 192: *PalDat*

Picture: *Podocarpus* sp., page: 192: *PalDat*

Picture: *Pinus mugo*, page: 192: Weber, Martina

Picture: *Pinus contorta*, page: 192: *PalDat*

Picture: Podocarpaceae, page: 193: Grímsson, Friðgeir

Picture: *Pherosphaera hookeriana*, page: 193: *PalDat*

Picture: *Pherosphaera hookeriana*, page: 193: *PalDat*

Picture: *Dacrycarpus dacrydioides*, page: 193: *PalDat*

Picture: *Abies concolor*, page: 193: *PalDat* Picture: *Abies concolor*, page: 193: *PalDat*

Picture: *Fritillaria pontica*, page: 194: Weber, Martina

Picture: *Veratrum album*, page: 194: *PalDat* Picture: *Galium odoratum*, page: 194: *PalDat* Picture: *Sparmannia africana*, page: 194: *PalDat* Picture: *Roemeria hybrida*, page: 194: *PalDat* Picture: *Bifora radians*, page: 194: *PalDat* Picture: *Artemisia pontica*, page: 195: *PalDat* Picture: *Carex alba*, page: 195: *PalDat* Picture: *Lachenalia aloides*, page: 195: *PalDat* Picture: *Luzula sylvatica*, page: 195: *PalDat* Picture: *Moehringia muscosa*, page: 195: *PalDat* Picture: *Anemone hortensis*, page: 195: *PalDat* Picture: *Dracontium asperum*, page: 196: Ulrich, Silvia

**Service Part** Picture: *Billbergia seidelii*, page: 196: *PalDat* Picture: *Lilium candidum*, page: 196: *PalDat* Picture: *Nuphar lutea*, page: 196: *PalDat* Picture: *Ginkgo biloba*, page: 196: *PalDat* Picture: *Asphodeline lutea*, page: 196: *PalDat* Picture: *Lysichiton americanus*, page: 197: *PalDat* Picture: *Piper nigrum*, page: 197: *PalDat* Picture: *Gagea lutea*, page: 197: *PalDat* Picture: *Sparganium erectum*, page: 197: *PalDat* Picture: *Dioon edule*, page: 197: *PalDat* Picture: *Symplocarpus foetidus*, page: 197: *PalDat* Picture: *Asphodelus fistulosus*, page: 198: *PalDat* Picture: *Eremurus thiodanthus*, page: 198: *PalDat* Picture: *Piper auritum*, page: 198: *PalDat* Picture: *Wachendorfia thyrsiflora*, page: 198: *PalDat* Picture: *Haemanthus coccineus*, page: 198: *PalDat* Picture: *Hymenocallis tubiflora*, page: 198: *PalDat* Picture: *Bougainvillea* sp., page: 199: *PalDat* Picture: *Heliconia* sp., page: 199: *PalDat* Picture: *Tilia euchlora*, page: 199: *PalDat* Picture: *Elaeagnus angustifolia*, page: 199: *PalDat* Picture: *Luzula campestris*, page: 199: *PalDat* Picture: *Tsuga canadensis*, page: 199: *PalDat* Picture: *Adenanthos sericeus*, page: 200: *PalDat* Picture: *Heliconia stricta*, page: 200: *PalDat* Picture: *Petrea volubilis*, page: 200: *PalDat* Picture: *Leucadendron brunioides*, page: 200:

*PalDat* Picture: *Cunninghamia lanceolata*, page: 200: *PalDat*

Picture: *Hibiscus schizopetalus*, page: 200: *PalDat*


Picture: *Leucadendron discolor*, page: 201: *PalDat* Picture: *Melastoma sanguineum*, page: 201: *PalDat*


Picture: *Grevillea banksii*, page: 202: *PalDat*

Picture: *Tropaeolum moritzianum*, page: 202: *PalDat*

© Springer International Publishing AG 2018 M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8 Picture: Poaceae, page: 203: Weber, Martina Picture: *Dorstenia contrajerva*, page: 203: *PalDat* Picture: *Guzmania elvallensis*, page: 203: *PalDat* Picture: *Smilax spinosa*, page: 203: *PalDat* Picture: *Callitriche stagnalis*, page: 203: *PalDat* Picture: *Vanilla planifolia*, page: 203: *PalDat* Picture: *Urtica dioica*, page: 204: *PalDat* Picture: *Populus alba*, page: 204: *PalDat* Picture: *Sesleria albicans*, page: 204: *PalDat* Picture: *Anthurium radicans*, page: 204: *PalDat*

Picture: *Coriaria nepalensis*, page: 204: *PalDat* Picture: *Orobanche hederae*, page: 204: *PalDat*

### **Chapter: "Aperture"**

Picture: *Symplocos* sp., page 208: Zetter, Reinhard Picture: *Acalypha macrostachya*, page 208: *PalDat*

Picture: *Tropaeolum majus*, page 208: *PalDat* Picture: *Anthyllis vulneraria*, page 208: *PalDat* Picture: *Cardiospermum halicacabum*, page 208: *PalDat*

Picture: *Oenothera biennis*, page 208: *PalDat* Picture: *Betula humilis*, page 209: *PalDat* Picture: *Triticum aestivum*, page 209: *PalDat* Picture: Poaceae, page 209: Weber, Martina Picture: *Secale cereale*, page 209: *PalDat* Picture: *Circaea lutetiana*, page 209: *PalDat* Picture: *Fumaria officinalis*, page 209: *PalDat* Picture: *Myriophyllum spicatum*, page 210: *PalDat* Picture: *Morina longifolia*, page 210: *PalDat* Picture: *Epilobium angustifolium*, page 210: *PalDat* Picture: *Dorstenia contrajerva*, page 210: *PalDat* Picture: *Trichosanthes cucumerina*, page 210:

*PalDat*


Picture: *Alnus glutinosa*, page 211: *PalDat*

Picture: *Tilia* sp., page 211: Weber, Martina

Picture: *Elaeagnus angustifolia*, page 211: Weber, Martina

Picture: *Rubus caesius*, page 211: Weber, Martina Picture: *Acinos alpinus*, page 211: *PalDat*

Picture: *Eupatorium cannabinum*, page 212: *PalDat*

Picture: *Clinopodium vulgare*, page 212: *PalDat* Picture: *Eucharis grandiflora*, page 212: *PalDat* Picture: *Alisma plantago-aquatica*, page 212: *PalDat*

	- Picture: *Petrea volubilis*, page 221: *PalDat*
	- Picture: *Succisa pratensis*, page 221: *PalDat*
	- Picture: *Tilia platyphyllos*, page 222: Weber, Martina
	- Picture: *Coriaria myrtifolia*, page 222: *PalDat*
	- Picture: *Dalechampia spathulata*, page 222: *PalDat*
	- Picture: *Symphytum orientale*, page 222: *PalDat*
	- Picture: *Borago pygmaea*, page 222: *PalDat*
	- Picture: *Tabernaemontana simulans*, page 222: *PalDat*

Picture: *Elaeagnus angustifolia*, page 223: *PalDat* Picture: *Malus baccata*, page 223: Weber, Martina Picture: *Bifora radians*, page 223: *PalDat* Picture: *Gazania rigens*, page 223: *PalDat* Picture: *Cunonia capensis*, page 223: *PalDat* Picture: *Colutea arborescens*, page 223: Weber, Martina

Picture: *Typha latifolia*, page 224: *PalDat* Picture: *Epilobium hirsutum*, page 224: *PalDat* Picture: *Ludwigia octovalvis*, page 224: *PalDat* Picture: *Thelethylax minutiflora*, page 224: *PalDat* Picture: *Aechmea subintegerrima*, page 224: *PalDat*

Picture: *Aechmea subintegerrima*, page 224: *PalDat*


Picture: *Impatiens parviflora*, page 229: Weber, Martina

Picture: *Impatiens glandulifera*, page 229: *PalDat* Picture: *Impatiens parviflora*, page 229: *PalDat* Picture: *Sideritis romana*, page 229: *PalDat* Picture: *Didymaea mexicana*, page 229: *PalDat* Picture: *Mendoncia albida*, page 229: *PalDat* Picture: *Cruciata laevipes*, page 230: *PalDat*


Picture: *Clinopodium vulgare*, page 230: *PalDat* Picture: *Galium glaucum*, page 231: *PalDat*

**Service Part** Picture: *Galium lucidum*, page 231: *PalDat* Picture: *Sherardia arvensis*, page 231: *PalDat* Picture: *Galium odoratum*, page 231: *PalDat* Picture: *Codonopsis pilosula*, page 231: *PalDat* Picture: *Sechium edule*, page 231: *PalDat* Picture: *Sarcocapnos enneaphylla*, page 232: *PalDat* Picture: *Talinum paniculatum*, page 232: *PalDat* Picture: *Pseudofumaria lutea*, page 232: *PalDat* Picture: *Maripa nicaraguensis*, page 232: *PalDat* Picture: *Mollugo verticillata*, page 232: *PalDat* Picture: *Turbinicarpus pseudomacrochele*, page 232: *PalDat* Picture: *Justicia procumbens*, page 233: *PalDat* Picture: *Justicia procumbens*, page 233: *PalDat* Picture: *Adhatoda schimperiana*, page 233: *PalDat* Picture: *Justicia macrantha*, page 233: *PalDat* Picture: *Justicia carnea*, page 233: Weber, Martina Picture: *Justicia xylosteoides*, page 233: *PalDat* Picture: *Lathyrus vernus*, page 234: *PalDat* Picture: *Kraussia floribunda*, page 234: *PalDat* Picture: *Hieracium hoppeanum*, page 234: *PalDat* Picture: *Erica herbacea*, page 234: *PalDat* Picture: *Rumex acetosa*, page 234: *PalDat* Picture: *Aruncus dioicus*, page 234: *PalDat* Picture: *Tricolporopollenites wackersdorfensis*, page 235: Zetter, Reinhard Picture: Fabaceae, page 235: Weber, Martina Picture: *Fagopyrum* sp., page 235: Weber, Martina Picture: *Lathyrus sylvestris*, page 235: Weber, Martina Picture: *Rhus* sp., page 235: Weber, Martina Picture: *Euphorbia peplus*, page 235: Weber, Martina Picture: *Centaurea scabiosa*, page 236: *PalDat* Picture: *Cirsium oleraceum*, page 236: *PalDat* Picture: *Echium vulgare*, page 236: *PalDat* Picture: *Gardenia thunbergia*, page 236: *PalDat* Picture: *Myrrhis odorata*, page 236: *PalDat* Picture: *Fatsia japonica*, page 236: *PalDat* Picture: *Citrus swinglei*, page 237: *PalDat* Picture: *Pulmonaria mollis*, page 237: *PalDat* Picture: *Nicotiana tabacum*, page 237: *PalDat* Picture: *Genlisea violacea*, page 237: *PalDat* Picture: *Poncirus trifoliata*, page 237: *PalDat* Picture: *Tridax procumbens*, page 237: *PalDat* Picture: *Viola arvensis*, page 238: *PalDat* Picture: *Justicia menesii*, page 238: *PalDat* Picture: *Sanguisorba officinalis*, page 238: *PalDat* Picture: *Cerinthe minor*, page 238: *PalDat* Picture: *Pinguicula ehlersiae*, page 238: *PalDat* Picture: *Polygala chamaebuxus*, page 238: *PalDat* Picture: *Moltkia petraea*, page 239: *PalDat* Picture: *Symphytum caucasicum*, page 239: *PalDat*

Picture: *Buglossoides arvensis*, page 239: *PalDat* Picture: *Echinopepon wrightii*, page 239: *PalDat* Picture: *Utricularia vulgaris*, page 239: Weber,

Martina Picture: *Lathyrus sylvestris*, page 240: Weber, Martina

Picture: *Tussilago farfara*, page 240: Weber, Martina

Picture: *Parthenocissus* sp., page 240: Weber, Martina

Picture: *Centaurea jacea*, page 240: Weber, Martina

Picture: *Cichorium intybus*, page 240: Weber, Martina

Picture: *Lysimachia punctata*, page 240: Weber, Martina

Picture: *Dictamnus albus*, page 241: Weber, Martina

Picture: *Scaevola aemula*, page 241: Weber, Martina

Picture: *Vicia faba*, page 241: Weber, Martina Picture: *Vitaliana primuliflora*, page 241: *PalDat* Picture: *Rumex* sp., page 241: Zetter, Reinhard Picture: *Phacelia campanularia*, page 241: *PalDat* Picture: *Lythrum hyssopifolia*, page 242: *PalDat* Picture: *Pardoglossum* sp., page 242: *PalDat* Picture: *Tetramerium nervosum*, page 242: *PalDat* Picture: *Cynoglossum officinale*, page 242: *PalDat* Picture: *Phacelia tanacetifolia*, page 242: *PalDat* Picture: *Myosotis ramosissima*, page 242: *PalDat* Picture: *Pseuderanthemum alatum*, page 243: *PalDat*

Picture: *Combretum fruticosum*, page 243: *PalDat* Picture: *Omphalodes linifolia*, page 243: *PalDat* Picture: *Medinilla scortechinii*, page 243: *PalDat* Picture: *Meriania selvaflorensis*, page 243: *PalDat* Picture: *Phacelia campanularia*, page 243: *PalDat* Picture: *Alocasia odora*, page 244: Ulrich, Silvia Picture: *Stylochaeton bogneri*, page 244: Ulrich, Silvia

Picture: *Ambrosina bassii*, page 244: Ulrich, Silvia Picture: *Synandrospadix vermitoxicus*, page 244: Ulrich, Silvia

Picture: *Spathicarpa sagittifolia*, page 244: Ulrich, Silvia

Picture: *Spathiphyllum* sp., page 244: Ulrich, Silvia Picture: *Pinellia ternata*, page 245: *PalDat*

Picture: *Populus alba*, page 245: *PalDat* Picture: *Chlorospatha dodsonii*, page 245: *PalDat*

Picture: *Aglaodorum griffithii*, page 245: *PalDat*

Picture: *Phoebe sheareri*, page 245: *PalDat*

Picture: *Posidonia* sp., page 245: *PalDat*

Picture: *Orchidantha maxillarioides*, page 246: *PalDat*

Picture: *Aristolochia arborea*, page 246: *PalDat* Picture: *Triglochin maritima*, page 246: *PalDat* Picture: *Gnetum gnemon*, page 246: *PalDat*

© Springer International Publishing AG 2018 M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8 Picture: *Polygala major*, page 239: *PalDat*

Picture: *Cytinus hypocistis*, page 246: *PalDat* Picture: *Trillium chloropetalum*, page 246: *PalDat* Picture: *Cedrus atlantica*, page 247: Weber,

Martina Picture: *Picea* sp., page 247: Ulrich, Silvia Picture: *Pinus strobus*, page 247: Weber, Martina

Picture: *Pinus cembra*, page 247: Weber, Martina Picture: *Picea* sp., page 247: Weber, Martina Picture: *Pinus cembra*, page 247: Weber, Martina

Picture: *Discocleidion rufescens*, page 248: *PalDat*

Picture: *Euphorbia peplus*, page 248: Weber, Martina

Picture: *Fatsia japonica*, page 248: *PalDat* Picture: *Begonia heracleifolia*, page 248: *PalDat* Picture: *Lysimachia vulgaris*, page 248: *PalDat*

Picture: *Limnanthes douglasii*, page 248: *PalDat* Picture: *Merinthopodium neuranthum*, page 249: *PalDat*

Picture: *Merinthopodium neuranthum*, page 249: *PalDat*

Picture: *Butomus umbellatus*, page 249: *PalDat*

Picture: *Blumenbachia hieronymi*, page 249: *PalDat*

Picture: *Omphalodes verna*, page 249: *PalDat* Picture: *Rhamnus cathartica*, page 249: *PalDat* Picture: *Salix retusa*, page 250: *PalDat*

Picture: *Anchusa cretica*, page 250: *PalDat*

Picture: *Nematanthus strigillosus*, page 250: *PalDat*

Picture: *Fouquieria columnaris*, page 250: *PalDat*

Picture: *Coris monspeliensis*, page 250: *PalDat*

Picture: *Astragalus tragacantha*, page 250: *PalDat*

Picture: *Dianthus carthusianorum*, page 251: *PalDat*

Picture: *Teucrium pyrenaicum*, page 251: *PalDat* Picture: *Babiana ecklonii*, page 251: *PalDat* Picture: *Zea mays*, page 251: *PalDat* Picture: *Dionaea muscipula*, page 251: *PalDat* Picture: *Potentilla incana*, page 251: *PalDat* Picture: *Knautia drymeia*, page 252: *PalDat* Picture: *Tulipa sylvestris*, page 252: *PalDat* Picture: *Cucurbita pepo*, page 252: *PalDat* Picture: *Camellia japonica*, page 252: *PalDat* Picture: *Agrostemma githago*, page 252: *PalDat* Picture: *Passiflora citrina*, page 252: *PalDat* Picture: *Rosa pendulina*, page 253: *PalDat* Picture: *Passiflora suberosa*, page 253: *PalDat* Picture: *Agave asperrima*, page 253: *PalDat* Picture: *Erythronium dens-canis*, page 253: *PalDat* Picture: *Erythronium dens-canis*, page 253: *PalDat* Picture: *Potentilla erecta*, page 253: *PalDat* Picture: *Avena sativa*, page 254: *PalDat* Picture: *Gladiolus illyricus*, page 254: *PalDat* Picture: *Poa pratensis*, page 254: *PalDat* Picture: *Triticum aestivum*, page 254: *PalDat* Picture: *Plantago lanceolata*, page 254: *PalDat* Picture: *Poa angustifolia*, page 254: *PalDat*

Picture: *Anemone transsilvanica*, page 255: *PalDat* Picture: *Opuntia basilaris*, page 255: *PalDat* Picture: *Ranunculus lanuginosus*, page 255: *PalDat* Picture: *Portulaca grandiflora*, page 255: *PalDat* Picture: *Sideritis syriaca*, page 255: *PalDat* Picture: *Corydalis cava*, page 255: *PalDat* Picture: *Stigmaphyllon lindenianum*, page 256: *PalDat*

Picture: *Tristellateia australasiae*, page 256: *PalDat* Picture: *Banisteria muricata*, page 256: *PalDat* Picture: *Malpighia glabra*, page 256: *PalDat* Picture: *Malva moschata*, page 256: Weber, Martina

Picture: *Malva alcea*, page 256: Weber, Martina Picture: *Fumaria officinalis*, page 257: *PalDat* Picture: *Cucurbita pepo*, page 257: *PalDat* Picture: *Ribes aureum*, page 257: *PalDat* Picture: *Costus barbatus*, page 257: *PalDat* Picture: *Amphitecna macrophylla*, page 257: *PalDat*

Picture: *Opuntia phaeacantha*, page 257: *PalDat* Picture: *Cobaea scandens*, page 258: Weber, Martina

Picture: *Juglans* sp., page 258: Preusche, Philipp Picture: *Phaleria capitata*, page 258: *PalDat* Picture: *Dysphania ambrosioides*, page 258: *PalDat* Picture: *Atriplex patula*, page 258: *PalDat* Picture: *Stellaria graminea*, page 258: *PalDat*

Picture: *Cryptomeria* sp., page 259: Zetter, Reinhard

Picture: *Cryptomeria japonica*, page 259: *PalDat* Picture: *Cryptomeria japonica*, page 259: *PalDat* Picture: *Metasequoia glyptostroboides*, page 259:

*PalDat* Picture: *Metasequoia glyptostroboides*, page 259: *PalDat*

Picture: *Cunninghamia lanceolata*, page 259: *PalDat*

Picture: *Pachira sessilis*, page 260: Weber, Martina Picture: *Pachira quinata*, page 260: *PalDat* Picture: *Centaurea segetum*, page 260: *PalDat* Picture: *Arbutus unedo*, page 260: *PalDat* Picture: *Persicaria bistorta*, page 260: *PalDat* Picture: *Tilia platyphyllos*, page 260: Weber, Martina Picture: *Euphorbia tithymaloides*, page 261: *PalDat* Picture: *Justicia brandegeeana*, page 261: *PalDat* Picture: *Schaueria flavicoma*, page 261: *PalDat* Picture: *Echinops exaltatus*, page 261: *PalDat* Picture: *Pachira aquatica*, page 261: *PalDat* Picture: *Pachira aquatica*, page 261: *PalDat* Picture: *Sanguisorba cretica*, page 262: *PalDat* Picture: *Sanguisorba cretica*, page 262: *PalDat* Picture: *Sarcopoterium spinosum*, page 262: *PalDat* Picture: *Sarcopoterium spinosum*, page 262: *PalDat* Picture: *Sanguisorba minor*, page 262: *PalDat* Picture: *Veratrum nigrum*, page 262: *PalDat*

**Service Part** Picture: *Aechmea allenii*, page 263: *PalDat* Picture: *Colchicum autumnale*, page 263: *PalDat* Picture: *Sanchezia nobilis*, page 263: *PalDat* Picture: *Whitfieldia lateralis*, page 263: *PalDat* Picture: *Broussonetia papyrifera*, page 263: *PalDat* Picture: *Quesnelia lateralis*, page 263: *PalDat* Picture: *Betula humilis*, page 264: Weber, Martina Picture: *Betula pendula*, page 264: *PalDat* Picture: *Campanula saxatilis*, page 264: *PalDat* Picture: *Oenothera fruticosa*, page 264: *PalDat* Picture: *Cucumis melo*, page 264: *PalDat* Picture: *Carya* sp., page 264: Zetter, Reinhard Picture: *Amsonia ciliata*, page 265: *PalDat* Picture: *Tetrapollinia caerulescens*, page 265:

*PalDat* Picture: *Knautia arvensis*, page 265: *PalDat* Picture: *Clarkia unguiculata*, page 265: *PalDat* Picture: *Cordia cylindrostachya*, page 265: *PalDat* Picture: *Maclura pomifera*, page 265: *PalDat* Picture: *Phyteuma spicatum*, page 266: Weber,

Martina

Picture: *Myriophyllum spicatum*, page 266: *PalDat* Picture: *Campanula alpina*, page 266: *PalDat* Picture: *Aechmea fulgens*, page 266: *PalDat* Picture: *Aechmea tomentosa*, page 266: *PalDat* Picture: *Asyneuma canescens*, page 266: *PalDat*

Picture: *Alnus glutinosa*, page 267: *PalDat*

Picture: *Legousia speculum-veneris*, page 267: *PalDat*

Picture: *Campanula garganica*, page 267: *PalDat* Picture: *Irlbachia pendula*, page 267: *PalDat* Picture: *Elatostema ambiguum*, page 267: *PalDat* Picture: *Campanula rapunculoides*, page 267:

*PalDat*

Picture: *Alnus viridis*, page 268: *PalDat* Picture: *Pterocarya* sp., page 268: Zetter, Reinhard

Picture: *Drosera kansaiensis*, page 268: *PalDat*

Picture: *Paramoltkia doerfleri*, page 268: *PalDat*

Picture: *Ulmus minor*, page 268: *PalDat*

Picture: *Megaskepasma erythrochlamys*, page 268: *PalDat*

Picture: *Stellaria holostea*, page 269: *PalDat* Picture: *Cobaea scandens*, page 269: *PalDat* Picture: *Ipomoea batatas*, page 269: *PalDat* Picture: *Helianthium bolivianum*, page 269: *PalDat* Picture: *Calystegia sepium*, page 269: *PalDat* Picture: *Aechmea azurea*, page 269: *PalDat* Picture: *Bassia scoparia*, page 270: *PalDat* Picture: *Bassia scoparia*, page 270: *PalDat* Picture: *Plantago major*, page 270: *PalDat* Picture: *Juglans regia*, page 270: *PalDat* Picture: *Buxus sempervirens*, page 270: *PalDat* Picture: *Kallstroemia maxima*, page 270: *PalDat* Picture: *Liquidambar* sp., page 271: Zetter, Reinhard

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8

© Springer International Publishing AG 2018

Picture: *Pavonia multiflora*, page 271: Weber, Martina

Picture: *Arenaria ciliata*, page 271: *PalDat* Picture: *Lavatera thuringiaca*, page 271: *PalDat* Picture: *Whitfieldia elongata*, page 271: *PalDat* Picture: *Alisma lanceolatum*, page 271: *PalDat* Picture: *Carex remota*, page 272: *PalDat* Picture: *Cercidiphyllum japonicum*, page 272: *PalDat*

Picture: *Sagittaria sagittifolia*, page 272: *PalDat* Picture: *Caldesia parnassifolia*, page 272: *PalDat* Picture: *Schoenoplectus lacustris*, page 272: *PalDat* Picture: *Scirpus sylvaticus*, page 272: *PalDat* Picture: *Lythrum salicaria*, page 273: *PalDat* Picture: *Asperugo procumbens*, page 273: *PalDat* Picture: *Lumnitzera racemosa*, page 273: *PalDat* Picture: *Cynoglossum officinale*, page 273: *PalDat* Picture: *Justicia furcata*, page 273: *PalDat* Picture: *Pachystachys lutea*, page 273: *PalDat* Picture: *Zamioculcas zamiifolia*, page 274: Zetter,

Reinhard

Picture: *Zamioculcas zamiifolia*, page 274: Zetter, Reinhard

Picture: *Zamioculcas zamiifolia*, page 274: *PalDat* Picture: *Monstera deliciosa*, page 274: *PalDat* Picture: *Gonatopus angustus*, page 274: *PalDat* Picture: *Gonatopus angustus*, page 274: *PalDat* Picture: *Pedicularis palustris*, page 275: *PalDat* Picture: *Pedicularis palustris*, page 275: *PalDat*

Picture: *Pedicularis rostratocapitata*, page 275: *PalDat*

Picture: *Iris histrioides*, page 275: *PalDat* Picture: *Cephalostemon riedelianus*, page 275: *PalDat*

Picture: *Limnanthes douglasii*, page 275: *PalDat* Picture: *Gonatopus boivinii*, page 276: Ulrich, Silvia Picture: *Victoria regia*, page 276: *PalDat* Picture: *Pedicularis gyroflexa*, page 276: *PalDat* Picture: *Passiflora amethystina*, page 276: *PalDat* Picture: *Acacia dealbata*, page 276: *PalDat* Picture: *Acacia dealbata*, page 276: *PalDat* Picture: *Mimulus* sp., page 277: Weber, Martina Picture: *Thunbergia alata*, page 277: Weber,

Martina Picture: *Mimulus guttatus*, page 277: *PalDat* Picture: *Thunbergia alata*, page 277: *PalDat* Picture: *Claytonia perfoliata*, page 277: *PalDat* Picture: *Berberis vulgaris*, page 277: *PalDat*

Picture: *Aphyllanthes monspeliensis*, page 278: *PalDat*

Picture: *Berberis amurensis*, page 278: *PalDat* Picture: *Crocus speciosus*, page 278: *PalDat* Picture: *Mimulus guttatus*, page 278: *PalDat* Picture: *Thunbergia laurifolia*, page 278: *PalDat* Picture: *Bignonia magnifica*, page 278: *PalDat*

	- Grímsson, Friðgeir

### **Chapter: "Ornamentation"**

**Service Part** Picture: *Peperomia rubella*, page 296: *PalDat* Picture: *Dracunculus vulgaris*, page 296: *PalDat* Picture: *Mimosa pudica*, page 296: *PalDat* Picture: *Poikilacanthus macranthus*, page 296: *PalDat* Picture: *Cynodon dactylon*, page 296: *PalDat* Picture: *Justicia brandegeeana*, page 296: *PalDat* Picture: *Justicia carnea*, page 297: Weber, Martina Picture: *Justicia carnea*, page 297: *PalDat* Picture: *Peperomia polybotrya*, page 297: *PalDat* Picture: *Peperomia polybotrya*, page 297: *PalDat* Picture: *Megaskepasma erythrochlamys*, page 297: *PalDat* Picture: *Megaskepasma erythrochlamys*, page 297: *PalDat* Picture: *Viscum album*, page 298: *PalDat* Picture: *Viscum album*, page 298: *PalDat* Picture: *Viscum album*, page 298: *PalDat* Picture: *Nymphaea alba*, page 298: Weber, Martina Picture: *Erythrochiton brasiliensis*, page 298: *PalDat* Picture: *Erythrochiton brasiliensis*, page 298: *PalDat* Picture: *Ocimum basilicum*, page 299: *PalDat* Picture: *Ocimum basilicum*, page 299: *PalDat* Picture: *Ocimum basilicum*, page 299: *PalDat* Picture: *Salvia hians*, page 299: *PalDat* Picture: *Salvia argentea*, page 299: *PalDat* Picture: *Salvia glutinosa*, page 299: *PalDat* Picture: *Phlox drummondii*, page 300: *PalDat* Picture: *Phlox paniculata*, page 300: *PalDat* Picture: *Agastache mexicana*, page 300: *PalDat* Picture: *Agastache mexicana*, page 300: *PalDat* Picture: *Vitex trifolia*, page 300: *PalDat* Picture: *Pachystachys lutea*, page 300: *PalDat* Picture: *Plectranthus ornatus*, page 301: *PalDat* Picture: *Hyptis suaveolens*, page 301: *PalDat* Picture: *Sparmannia africana*, page 301: *PalDat* Picture: *Melittis melissophyllum*, page 301: *PalDat* Picture: *Prunella grandiflora*, page 301: *PalDat* Picture: *Andrographis paniculata*, page 301: *PalDat* Picture: *Iris planifolia*, page 302: *PalDat* Picture: *Ilex* sp., page 302: *PalDat* Picture: *Ilex aquifolium*, page 302: *PalDat* Picture: *Ilex aquifolium*, page 302: *PalDat*

© Springer International Publishing AG 2018 Picture: *Linum flavum*, page 302: *PalDat* Picture: *Aratitiyopea lopezii*, page 302: *PalDat* Picture: *Linum* sp., page 303: *PalDat* Picture: *Hymenocallis tubiflora*, page 303: *PalDat* Picture: *Linum capitatum*, page 303: *PalDat* Picture: *Linum capitatum*, page 303: *PalDat* Picture: *Geranium robertianum*, page 303: *PalDat* Picture: *Geranium robertianum*, page 303: *PalDat* Picture: *Jatropha multifida*, page 304: *PalDat* Picture: *Plumbago auriculata*, page 304: *PalDat*

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8

Picture: *Hymenocallis littoralis*, page 304: *PalDat* Picture: *Viburnum lantana*, page 304: *PalDat* Picture: *Geranium pratense*, page 304: *PalDat* Picture: *Geranium sibiricum*, page 304: *PalDat* Picture: *Ibicella lutea*, page 305: *PalDat* Picture: *Ibicella lutea*, page 305: *PalDat* Picture: *Proboscidea fragrans*, page 305: *PalDat* Picture: *Pseudofumaria lutea*, page 305: *PalDat* Picture: *Iris bucharica*, page 305: *PalDat* Picture: *Iris bucharica*, page 305: *PalDat* Picture: *Catalpa bignonioides*, page 306: *PalDat* Picture: *Iris graeberiana*, page 306: *PalDat* Picture: *Lophophora williamsii*, page 306: *PalDat* Picture: *Phyllanthus* sp., page 306: *PalDat* Picture: *Irlbachia pedunculata*, page 306: *PalDat* Picture: *Banisteria muricata*, page 306: *PalDat* Picture: *Croton triqueter*, page 307: *PalDat* Picture: *Croton triqueter*, page 307: Weber, Martina

Picture: *Jatropha podagrica*, page 307: *PalDat* Picture: *Daphne laureola*, page 307: *PalDat* Picture: *Daphne cneorum*, page 307: *PalDat* Picture: *Thymelaea passerina*, page 307: *PalDat* Picture: *Garcia nutans*, page 308: Weber, Martina Picture: *Garcia nutans*, page 308: *PalDat* Picture: *Daphne tangutica*, page 308: *PalDat* Picture: *Callitriche stagnalis*, page 308: *PalDat* Picture: *Croton triqueter*, page 308: *PalDat* Picture: *Thymelaea passerina*, page 308: *PalDat* Picture: *Galinsoga ciliata*, page 309: *PalDat* Picture: *Carduus acanthoides*, page 309: *PalDat* Picture: *Hibiscus trionum*, page 309: *PalDat* Picture: *Lavatera thuringiaca*, page 309: *PalDat* Picture: *Pinellia ternata*, page 309: *PalDat* Picture: *Pinellia ternata*, page 309: *PalDat* Picture: *Pavonia multiflora*, page 310: Weber,

Martina Picture: *Abutilon megapotamicum*, page 310: Weber, Martina

Picture: *Calendula officinalis*, page 310: Weber, Martina

Picture: *Drosera scorpioides*, page 310: Weber, Martina

Picture: *Nuphar lutea*, page 310: Weber, Martina Picture: *Alocasia acuminata*, page 310: Ulrich, Silvia

Picture: *Patrinia gibbosa*, page 311: *PalDat* Picture: *Hieracium hoppeanum*, page 311: *PalDat* Picture: *Ambrosia artemisiifolia*, page 311: *PalDat* Picture: *Aster amellus*, page 311: *PalDat* Picture: *Portulaca grandiflora*, page 311: *PalDat* Picture: *Nuphar lutea*, page 311: *PalDat* Picture: *Petasites albus*, page 312: *PalDat* Picture: *Antennaria dioica*, page 312: *PalDat* Picture: *Malva neglecta*, page 312: *PalDat* Picture: *Tanacetum corymbosum*, page 312: *PalDat*

Picture: *Lonicera fragrantissima*, page 312: *PalDat* Picture: *Ipomoea purpurea*, page 312: *PalDat* Picture: *Arenga pinnata*, page 313: *PalDat* Picture: *Zomicarpa riedeliana*, page 313: *PalDat* Picture: *Helianthus annuus*, page 313: *PalDat* Picture: *Knautia drymeia*, page 313: *PalDat* Picture: *Campanula alpina*, page 313: *PalDat* Picture: *Ulearum sagittatum*, page 313: *PalDat* Picture: *Scorzonera austriaca*, page 314: *PalDat* Picture: *Ipomoea batatas*, page 314: *PalDat* Picture: *Tetrapollinia caerulescens*, page 314: *PalDat*

Picture: *Gynura scandens*, page 314: *PalDat* Picture: *Stratiotes aloides*, page 314: *PalDat* Picture: *Hibiscus schizopetalus*, page 314: *PalDat* Picture: *Mendoncia albida*, page 315: *PalDat* Picture: *Erica herbacea*, page 315: *PalDat* Picture: *Rhododendron hirsutum*, page 315: *PalDat* Picture: *Ledum palustre*, page 315: *PalDat* Picture: *Aristolochia manshuriensis*, page 315:

*PalDat* Picture: *Moneses uniflora*, page 315: *PalDat* Picture: *Pithecellobium dulce*, page 316: *PalDat* Picture: *Pithecellobium dulce*, page 316: *PalDat* Picture: *Gaultheria myrsinoides*, page 316: *PalDat* Picture: *Besleria hirsuta*, page 316: *PalDat* Picture: *Lagerstroemia indica*, page 316: *PalDat* Picture: *Lagerstroemia indica*, page 316: *PalDat* Picture: *Streptocalyx poeppigii*, page 317: *PalDat* Picture: *Canistrum camacaense*, page 317: *PalDat* Picture: *Lavandula angustifolia*, page 317: *PalDat* Picture: *Aechmea araneosa*, page 317: *PalDat* Picture: *Glossoloma ichthyoderma*, page 317:

*PalDat* Picture: *Glossoloma ichthyoderma*, page 317:

*PalDat*

Picture: *Nematanthus strigillosus*, page 318: *PalDat*

Picture: *Cyrtosperma beccarianum*, page 318: *PalDat*

Picture: *Maianthemum stellatum*, page 318: *PalDat* Picture: *Hohenbergia stellata*, page 318: *PalDat* Picture: *Aechmea ampla*, page 318: *PalDat* Picture: *Aechmea ampla*, page 318: *PalDat* Picture: *Ruellia makoyana*, page 319: *PalDat* Picture: *Impatiens parviflora*, page 319: *PalDat* Picture: *Erophila verna*, page 319: *PalDat* Picture: *Bougainvillea* sp., page 319: *PalDat* Picture: *Viburnum opulus*, page 319: *PalDat* Picture: *Thladiantha hookeri*, page 319: *PalDat* Picture: *Ruellia devosiana*, page 320: *PalDat* Picture: *Persicaria chinensis*, page 320: *PalDat* Picture: *Fouquieria macdougalii*, page 320: *PalDat* Picture: *Ardisia elliptica*, page 320: *PalDat* Picture: *Schotia brachypetala*, page 320: *PalDat* Picture: *Passiflora amethystina*, page 320: *PalDat* Picture: *Dionaea muscipula*, page 321: *PalDat*

Picture: *Cephalopentandra ecirrhosa*, page 321: *PalDat*

Picture: *Stenandrium dulce*, page 321: *PalDat* Picture: *Asarum europaeum*, page 321: *PalDat* Picture: *Fatsia japonica*, page 321: *PalDat* Picture: *Hakea kippistiana*, page 321: *PalDat* Picture: *Turnera ulmifolia*, page 322: *PalDat* Picture: *Wachendorfia thyrsiflora*, page 322: *PalDat* Picture: *Cantua buxifolia*, page 322: *PalDat* Picture: *Cantua buxifolia*, page 322: *PalDat* Picture: *Roridula gorgonias*, page 322: *PalDat* Picture: *Calluna vulgaris*, page 322: *PalDat* Picture: *Hemiptelia* sp., page 323: Grímsson, Friðgeir Picture: *Clarkia pulchella*, page 323: *PalDat* Picture: *Luzula campestris*, page 323: *PalDat* Picture: *Larix decidua*, page 323: *PalDat* Picture: *Humulus lupulus*, page 323: *PalDat* Picture: *Clarkia purpurea*, page 323: *PalDat* Picture: *Hedera helix*, page 324: *PalDat* Picture: *Lachenalia aloides*, page 324: *PalDat* Picture: *Fritillaria meleagris*, page 324: *PalDat* Picture: *Anthericum ramosum*, page 324: *PalDat* Picture: *Billbergia seidelii*, page 324: *PalDat* Picture: *Limodorum abortivum*, page 324: *PalDat* Picture: *Ornithogalum narbonense*, page 325:

*PalDat* Picture: *Ornithogalum narbonense*, page 325: *PalDat*

Picture: *Bomarea hirsuta*, page 325: *PalDat*

Picture: *Bessera elegans*, page 325: Weber, Martina

Picture: *Chelonanthus alatus*, page 325: *PalDat*

Picture: *Plectranthus esculentus*, page 325: Weber, Martina

Picture: *Acantholimon glumaceum*, page 326: *PalDat*

Picture: *Abeliophyllum distichum*, page 326: *PalDat* Picture: *Eranthemum wattii*, page 326: *PalDat* Picture: *Strobilanthes roseus*, page 326: *PalDat* Picture: *Ruellia* sp., page 326: *PalDat* Picture: *Thlaspi montanum*, page 326: *PalDat* Picture: *Armeria pinifolia*, page 327: *PalDat* Picture: *Armeria pinifolia*, page 327: *PalDat* Picture: *Impatiens parviflora*, page 327: Weber, Martina Picture: *Persicaria chinensis*, page 327: *PalDat* Picture: *Kallstroemia maxima*, page 327: *PalDat* Picture: *Ruellia tuberosa*, page 327: *PalDat* Picture: *Leontodon saxatilis*, page 328: *PalDat*

Picture: *Cichorium intybus*, page 328: *PalDat*

Picture: *Opuntia basilaris*, page 328: *PalDat*

Picture: *Pfaffia tuberosa*, page 328: *PalDat*

Picture: *Gazania* sp., page 328: *PalDat*

Picture: *Hieracium hoppeanum*, page 328: *PalDat* Picture: *Tragopogon orientalis*, page 329: Weber,

Martina

Picture: *Prenanthes purpurea*, page 329: Weber, Martina

Picture: *Cichorium intybus*, page 329: Weber, Martina

Picture: *Taraxacum* sp., page 329: Zetter, Reinhard Picture: *Tragopogon dubius*, page 329: *PalDat* Picture: *Crepis biennis*, page 329: *PalDat* Picture: *Cyanthillium cinereum*, page 330: *PalDat* Picture: *Gomphrena celosioides*, page 330: *PalDat* Picture: *Opuntia polyacantha*, page 330: *PalDat* Picture: *Scorzonera aristata*, page 330: *PalDat* Picture: *Ipomoea caerulea*, page 330: *PalDat* Picture: *Herniaria alpina*, page 330: *PalDat* Picture: *Heloniopsis kawanoi*, page 331: *PalDat* Picture: *Trillium grandiflorum*, page 331: *PalDat* Picture: *Kickxia spuria*, page 331: *PalDat* Picture: *Lamium purpureum*, page 331: *PalDat* Picture: *Aspidistra elatior*, page 331: *PalDat* Picture: *Callisia fragrans*, page 331: *PalDat* Picture: *Orobanche lutea*, page 332: *PalDat* Picture: *Trillium chloropetalum*, page 332: *PalDat* Picture: *Claytonia perfoliata*, page 332: *PalDat* Picture: *Petrorhagia prolifera*, page 332: *PalDat* Picture: *Veronica prostrata*, page 332: *PalDat* Picture: *Ptelea trifoliolata*, page 332: *PalDat* Picture: *Podophyllum peltatum*, page 333: *PalDat* Picture: *Solanum torvum*, page 333: *PalDat* Picture: *Campanula persicifolia*, page 333: *PalDat* Picture: *Melampyrum pratense*, page 333: *PalDat* Picture: *Tilia* sp., page 333: Grímsson, Friðgeir Picture: *Tilia* sp., page 333: Grímsson, Friðgeir Picture: *Tradescantia spathacea*, page 334: *PalDat*

Picture: *Aspidistra locii*, page 334: *PalDat* Picture: *Smilax spinosa*, page 334: *PalDat* Picture: *Clethra arborea*, page 334: *PalDat* Picture: *Quercus robur*, page 334: *PalDat* Picture: *Erica pageana*, page 334: *PalDat* Picture: *Elaeagnus rhamnoides*, page 335: *PalDat* Picture: *Polyscias filicifolia*, page 335: *PalDat* Picture: *Cytisus nigricans*, page 335: *PalDat* Picture: *Reseda luteola*, page 335: *PalDat* Picture: *Anemone pratensis*, page 335: *PalDat* Picture: *Drosera kansaiensis*, page 335: *PalDat* Picture: *Babiana ecklonii*, page 336: *PalDat* Picture: *Galium lucidum*, page 336: *PalDat* Picture: *Scirpoides holoschoenus*, page 336: *PalDat* Picture: Poaceae, page 336: Grímsson, Friðgeir Picture: *Callisia fragrans*, page 336: *PalDat* Picture: *Oenothera fruticosa*, page 336: *PalDat* Picture: *Dianella tasmanica*, page 337: *PalDat* Picture: *Veronica longifolia*, page 337: *PalDat* Picture: *Symphytum caucasicum*, page 337: *PalDat*

Picture: *Hordeum bulbosum*, page 337: *PalDat* Picture: *Juglans* sp., page 337: Grímsson, Friðgeir

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8

© Springer International Publishing AG 2018

Picture: Amaranthaceae, page 337: Grímsson, Friðgeir

Picture: *Carex filiformis*, page 338: *PalDat*

Picture: Cyperaceae, page 338: Grímsson, Friðgeir

Picture: *Viola calcarata*, page 338:

Picture: *Tilia* sp., page 338: Grímsson, Friðgeir


Picture: *Drosera binata*, page 340: *PalDat* Picture: *Alisma* sp., page 340: Grímsson, Friðgeir Picture: *Heliohebe raoulii*, page 340: *PalDat* Picture: *Maclura pomifera*, page 340: *PalDat* Picture: *Trapa* sp., page 341: Grímsson, Friðgeir Picture: Onagraceae, page 341: Grímsson, Friðgeir Picture: *Pedicularis verticillata*, page 341: *PalDat* Picture: *Tetragonia tetragonioides*, page 341: *PalDat*


Picture: *Pistia stratiotes*, page 344: *PalDat* Picture: *Pistia stratiotes*, page 344: *PalDat* Picture: *Welwitschia mirabilis*, page 345: *PalDat* Picture: *Amorphophallus lacourii*, page 345: *PalDat*

Picture: *Amorphophallus serrulatus*, page 345: Ulrich, Silvia

Picture: *Brillantaisia owariensis*, page 345: *PalDat*


Picture: *Androlepis skinneri*, page 347: *PalDat* Picture: *Trigonia nivea*, page 347: *PalDat* Picture: *Maxillaria densa*, page 347: *PalDat* Picture: *Cheilocostus speciosus*, page 347: *PalDat* Picture: *Whitfieldia lateritia*, page 347: *PalDat* Picture: *Cardamine pratensis*, page 348: *PalDat* Picture: *Luffa cylindrica*, page 348: *PalDat* Picture: *Physostegia virginiana*, page 348: *PalDat* Picture: *Aechmea azurea*, page 348: *PalDat* Picture: *Beloperone guttata*, page 348: *PalDat* Picture: *Razisea citrina*, page 348: *PalDat* Picture: *Salix daphnoides*, page 349: *PalDat* Picture: *Persicaria mitis*, page 349: *PalDat* Picture: *Paradisea liliastrum*, page 349: *PalDat* Picture: *Trifolium rubens*, page 349: *PalDat* Picture: *Ecballium elaterium*, page 349: *PalDat* Picture: *Ajuga genevensis*, page 349: *PalDat* Picture: *Ruellia brevifolia*, page 350: *PalDat* Picture: *Buxus sempervirens*, page 350: *PalDat* Picture: *Impatiens glandulifera*, page 350: *PalDat* Picture: *Lupinus polyphyllus*, page 350: *PalDat* Picture: *Opuntia paraguayensis*, page 350: *PalDat* Picture: *Theobroma cacao*, page 350: *PalDat* Picture: *Tropaeolum majus*, page 351: *PalDat* Picture: *Veratrum album*, page 351: *PalDat* Picture: *Werauhia tarmaensis*, page 351: *PalDat* Picture: *Erythronium dens-canis*, page 351: *PalDat* Picture: *Poncirus trifoliata*, page 351: *PalDat* Picture: *Melilotus officinalis*, page 351: *PalDat* Picture: *Vigna speciosa*, page 352: *PalDat* Picture: *Epipactis helleborine*, page 352: *PalDat* Picture: *Harpochilus neesianus*, page 352: *PalDat* Picture: *Polygala major*, page 352: *PalDat* Picture: *Adenia fruticosa*, page 352: *PalDat* Picture: *Pisum sativum*, page 352: *PalDat* Picture: *Thladiantha hookeri*, page 353: *PalDat* Picture: *Pinguicula alpina*, page 353: *PalDat* Picture: *Billardiera heterophylla*, page 353: *PalDat*

Picture: *Cephalanthera longifolia*, page 353: *PalDat*

Picture: *Aechmea allenii*, page 353: *PalDat* Picture: *Lathyrus vernus*, page 353: *PalDat* Picture: *Jasminum nudiflorum*, page 354: Weber, Martina

Picture: Lamiaceae, page 354: Weber, Martina Picture: *Iris pseudacorus*, page 354: Weber, Martina Picture: *Iris domestica*, page 354: Weber, Martina Picture: *Fraxinus excelsior*, page 354: *PalDat* Picture: *Persicaria* sp., page 354: Zetter, Reinhard Picture: *Lilium martagon*, page 355: *PalDat* Picture: *Lilium candidum*, page 355: *PalDat* Picture: *Phaleria capitata*, page 355: Weber, Martina

Picture: *Phaleria capitata*, page 355: *PalDat* Picture: *Fumana procumbens*, page 355: *PalDat* Picture: *Erdtmanipollis*, page 355: Zetter, Reinhard Picture: *Anthurium gracile*, page 356: *PalDat* Picture: *Anthurium gracile*, page 356: *PalDat* Picture: *Geranium pratense*, page 356: Weber, Martina

Picture: *Geranium reuteri*, page 356: *PalDat* Picture: *Sarcococca pruniformis*, page 356: *PalDat* Picture: *Sarcococca pruniformis*, page 356: *PalDat* Picture: Liliaceae, page 357: Weber, Martina Picture: *Pachira aquatica*, page 357: *PalDat* Picture: *Pachysandra terminalis*, page 357: *PalDat* Picture: *Mercurialis perennis*, page 357: *PalDat* Picture: *Aponogeton masoalaensis*, page 357: *PalDat*

Picture: *Armeria maritima*, page 357: *PalDat* Picture: *Securigera varia*, page 358: *PalDat* Picture: *Zelkova* sp., page 358: Zetter, Reinhard Picture: *Peucedanum cervaria*, page 358: *PalDat* Picture: *Fagus* sp., page 358: *PalDat* Picture: *Sedum acre*, page 358: *PalDat* Picture: *Circaea lutetiana*, page 358: *PalDat* Picture: *Carpinus betulus*, page 359: *PalDat* Picture: *Acer negundo*, page 359: *PalDat* Picture: *Nymphoides peltata*, page 359: *PalDat* Picture: *Leucadendron discolor*, page 359: *PalDat* Picture: *Nicotiana tabacum*, page 359: *PalDat* Picture: *Myrrhis odorata*, page 359: *PalDat* Picture: *Fagus* sp., page 360: Zetter, Reinhard Picture: *Populus alba*, page 360: Weber, Martina Picture: *Dioon edule*, page 360: Weber, Martina Picture: *Betula humilis*, page 360: Weber, Martina Picture: *Ceratozamia mexicana*, page 360: Weber, Martina

Picture: *Sanguisorba officinalis*, page 360: Weber, Martina

Picture: *Acer pseudoplatanus*, page 361: *PalDat* Picture: *Prunus avium*, page 361: *PalDat* Picture: *Potentilla inclinata*, page 361: *PalDat* Picture: *Veronica cinerea*, page 361: *PalDat*

**Service Part** Picture: *Amherstia nobilis*, page 361: *PalDat* Picture: *Gentiana lutea*, page 361: *PalDat* Picture: *Saxifraga rotundifolia*, page 362: *PalDat* Picture: *Aesculus hippocastanum*, page 362:

*PalDat*

Picture: *Lycium barbarum*, page 362: *PalDat* Picture: *Chaenomeles sinensis*, page 362: *PalDat* Picture: *Rubus caesius*, page 362: *PalDat* Picture: *Allium flavum*, page 362: *PalDat* Picture: *Geum reptans*, page 363: *PalDat* Picture: *Sanguisorba minor*, page 363: *PalDat* Picture: *Saxifraga taygetea*, page 363: *PalDat* Picture: *Saxifraga taygetea*, page 363: *PalDat* Picture: *Begonia heracleifolia*, page 363: *PalDat* Picture: *Cabomba palaeformis*, page 363: *PalDat* Picture: *Cuphea llavea*, page 364: *PalDat* Picture: *Crataegus laevigata*, page 364: *PalDat* Picture: *Malus sylvestris*, page 364: *PalDat*

Picture: *Neoalsomitra sarcophylla*, page 364: *PalDat*

Picture: *Ruta graveolens*, page 364: *PalDat*

Picture: *Helianthemum nummularium*, page 364: *PalDat*

Picture: *Menyanthes trifoliata*, page 365: *PalDat* Picture: *Prunus laurocerasus*, page 365: *PalDat* Picture: *Saxifraga tridactylites*, page 365: *PalDat*

Picture: *Acer pseudoplatanus*, page 365: *PalDat*

Picture: *Amorphophallus interruptus*, page 365: Ulrich, Silvia

Picture: *Amorphophallus serrulatus*, page 365: Ulrich, Silvia

Picture: *Blackstonia perfoliata*, page 366: *PalDat*

Picture: *Erodium cicutarium*, page 366: *PalDat*

Picture: *Gentianella austriaca*, page 366: *PalDat*

Picture: *Gelsemium sempervirens*, page 366: *PalDat*

Picture: *Cotinus coggygria*, page 366: *PalDat*

Picture: *Cotinus coggygria*, page 366: *PalDat* Picture: *Pelargonium tetragonum*, page 367:

*PalDat* Picture: *Pelargonium carnosum*, page 367: *PalDat*

Picture: *Fouquieria macdougalii*, page 367: *PalDat*

Picture: *Fouquieria macdougalii*, page 367: *PalDat*

Picture: *Ailanthus altissima*, page 367: *PalDat*

Picture: *Cistus clusii*, page 367: *PalDat*

Picture: *Solandra longiflora*, page 368: *PalDat*

Picture: *Pelargonium punctatum*, page 368: *PalDat*

Picture: *Gentiana acaulis*, page 368: *PalDat*

Picture: *Polemonium caeruleum*, page 368: *PalDat* Picture: *Polemonium caeruleum*, page 368: Weber,

Martina

Picture: *Polemonium pauciflorum*, page 368: *PalDat*

© Springer International Publishing AG 2018

Picture: *Brugmansia suaveolens*, page 369: *PalDat* Picture: *Chlorospatha pubescens*, page 369:

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8

*PalDat*

Picture: *Schinus molle*, page 369: *PalDat* Picture: *Solandra maxima*, page 369: *PalDat* Picture: *Plagiorhegma dubium*, page 369: *PalDat* Picture: *Amherstia nobilis*, page 369: *PalDat* Picture: *Anthurium gracile*, page 370: *PalDat* Picture: *Anthurium gracile*, page 370: *PalDat* Picture: *Triglochin maritima*, page 370: *PalDat* Picture: *Ascarina lucida*, page 370: Grímsson, Friðgeir

Picture: *Lophophora williamsii*, page 370: *PalDat* Picture: *Phillyrea angustifolia*, page 370: *PalDat* Picture: *Fraxinus* sp., page 371: Grímsson, Friðgeir Picture: *Tulipa turkestanica*, page 371: *PalDat* Picture: *Helleborus foetidus*, page 371: *PalDat* Picture: *Cercidiphyllum* sp., page 371: Grímsson,

Friðgeir Picture: *Parentucellia latifolia*, page 371: *PalDat* Picture: *Viburnum utile*, page 371: *PalDat* Picture: *Linum bienne*, page 372: *PalDat* Picture: *Saxifraga hostii*, page 372: *PalDat* Picture: *Betula* sp., page 372: Grímsson, Friðgeir Picture: *Piper auritum*, page 372: *PalDat* Picture: *Ulmus* sp., page 372: Grímsson, Friðgeir Picture: *Tsuga* sp., page 372: Grímsson, Friðgeir Picture: *Ceratostigma plumbaginoides*, page 373:

*PalDat* Picture: *Ilex* sp., page 373: Grímsson, Friðgeir Picture: *Akebia quinata*, page 373: *PalDat* Picture: *Polemonium caeruleum*, page 373: *PalDat* Picture: *Ouratea hexasperma*, page 373: *PalDat* Picture: *Coffea arabica*, page 373: *PalDat* Picture: *Geranium robertianum*, page 374: *PalDat* Picture: *Geranium robertianum*, page 374: Weber, Martina

Picture: *Atriplex sagittata*, page 374: *PalDat* Picture: *Pulsatilla grandis*, page 374: *PalDat* Picture: *Quercus robur*, page 374: *PalDat* Picture: *Plantago maritima*, page 374: *PalDat* Picture: *Aristolochia arborea*, page 375: *PalDat* Picture: *Plantago media*, page 375: *PalDat* Picture: *Aristolochia tricaudata*, page 375: *PalDat* Picture: *Calliandra tergemina*, page 375: *PalDat* Picture: *Corydalis cava*, page 375: *PalDat* Picture: *Corydalis cava*, page 375: Weber, Martina Picture: *Teucrium chamaedrys*, page 376: *PalDat* Picture: *Cyanastrum cordifolium*, page 376: *PalDat* Picture: *Platycapnos tenuilobus*, page 376: *PalDat* Picture: *Platycapnos tenuilobus*, page 376: *PalDat* Picture: *Plantago reniformis*, page 376: *PalDat* Picture: *Plantago lanceolata*, page 376: *PalDat* Picture: *Amorphophallus stuhlmannii*, page 377:

Ulrich, Silvia Picture: *Amorphophallus stuhlmannii*, page 377:

*PalDat*

Picture: *Sarcocapnos enneaphylla*, page 377: *PalDat*

Picture: *Commelina erecta*, page 377: *PalDat* Picture: *Stanfieldiella imperforata*, page 377: *PalDat*

Picture: *Callistemon comboynensis*, page 377: *PalDat*

#### **Chapter: "Pollen Wall"**

Picture: *Plantago major*, page 380: *PalDat* Picture: *Salvia verticillata*, page 380: *PalDat* Picture: *Galeopsis tetrahit*, page 380: *PalDat* Picture: *Galeopsis tetrahit*, page 380: *PalDat* Picture: *Salvia nemorosa*, page 380: *PalDat* Picture: *Salvia nemorosa*, page 380: *PalDat* Picture: *Corylus avellana*, page 381: *PalDat* Picture: *Ambrosia artemisiifolia*, page 381: *PalDat* Picture: *Plantago maritima*, page 381: *PalDat* Picture: *Adonis aestivalis*, page 381: *PalDat* Picture: *Quercus robur*, page 381: *PalDat* Picture: *Apium nodiflorum*, page 381: *PalDat* Picture: *Ailanthus altissima*, page 382: *PalDat* Picture: *Ailanthus altissima*, page 382: *PalDat* Picture: *Fraxinus excelsior*, page 382: *PalDat* Picture: *Fraxinus excelsior*, page 382: *PalDat* Picture: *Odontites luteus*, page 382: *PalDat* Picture: *Odontites luteus*, page 382: *PalDat* Picture: *Salix fragilis*, page 383: *PalDat* Picture: *Salix fragilis*, page 383: *PalDat* Picture: *Alangium* sp., page 383: *PalDat* Picture: *Lomatogonium carinthiacum*, page 383: *PalDat* Picture: *Quesnelia lateralis*, page 383: *PalDat* Picture: *Pachysandra terminalis*, page 383: *PalDat* Picture: *Rhaphidophora africana*, page 384: *PalDat* Picture: *Orobanche hederae*, page 384: *PalDat* Picture: *Sauromatum venosum*, page 384: *PalDat* Picture: *Globba schomburgkii*, page 384: *PalDat* Picture: *Sauromatum venosum*, page 384: Weber, Martina Picture: *Spathicarpa* sp., page 384: Weber, Martina Picture: *Abies* sp., page 385: Zetter, Reinhard Picture: *Pinus* sp., page 385: Zetter, Reinhard Picture: *Pinus* sp., page 385: Zetter, Reinhard Picture: *Tsuga* sp., page 385: Zetter, Reinhard Picture: *Gonatopus angustus*, page 385: *PalDat*

Picture: *Zamioculcas zamiifolia*, page 385: *PalDat* Picture: *Solidago canadensis*, page 386: *PalDat*

Picture: *Cichorium intybus*, page 386: Weber, Martina

Picture: Asteraceae, page 386: Ulrich, Silvia Picture: *Solidago canadensis*, page 386: *PalDat* Picture: Asteraceae, page 386: Ulrich, Silvia Picture: *Chaenorhinum minus*, page 386: *PalDat* Picture: *Mentha aquatica*, page 387: *PalDat*

Picture: *Cobaea scandens*, page 387: Weber, Martina Picture: *Mentha aquatica*, page 387: *PalDat* Picture: Brassicaceae, page 387: Ulrich, Silvia Picture: *Ligustrum vulgare*, page 387: Weber, Martina Picture: *Melampyrum pratense*, page 387: *PalDat* Picture: *Bassia scoparia*, page 388: *PalDat* Picture: Asteraceae, page 388: Halbritter, Heidemarie Picture: Asteraceae, page 388: Ulrich, Silvia Picture: *Tanacetum corymbosum*, page 388: Weber, Martina Picture: Asteraceae, page 388: Ulrich, Silvia Picture: *Artemisia vulgaris*, page 388: Weber, Martina Picture: *Amydrium medium*, page 389: *PalDat* Picture: *Corylus colurna*, page 389: *PalDat* Picture: *Viola tricolor*, page 389: *PalDat* Picture: *Juglans regia*, page 389: *PalDat* Picture: *Ulmus laevis*, page 389: *PalDat* Picture: *Plantago lanceolata*, page 389: *PalDat* Picture: *Dieffenbachia humilis*, page 390: *PalDat* Picture: *Mahonia aquifolium*, page 390: *PalDat* Picture: *Berberis vulgaris*, page 390: *PalDat* Picture: *Iris pumila*, page 390: *PalDat* Picture: *Argyranthemum* sp., page 391: Frosch-Radivo, Andrea Picture: *Tanacetum corymbosum*, page 391: Weber, Martina Picture: *Cyanus segetum*, page 391: *PalDat* Picture: *Cyanus segetum*, page 392: *PalDat* Picture: *Centaurea jacea*, page 392: Weber, Martina Picture: *Agrimonia eupatoria*, page 392: *PalDat* Picture: *Ageratum* sp., page 392: Weber, Martina Picture: *Nigella arvensis*, page 392: *PalDat* Picture: *Artemisia vulgaris*, page 392: Weber, Martina Picture: *Chaenorhinum minus*, page 393: *PalDat* Picture: *Plantago maritima*, page 393: *PalDat* Picture: *Clinopodium vulgare*, page 393: *PalDat* Picture: *Acinos alpinus*, page 393: *PalDat* Picture: *Pachypodium succulentum*, page 393: *PalDat* Picture: *Fraxinus excelsior*, page 393: *PalDat* Picture: *Odontites luteus*, page 394: *PalDat* Picture: *Ailanthus altissima*, page 394: *PalDat* Picture: *Androsace maxima*, page 394: *PalDat* Picture: *Adonis aestivalis*, page 394: *PalDat* Picture: *Acer platanoides*, page 394: *PalDat* Picture: *Chaenorhinum minus*, page 394: *PalDat* Picture: *Glaucium flavum*, page 395: *PalDat*

Picture: *Odontites vulgaris*, page 395: *PalDat* Picture: *Ranunculus trichophyllus*, page 395: *PalDat* Picture: *Delphinium elatum*, page 395: *PalDat*

**Service Part** Picture: *Plantago maritima*, page 395: *PalDat* Picture: *Stachys officinalis*, page 395: *PalDat* Picture: *Pistia stratiotes*, page 396: *PalDat* Picture: *Amorphophallus mossambicensis*, page

396: Ulrich, Silvia Picture: *Mentha aquatica*, page 396: *PalDat* Picture: *Spathiphyllum blandum*, page 396: *PalDat* Picture: *Arophyton buchetii*, page 396: *PalDat* Picture: *Homalomena wallisii*, page 396: *PalDat* Picture: *Orobanche hederae*, page 397: *PalDat* Picture: *Thalictrum flavum*, page 397: *PalDat* Picture: *Ambrosia artemisiifolia*, page 397: *PalDat* Picture: *Corylus avellana*, page 398: *PalDat* Picture: *Corylus avellana*, page 398: *PalDat* Picture: *Syringa vulgaris*, page 398: *PalDat* Picture: *Syringa vulgaris*, page 398: *PalDat* Picture: *Acinos alpinus*, page 398: *PalDat* Picture: *Acinos alpinus*, page 398: *PalDat* Picture: *Brassica napus*, page 399: *PalDat* Picture: *Chenopodium album*, page 399: *PalDat* Picture: *Trisetum flavescens*, page 399: *PalDat* Picture: *Cereus* sp., page 399: *PalDat* Picture: *Avena sativa*, page 399: *PalDat* Picture: *Ornithogalum nutans*, page 399: *PalDat* Picture: *Apium nodiflorum*, page 400: *PalDat* Picture: *Apium nodiflorum*, page 400: *PalDat* Picture: *Quercus robur*, page 400: *PalDat* Picture: *Acinos alpinus*, page 401: *PalDat* Picture: *Acinos alpinus*, page 401: *PalDat*


Picture: *Centaurea* sp., page 404: Weber, Martina Picture: *Pelargonium carnosum*, page 404: Weber, Martina

Picture: *Citrus aurantiifolia*, page 404: Weber, Martina


Picture: *Galium* sp., page 411: Weber, Martina Picture: *Jasminum nudiflorum*, page 412: *PalDat* Picture: *Nigella arvensis*, page 412: *PalDat* Picture: *Melampyrum nemorosum*, page 412:

*PalDat* Picture: *Betonica officinalis*, page 412: *PalDat* Picture: *Anemone ranunculoides*, page 412: *PalDat* Picture: *Consolida regalis*, page 412: *PalDat* Picture: *Salvia glutinosa*, page 413: *PalDat* Picture: *Linaria vulgaris*, page 413: *PalDat* Picture: *Salix x fragilis*, page 413: *PalDat* Picture: *Mentha aquatica*, page 413: *PalDat* Picture: *Syringa vulgaris*, page 413: *PalDat* Picture: *Scilla bifolia*, page 413: *PalDat* Picture: *Apium nodiflorum*, page 414: *PalDat* Picture: *Convolvulus tricolor*, page 414: *PalDat* Picture: *Veronica spicata*, page 414: *PalDat*

Picture: *Sambucus nigra*, page 414: *PalDat* Picture: *Atriplex tatarica*, page 414: *PalDat*


Picture: *Oenothera biennis*, page 419: *PalDat* Picture: *Oenothera biennis*, page 419: *PalDat* Picture: *Clarkia pulchella*, page 419: *PalDat* Picture: *Clarkia unguiculata*, page 419: *PalDat* Picture: *Circaea lutetiana*, page 420: Weber, Martina Picture: *Circaea lutetiana*, page 420: *PalDat* Picture: *Fuchsia magellanica*, page 420: *PalDat* Picture: *Lopezia racemosa*, page 420: Weber, Martina Picture: *Clarkia purpurea*, page 420: *PalDat* Picture: *Rhododendron hirsutum*, page 420: *PalDat* Picture: *Trisetum flavescens*, page 421: *PalDat* Picture: *Corylus avellana*, page 421: *PalDat* Picture: *Metasequoia glyptostroboides*, page 421: Weber, Martina Picture: Cupressaceae, page 421: Grímsson, Friðgeir Picture: Cupressaceae, page 421: Grímsson, Friðgeir Picture: *Stellaria graminea*, page 421: *PalDat* Picture: *Chamaecyparis lawsoniana*, page 422: *PalDat* Picture: *Chamaecyparis lawsoniana*, page 422: *PalDat* Picture: *Gladiolus illyricus*, page 422: *PalDat* Picture: *Quercus robur*, page 422: PalDat Picture: *Ruspolia seticalyx*, page 422: *PalDat* Picture: *Ruspolia seticalyx*, page 422: *PalDat* Picture: *Ephedra foeminea*, page 423: *PalDat* Picture: *Odontites luteus*, page 423: *PalDat* Picture: *Nigella arvensis*, page 423: *PalDat* Picture: *Ficaria verna*, page 423: *PalDat* Picture: *Tilia platyphyllos*, page 423: *PalDat* Picture: *Sauromatum venosum*, page 423: *PalDat* Picture: *Akebia quinata*, page 424: *PalDat* Picture: *Atriplex sagittata*, page 424: *PalDat* Picture: *Acacia binervia*, page 424: *PalDat* Picture: *Justicia brandegeeana*, page 424: *PalDat* Picture: *Calluna vulgaris*, page 424: *PalDat* Picture: *Acacia binervia*, page 424: *PalDat* Picture: *Cyperus longus*, page 425: *PalDat* Picture: *Justicia brandegeeana*, page 425: *PalDat* Picture: *Delonix regia*, page 425: *PalDat* Picture: *Scaevola aemula*, page 425: *PalDat* Picture: *Orobanche hederae*, page 425: *PalDat*

Picture: *Kalmia latifolia*, page 419: *PalDat* Picture: *Ledum palustre*, page 419: *PalDat*


#### **Service Part Chapter: "Pollen Class"**

Picture: *Iris bucharica*, page 430: *PalDat* Picture: *Ibicella lutea*, page 430: *PalDat* Picture: *Bunias orientalis*, page 430: *PalDat* Picture: *Corylopsis glabrescens*, page 430: *PalDat* Picture: *Viola alba*, page 430: *PalDat* Picture: *Orlaya grandiflora*, page 430: *PalDat* Picture: *Zeylanidium subulatum*, page 431: *PalDat* Picture: *Podostemum munnarense*, page 431: *PalDat* Picture: *Hedychium gardnerianum*, page 431: *PalDat* Picture: *Sauromatum venosum,* page 431: *PalDat* Picture: *Prenanthes purpurea*, page 431: *PalDat* Picture: *Gazania* sp., page 431: *PalDat* Picture: *Ephedra distachya*, page 432: *PalDat*

Picture: *Welwitschia mirabilis*, page 432: *PalDat*

© Springer International Publishing AG 2018

M. Špero, H. Vavro, *Neuroradiology - Expect the Unexpected*, DOI https://doi.org/10.1007/978-3-319-73482-8

Picture: *Acacia dealbata*, page 432: *PalDat* Picture: *Chelonanthus purpurascens*, page 432: *PalDat*

Picture: *Silene flos-cuculi*, page 432: *PalDat* Picture: *Pachypodium saundersii*, page 432: *PalDat* Picture: *Abies cephalonica*, page 433: *PalDat* Picture: *Pinus mugo*, page 433: *PalDat* Picture: *Berberis thunbergii*, page 433: *PalDat* Picture: *Thunbergia alata*, page 433: *PalDat* Picture: *Sandersonia vaurantiaca*, page 433: *PalDat*

Picture: *Gagea villosa*, page 433: *PalDat* Picture: *Acca sellowiana*, page 434: *PalDat* Picture: *Primula farinosa*, page 434: *PalDat* Picture: *Moneses uniflora*, page 434: *PalDat* Picture: *Neottia ovata*, page 434: *PalDat* Picture: *Sesleria caerulea*, page 434: *PalDat* Picture: *Typha laxmannii,* page 434: *PalDat*

#### **Index**

**A** *Abeliophyllum distichum*, 326 *Abies* sp., 50, 51, 385 *A. cephalonica*, 68, 188, 190, 191, 433 *A. concolor*, 193 *A. nordmanniana*, 191 fossil, Quaternary, Austria, 51 fossil, middle Miocene, Austria, 385 *Abutilon* sp. *A. megapotamicum*, 310 *A. theophrasti*, 156 *Acacia* sp., 145, 146 *A. binervia*, 424 *A. dealbata*, 146, 276, 432 *Acalypha macrostachya*, 208 Acanthaceae, 13, 43, 51, 54, 164, 172–175, 177, 181, 185, 221, 229, 233, 238, 242, 243, 261, 263, 268, 271, 273, 277, 278, 280, 296, 297, 300, 301, 315, 319–321, 326, 327, 344, 345, 347, 348, 350, 352, 422, 424, 425, 433 *Acantholimon glumaceum*, 326 *Acca sellowiana*, 434 *Acer* sp. *A. negundo*, 359 *A. platanoides*, 394 *A. pseudoplatanus*, 160, 361, 365 fossil, middle Miocene, Austria, 228 Achariaceae, 157, 225 *Acicarpha tribuloides*, 163, 170, 186 *Acinos alpinus*, 28, 30, 211, 393, 398, 401 *Acokanthera oblongifolia*, 409 *Aconitum lycoctonum*, 219 *Actaea spicata*, 426 *Adansonia gregorii*, 181 *Adenanthos sericeus*, 180, 200 *Adenia fruticosa*, 352 *Adhatoda schimperiana*, 233 *Adonis aestivalis*, 381, 394 Adoxaceae, 177, 304, 319, 371, 414 *Aechmea* sp. *A. allenii*, 263, 353 *A. ampla*, 318 *A. araneosa*, 317 *A. azurea*, 269, 348 *A. caesia*, 214 *A. dealbata*, 158 *A. drakeana*, 184 *A. fulgens*, 266 *A. mentosa*, 266 *A. subintegerrima*, 224 *A. tomentosa,* 266 *Aerides multiflora*, 150 *Aesculus* sp., 173

*A. carnea*, 219 *A. flava*, 177 *A. hippocastanum*, 216, 362 *Aesculus x carnea*, 215 *Aetanthus* sp. *A. coriaceus*, 98 *A. macranthus*, 290 *Agapetes variegata*, 408 *Agastache mexicana*, 300 Agavaceae, 44, 45 *Agave asperrima*, 253 *Ageratum* sp., 392 *Aglaia elliptica*, 407 *Aglaodorum griffithii*, 245 *Agrimonia eupatoria*, 392 *Agrostemma githago*, 252 *Ailanthus altissima*, 367, 382, 394 Aizoaceae, 341 *Ajuga* sp. *A. genevensis*, 349 *A. reptans*, 30 *Akebia quinata*, 373, 424 *Alangium* sp., 383 fossil, 383 *Albizia julibrissin*, 81, 146 *Alisma* sp. *A. lanceolatum*, 58, 271 *A. plantago-aquatica*, 212 fossil, Miocene, China, 340 Alismataceae, 58, 182, 212, 269, 271, 272, 340 fossil, Miocene, China, 340 *Alkanna corcyrensis*, 180, 183 *Alliaria petiolata*, 415 *Allium* sp. *A. flavum*, 362 *A. oleraceum*, 158 *A. paradoxum*, 40 *A. sphaerocephalum*, 285 *A. ursinum*, 54, 56, 282 *Alnus* sp., 43, 407 *A. viridis*, 268 *A. glutinosa*, 201, 211, 267, 279, 407 *A. incana*, 407 *Alocasia* sp, 99 *A. acuminata*, 310 *A. odora*, 132, 244 *Alopecurus* sp., 14 Alstroemeriaceae, 325 Altingiaceae, fossil, middle Miocene, Austria, 271 Amaranthaceae, 219, 258, 270, 328, 330, 337, 339, 374, 388, 399, 401, 414, 424 fossil, Miocene, China, 337, 339

Amaryllidaceae, 54, 158, 169, 187, 198, 212, 282, 285, 287, 303, 304, 362 Amborellaceae, 293 *Amborella trichopoda*, 293 *Ambrosia artemisiifolia*, 47, 311, 381, 397, 411 *Ambrosina* sp., 15 *A. bassii*, 158, 244 *Amherstia nobilis*, 361, 369 *Amorphophallus* sp. *A. interruptus*, 101, 158, 365 *A. krausei*, 82, 100 *A. lacourii*, 345 *A. longituberosus*, 78, 83 *A. mangelsdorffii*, 46 *A. mossambicensis*, 396 *A. serrulatus*, 345, 365 *A. stuhlmannii*, 377, 411 *A. taurostigma*, 31 *A. yunnanensis*, 180 *Amphitecna macrophylla*, 257 *Amsonia ciliata*, 170, 265 *Amydrium medium*, 389 *Anacamptis pyramidalis*, 152 Anacardiaceae, 221, 235, 366, 369 *Anaphyllopsis americana*, 284 *Anchomanes welwitschii*, 29, 100 *Anchusa* sp. *A. officinalis*, 166 *A. cretica*, 250 *Andrographis paniculata*, 301 *Androlepis skinneri*, 142, 347, 405 *Androsace maxima*, 394 *Anemone* sp. *A. hortensis*, 195 *A. pratensis*, 335 *A. ranunculoides*, 412 *A. transsilvanica*, 255 Annonaceae, 44, 45, 57, 77, 144, 145, 406 *Annona muricata*, 44, 45, 406 *Antennaria dioica*, 312 *Anthericum* sp. *A. liliago*, 401 *A. ramosum*, 324 *Anthurium* sp., 15 *A. gracile*, 33, 356, 370 *A. ovatifolium*, 156 *A. radicans*, 204 *Anthyllis vulneraria*, 184, 208, 346 *Aphelandra arborea*, 173 *Aphyllanthes monspeliensis*, 278 Apiaceae, 24, 26, 32, 34, 62, 119, 161, 164, 165, 174, 176, 194, 201, 213, 223, 236, 358, 359, 381, 400, 403, 414, 430 *Apium nodiflorum*, 119, 213, 381, 400, 414 *Apoballis acuminatissima*, 402 Apocynaceae, 137, 144, 170, 222, 265, 346, 393, 402, 409, 432 Aponogetonaceae, 357 *Aponogeton masoalaensis*, 357 Aquifoliaceae, 302, 373 fossil, Miocene, China, 373 Araceae, 13, 15, 16, 24, 26, 27, 29, 31–34, 44, 46, 77, 79, 82, 83, 99–101, 132, 137, 142, 144, 156, 158, 180, 183, 196, 197, 204, 244, 245, 274, 276, 284, 293, 296, 309, 310, 313, 318, 339, 342, 344, 345, 356, 365, 369, 370, 377, 384, 385, 389, 390, 396, 402, 405, 411, 423, 431 Araliaceae, 236, 248, 321, 324, 335 *Aratitiyopea lopezii*, 302

*Arbutus unedo*, 140, 260 *Arceuthobium* sp., fossil, Miocene, China, 340 *Ardisia* sp. *A. crenata*, 202, 291 *A. elliptica*, 320 Arecaceae, 13, 286, 288, 313 fossil, early Miocene, South Africa, 288 *Arenaria* sp. *A. ciliata*, 271 *A. serpyllifolia*, 216 *Arenga pinnata*, 313 *Argyranthemum* sp., 391 Aristolochiaceae, 52, 157, 246, 315, 321, 375, 402 *Aristolochia* sp. *A. arborea*, 52, 246, 375, 402 *A. clematitis*, 402 *A. manshuriensis*, 315 *A. tricaudata*, 375 *Armeria* sp. *A. alpina*, 61 *A. maritima*, 357 *A. pinifolia*, 327 *Arophyton buchetii*, 396 *Artemisia* sp., 162 *A. pontica*, 160, 195 *A. vulgaris*, 388, 392 *Aruncus dioicus*, 234 *Asarum europaeum*, 321 *Ascarina lucida*, 370 Asclepiadaceae, 13, 57, 149, 153 *Asclepias* sp., 6 *Asimina triloba*, 144, 406 Asparagaceae, 179, 186, 187, 195, 253, 278, 281, 283, 285, 292, 318, 324, 325, 331, 334, 349, 399, 401, 413 *Asperugo procumbens*, 273 *Asperula tinctoria*, 176, 280 *Asphodeline lutea*, 196, 282 *Asphodelus fistulosus*, 198, 285 *Aspidistra* sp. *A. elatior*, 331 *A. locii*, 334 Asteraceae, 10, 39, 47, 49, 73, 75, 132, 133, 160–162, 164, 174, 181, 195, 212, 214, 218, 223, 234, 236, 237, 240, 260, 261, 309–314, 328–330, 381, 386, 388, 391, 392, 397, 404, 411, 431 fossil, Quaternary, Austria, 329 *Aster amellus*, 311 *Asterostigma lividum*, 29 *Astragalus* sp. *A. onobrychis*, 174 *A. tragacantha*, 250 *Asyneuma canescens*, 266 *Atriplex* sp. *A. patula*, 258 *A. sagittata*, 374, 424 *A. tatarica*, 401, 414 *Austobuxus nitidus*, 50 Austrobaileyaceae, 180 *Austrobaileya scandens*, 180 *Avena sativa*, 254, 399 *Axinaea lehmannii*, 410

#### **B**

*Babiana ecklonii*, 251, 336 Balsaminaceae, 158, 168, 221, 229, 319, 327, 350 *Banisteria muricata*, 256, 306 *Barringtonia asiatica*, 162, 290

*Basella alba*, 172, 182 Basellaceae, 172, 182 *Bassia scoparia*, 270, 388 Begoniaceae, 248, 363 *Begonia heracleifolia*, 248, 363 *Bellis perennis*, 39, 161 *Beloperone guttata*, 348 Berberidaceae, 227, 277, 278, 290, 333, 369, 390, 433 *Berberis* sp. *B. amurensis*, 278 *B. nervosa*, 290 *B. thunbergii*, 433 *B. vulgaris*, 277, 390 *Beschorneria yuccoides*, 44, 45 *Besleria hirsuta*, 316 *Bessera elegans*, 281, 285, 325 *Betonica officinalis*, 29, 30, 412 Betulaceae, 29, 43, 132, 168, 201, 209, 211, 220, 264, 267, 268, 279, 359, 360, 372, 381, 389, 398, 407, 421 fossil, Miocene, China, 372 *Betula* sp., 220, 372 *B. humilis*, 132, 209, 220, 264, 360 *B. pendula*, 88, 220, 264 fossil, Miocene, China, 372 *Bifora radians*, 176, 194, 223 Bignoniaceae, 142, 257, 278, 306 *Bignonia magnifica*, 278 *Billardiera heterophylla*, 353 *Billbergia* sp. *B. macrocalyx*, 216 *B. porteana*, 159 *B. pyramidalis*, 185 *B. seidelii*, 178, 196, 324 *Blackstonia perfoliata*, 366 *Blumenbachia hieronymi*, 249 *Bomarea hirsuta*, 325 Boraginaceae, 43, 78, 166, 174–178, 180, 183, 186, 216, 222, 236–239, 241–243, 249, 250, 265, 268, 273, 280, 291, 337, 339, 342, 343, 346 *Borago* sp. *B. officinalis*, 280 *B. pygmaea*, 222, 339 *Bougainvillea* sp., 199, 319 *Brassavola cucullata*, 150, 417 Brassicaceae, 13, 24, 28, 226, 319, 326, 348, 387, 399, 415, 430 *Brassica* sp. *B. napus*, 28, 399, 415 *B. nigra*, 415 *Brillantaisia owariensis*, 345 Bromeliaceae, 33, 58, 142, 158, 159, 170, 178, 179, 184, 185, 196, 203, 214, 216, 224, 263, 266, 269, 283, 286, 317, 318, 324, 347, 348, 351, 353, 383, 405 *Bromus erectus*, 292 *Broussonetia papyrifera*, 263 *Brugmansia suaveolens*, 184, 369 *Buglossoides* sp. *B. arvensis*, 239 *B. purpurocaerulea*, 175 *Bunias orientalis*, 430 *Bupleurum rotundifolium*, 164, 201 Butomaceae, 249 *Butomus umbellatus*, 249 Buxaceae, 270, 350, 355–357, 383 fossil, Upper Cretaceous, USA, 355 *Buxus sempervirens*, 270, 350

#### **C**

Cabombaceae, 282, 363 *Cabomba palaeformis*, 282, 363 Cactaceae, 167, 228, 232, 255, 257, 306, 328, 330, 350, 370, 399 *Caladenia latifolia*, 150, 416 *Caldesia parnassifolia*, 272 *Calendula officinalis*, 310 *Calla palustris*, 75, 79, 100, 142 *Calliandra* sp. *C. emarginata*, 72 *C. tergemina*, 74, 145, 375 *Callisia fragrans*, 331, 336 *Callistemon* sp. *C. coccineus*, 164, 291 *C. comboynensis*, 377 *Callitriche* sp., 76, 446 *C. palustris*, 80 *C. polymorpha*, 80 *C. stagnalis*, 203, 308 *Calluna vulgaris*, 142, 144, 180, 322, 424 *Calolisianthus pendulus*, 141, 406 *Calycanthus floridus*, 75 Calyceraceae, 163, 170, 186 *Calystegia sepium*, 215, 269 *Camellia japonica*, 252 *Campanula* sp. *C. alpina*, 266, 313 *C. fenestrellata*, 171 *C. garganica*, 267 *C. persicifolia*, 333 *C. rapunculoides*, 267 *C. saxatilis*, 264 Campanulaceae, 171, 175, 230, 231, 264, 266, 267, 280, 313, 333 *Canistrum camacaense*, 317 Cannabaceae, 323 *Cantua buxifolia*, 322 Caprifoliaceae, 163, 165, 169, 210, 221, 226, 252, 265, 311–313, 340 fossil, Miocene, China, 340 *Cardamine pratensis*, 348 *Cardiospermum* sp. *C. corindum*, 69 *C. halicacabum*, 184, 208 *Carduus acanthoides*, 309 *Carex* sp., 135 *C. alba*, 195, 212 *C. filiformis*, 338 *C. remota*, 272 *Carpinus betulus*, 29, 359 *Carthamus lanatus*, 75, 218 *Carya* sp., fossil, middle Miocene, Austria, 168, 264 Caryophyllaceae, 51, 53, 166, 167, 171, 181, 182, 195, 216, 251, 252, 258, 269, 271, 330, 332, 339, 421, 432 fossil, Miocene, China, 339 *Cassia pulcherrima*, 291 *Catalpa bignonioides*, 306 *Catopsis floribunda*, 283 *Cedrus atlantica*, 247 Celastraceae, 77, 171 *Centaurea* sp., 404 *C. jacea*, 240, 392 *C. scabiosa*, 236 *C. segetum*, 260 *Cephalanthera longifolia*, 148, 292, 353 *Cephalopentandra ecirrhosa*, 321 *Cephalostemon riedelianus*, 275 Cephalotaxaceae, 46

*Cephalotaxus* sp., 46 *Cerastium dubium*, 182 *Ceratostigma plumbaginoides*, 226, 373 *Ceratozamia* sp. *C. kuesteriana*, 285 *C. mexicana*, 360 Cercidiphyllaceae, 272, 371 fossil, Miocene, China, 371 *Cercidiphyllum* sp. *C. japonicum*, 272 fossil, Miocene, China, 371 *Cereus* sp., 399 *Cerinthe minor*, 177, 238 *Ceropegia sandersonii*, 153 *Chaenarrhinum* sp., 16 *Chaenomeles sinensis*, 362 *Chaenorhinum minus*, 178, 386, 393, 394 *Chamaecyparis lawsoniana*, 422 *Chamaedorea microspadix*, 286 *Cheilocostus speciosus*, 347 *Chelonanthus* sp. *C. alatus*, 140, 325 *C. purpurascens*, 145, 432 Chenopodiaceae, 156 *Chenopodium* sp. *C. album*, 399, 401, 414 *C. hybridum*, 219 Chloranthaceae, 156, 289, 370 *Chlorella* sp., 49 *Chlorospatha* sp. *C. dodsonii*, 142, 144, 245 *C. hannoniae*, 405 *C. kolbii*, 137, 144, 405 *C. oblongifolia*, 405 *C. pubescens*, 369, 405 *Cichorium intybus*, 240, 328, 329, 386 *Circaea lutetiana*, 165, 209, 358, 420 *Cirsium oleraceum*, 181, 236 Cistaceae, 33, 58, 182, 355, 364, 367 *Cistus* sp. *C. clusii*, 367 *C. creticus*, 58 *Citrus* sp. *C. aurantiifolia*, 404 *C. swinglei*, 237 *Clarkia* sp. *C. pulchella*, 217, 323, 419 *C. purpurea*, 170, 323, 420 *C. unguiculata*, 170, 185, 265, 419 *Claytonia perfoliata*, 277, 332 *Cleistocactus straussii*, 228 *Clematis heracleifolia*, 161 *Clerodendrum thomsoniae*, 218 *Clethra arborea*, 334 Clethraceae, 334 *Clinopodium vulgare*, 212, 230, 393 *Cobaea scandens*, 258, 269, 387 *Codiaeum* sp., 62 *Codonopsis pilosula*, 231 *Coelogyne fimbriata*, 151 *Coffea arabica*, 373 Colchicaceae, 263, 287, 433 *Colchicum autumnale*, 263 Coleochaete, 49 *Colocasia antiquorum*, 34 *Columnea magnifica*, 176

*Colutea arborescens*, 173, 223, 342 Combretaceae, 243, 273 *Combretum fruticosum*, 243 Commelinaceae, 70, 159, 218, 286, 331, 334, 336, 377 *Commelina erecta*, 159, 218, 377 *Consolida regalis*, 28, 412 Convolvulaceae, 80, 215, 216, 228, 230, 232, 269, 312, 314, 330, 414 *Convolvulus* sp. *C. arvensis*, 228 *C. tricolor*, 216, 414 *Cordia cylindrostachya*, 265 *Cordyline fruticosa*, 285 Coriariaceae, 204, 222 *Coriaria* sp. *C. myrtifolia*, 222 *C. nepalensis*, 204 *Coris monspeliensis*, 250 Cornaceae, 383 *Corydalis* sp. *C. cava*, 255, 375 *C. cheilanthifolia*, 184 *C. ophiocarpa*, 156 *Corylopsis* sp. *C. glabrescens*, 430 *C. platypetala*, 227 *Corylus* sp., 43 *C. avellana*, 168, 220, 381, 398, 421 *C. colurna*, 389 Costaceae, 347 *Costus barbatus*, 257 *Cotinus coggygria*, 366 Crassulaceae, 161, 165, 346, 358 *Crataegus laevigata*, 364 *Crepis biennis*, 329 *Crinum x amabile*, 287 *Crocus speciosus*, 278 *Crossandra flava*, 174, 177, 181 *Croton triqueter*, 307, 308 *Croton* type, 10 *Cruciata laevipes*, 230 *Cryptanthus bromelioides*, 33 *Cryptogamma crispa*, 44 *Cryptomeria* sp. *C. japonica*, 259 fossil, middle Miocene, Austria, 259 *Cucumis* sp. *C. melo*, 264 *C. sativus*, 342 Cucurbitaceae, 78, 82, 177, 210, 231, 239, 252, 257, 264, 319, 321, 342, 348, 349, 353, 364 *Cucurbita pepo*, 252 *Cunninghamia lanceolata*, 200, 259 *Cunonia capensis*, 165, 223 Cunoniaceae, 165, 223 *Cuphea* sp. *C. llavea*, 364 *C. procumbens*, 169, 291 Cupressaceae, 50, 53, 200, 259, 292, 421, 422 fossil, middle Miocene, Austria, 259 fossil, Miocene, China, 421 *Cuscuta lupuliformis*, 76, 80 *Cutandia* sp., 14 *Cyanastrum cordifolium*, 376 *Cyanthillium cinereum*, 330 *Cyanus segetum*, 391, 392 *Cynodon dactylon*, 296

*Cynoglossum officinale*, 242, 273 Cyperaceae, 135, 136, 183, 195, 212, 272, 336, 338, 425 fossil, Miocene, China, 338 *Cyperus longus*, 183, 425 *Cyrtopodium polyphyllum*, 417 *Cyrtosperma* sp., 15 *C. beccarianum*, 293, 318 Cytinaceae, 246 *Cytinus hypocistis*, 246 *Cytisus nigricans*, 335

#### **D**

*Dacrycarpus* sp., 50 *D. dacrydioides*, 188, 193 *Dactylis* sp., 14, 133 *Dactylorhiza maculata*, 417 *Dalechampia spathulata*, 222 *Daphne* sp. *D. cneorum*, 307 *D. laureola*, 307 *D. tangutica*, 308 *Delonix regia*, 425 *Delphinium elatum*, 395 *Dianella* sp. *D. caerulea*, 288 *D. intermedia*, 288 *D. tasmanica*, 70, 288, 337 *Dianthus carthusianorum*, 251 *Dictamnus albus*, 241 *Didymaea mexicana*, 229 *Dieffenbachia humilis*, 390, 402 *Dionaea muscipula*, 251, 321 *Dioon edule*, 360 *Dioscorea* sp., 75 Dipsacaceae, 46 *Dipsacus fullonum*, 165 *Discocleidion rufescens*, 248 *Dodecatheon meadia*, 290 *Dorstenia contrajerva*, 203, 210 Doryanthaceae, 214, 282 *Doryanthes palmeri*, 214, 282 *Dracocephalum austriacum*, 279, 280 *Dracontium asperum*, 27, 196 *Dracunculus* sp., 15 *D. vulgaris*, 296 *Drimys granadensis*, 293 Droseraceae, 138, 141, 144, 251, 268, 310, 321, 335, 340, 406 *Drosera* sp. *D. binata*, 141, 340 *D. kansaiensis*, 268, 335 *D. rotundifolia*, 406 *D. scorpioides*, 138, 310, 406 *Dysphania ambrosioides*, 258

#### **E**

*Ecballium elaterium*, 349 *Echinopepon wrightii*, 239 *Echinops* sp. *E. exaltatus*, 261 *E. ritro*, 164 *Echium* sp. *E. italicum*, 180 *E. plantagineum*, 183 *E. vulgare*, 236 *Eichhornia crassipes*, 75, 287 Elaeagnaceae, 178, 199, 211, 223, 335 *Elaeagnus* sp. *E. angustifolia*, 178, 199, 211, 223 *E. rhamnoides*, 335 *Elatostema ambiguum*, 267 Ephedraceae, 183, 344, 423, 432 fossil, Miocene, Austria, 344 fossil, Miocene, China, 344 *Ephedra* sp. *E. distachya*, 344, 432 *E. foeminea*, 183, 423 fossil, Miocene, Austria, 344 fossil, Miocene, China, 344 *Epilobium* sp. *E. angustifolium*, 210, 418 *E. dodonaei*, 418 *E. fleischeri*, 418 *E. hirsutum*, 138, 224, 418 *E. palustre*, 58, 339 *E. parviflorum*, 140, 418 *Epipactis* sp. *E. helleborine*, 137, 352 *E. muelleri*, 416 *Eranthemum wattii*, 326 *Eranthis hyemalis*, 426 *Erdtmanipollis* sp., fossil, Upper Cretaceous, USA, 355 *Eremogone procera*, 182 *Eremurus* sp. *E. robustus*, 166, 187 *E. thiodanthus*, 198 Ericaceae, 42, 137–142, 144, 160, 176, 179, 180, 201, 202, 234, 260, 315, 316, 322, 334, 406, 408, 409, 419, 420, 424, 434 *Erica* sp. *E. arborea*, 179, 201 *E. herbacea*, 137, 234, 315 *E. pageana*, 334 *Erodium cicutarium*, 366 *Erophila verna*, 319 *Erysimum odoratum*, 226 *Erythrochiton brasiliensis*, 298 *Erythronium dens-canis*, 253, 351 *Eucharis grandiflora*, 212 *Eucommia* sp., fossil, Middle Eocene, Western Greenland, 43 *Eucommiidites* sp., fossil, Lower Cretaceous of U.S.A., 71 *Eupatorium cannabinum*, 212 Euphoebiaceae, 235 *Euphorbia* sp. *E. helioscopia*, 343 *E. palustris*, 342 *E. peplus*, 235, 248 *E. tithymaloides*, 261 *Exospermum stipitatum*, 89–92

#### **F**

Fabaceae, 33, 74, 77, 81, 132, 133, 145, 146, 173–175, 184, 208, 215, 218, 223, 234, 235, 240, 241, 250, 276, 291, 316, 320, 335, 342, 343, 346, 349–353, 358, 361, 369, 375, 424, 425 fossil, middle Miocene, Austria, 235 Fagaceae, 29, 41, 43, 334, 358, 360, 374, 381, 400, 422, 426 fossil, Miocene, Austria, 358 fossil, Quaternary, Austria, 41, 360 *Fagopyrum* sp., 235 *Fagus* sp., 41 fossil, Miocene, Austria, 358 fossil, Quaternary, Austria, 360 *Fallopia convolvulus*, 55 *Fargesia* sp., 14

*Fatsia japonica*, 236, 248, 321 *Ficaria verna*, 215, 423 *Filarum manserichense*, 31 Fouquieriaceae, 250, 320, 367 *Fouquieria* sp. *F. columnaris*, 250 *F. macdougalii*, 320, 367 *Fraxinus* sp. *F. excelsior*, 227, 228, 354, 382, 393 *F. ornus*, 157 fossil, Miocene, China, 371 *Freesia* sp., 281 *Frerea indica*, 153 *Fritillaria* sp. *F. meleagris*, 324 *F. pontica*, 194, 281 *Fuchsia* sp. *F. magellanica*, 106, 420 *F. paniculata*, 185 *Fumana procumbens*, 355 *Fumaria officinalis*, 209, 257

#### **G**

*Gagea* sp. *G. lutea*, 197 *G. villosa*, 217, 433 *Galanthus nivalis*, 187, 282 *Galeopsis tetrahit*, 159, 217, 380 *Galinsoga ciliata*, 309 *Galium* sp., 411 *G. glaucum*, 231 *G. lucidum*, 157, 231, 336 *G. odoratum*, 27, 32, 194, 231 *Garcia nutans*, 308 *Gardenia thunbergia*, 236 *Gaultheria myrsinoides*, 316 *Gaura lindheimeri*, 187 *Gazania* sp., 328, 431 *G. rigens*, 223 Gelsemiaceae, 366 *Gelsemium sempervirens*, 366 *Genlisea violacea*, 237 *Gennaria diphylla*, 147, 152 Gentianaceae, 77, 140, 141, 145, 265, 267, 306, 325, 361, 366, 368, 383, 406, 432 *Gentiana* sp. *G. acaulis*, 368 *G. lutea*, 361 *Gentianella austriaca*, 366 Geraniaceae, 162, 213, 303, 304, 356, 366–368, 374, 404 *Geranium* sp. *G. pratense*, 304, 356 *G. reuteri*, 356 *G. robertianum*, 213, 303, 374 *G. sibiricum*, 304 Gesneriaceae, 176, 316–318 *Geum reptans*, 363 Ginkgoaceae, 157, 196 *Ginkgo* sp., 50 *G. biloba*, 157, 196 *Gladiolus illyricus*, 254, 422 *Glaucium flavum*, 395 *Globba schomburgkii*, 384 *Glossoloma ichthyoderma*, 317 Gnetaceae, 246 *Gnetum gnemon*, 246

*Gomphrena celosioides*, 330 *Gonatopus* sp. *G. angustus*, 274, 342, 385 *G. boivinii*, 276 Goodeniaceae, 62, 241, 425 *Goodyera repens*, 152 *Grevillea banksii*, 202 Grossulariaceae, 257 Gunneraceae, 160 *Gunnera tinctoria*, 160 *Guzmania elvallensis*, 203, 286 *Gynura scandens*, 314

#### **H**

*Habenaria* sp., 81 *H. tridactylites*, 416 *Hacquetia epipactis*, 26 *Haemanthus coccineus*, 198 Haemodoraceae, 198, 286, 322 *Hakea kippistiana*, 169, 321 Haloragaceae, 210, 266, 341 fossil, Miocene, China, 341 *Haloragis erecta*, 341 Hamamelidaceae, 227, 430 *Haraella odorata*, 151 *Harpochilus neesianus*, 352 *Hedera helix*, 306 *Hedychium gardnerianum*, 346, 431 *Hedyosmum* sp. *H. brasiliense*, 289 *H. scaberrimum*, 156, 289 *Helianthemum nummularium*, 364 *Helianthium bolivianum*, 269 *Helianthus annuus*, 313 *Helicodiceros muscivorus*, 411 Heliconiaceae, 170, 179, 183, 199, 200, 293 *Heliconia* sp., 170, 179, 199, 293 *H. rostrata*, 183 *H. stricta*, 200 *Heliohebe raoulii*, 340 *Helleborus foetidus*, 371 *Heloniopsis kawanoi*, 331 *Hemigraphis* sp., 10 *H. primulaefolia*, 344 *Hemiptelia* sp., fossil, Miocene, China, 323 *Herminium monorchis*, 147, 152 *Herniaria* sp. *H. alpina*, 330 *H. glabra*, 166, 182 *Hibiscus* sp. *H. schizopetalus*, 200, 314 *H. trionum*, 309 *Hieracium hoppeanum*, 234, 311, 328 Hippocrateaceae, 77 *Hippocrepis emerus*, 343 *Hohenbergia stellata*, 318 *Homalomena wallisii*, 396 *Hordeum bulbosum*, 337 *Hoya* sp. *H. carnosa*, 153 *H. multiflora*, 153 *Humulus lupulus*, 323 *Hyacinthoides italica*, 187 Hydrocharitaceae, 314 *Hymenocallis* sp.

*H. littoralis*, 304 *H. tubiflora*, 169, 198, 303 *Hypecoum* sp. *H. imberbe*, 162, 225 *H. procumbens*, 225 *Hypoestes phyllostachya*, 164 *Hyptis suaveolens*, 301

#### **I**

*Ibicella lutea*, 305, 430 *Ilex* sp., 93, 302 fossil, Miocene, China, 373 *I. aquifolium*, 302 *Impatiens* sp., 168 *I. columbaria*, 221 *I. glandulifera*, 168, 229, 350 *I. parviflora*, 158, 229, 319, 327 *Ipomoea* sp. *I. batatas*, 269, 314 *I. caerulea*, 330 *I. purpurea*, 312 Iridaceae, 28, 48, 171, 251, 254, 275, 278, 281, 283, 302, 305, 306, 336, 354, 390, 422, 430 *Iris* sp. *I. bucharica*, 305, 430 *I. domestica*, 281, 354 *I. graeberiana*, 306 *I. histrioides*, 275 *I. planifolia*, 302 *I. pseudacorus*, 281, 283, 354 *I. pumila*, 28, 48, 171, 390 *I. reichenbachii*, 283 *Irlbachia* sp. *I. pedunculata*, 306 *I. pendula*, 267

#### **J**

*Jasminum nudiflorum*, 32, 354, 412 *Jatropha* sp. *J. multifida*, 304 *J. podagrica*, 307 Juglandaceae, 56, 258, 270, 389 fossil, middle Miocene, Austria, 168, 264, 268 fossil, Miocene, China, 337, 338 *Juglans* sp., 56, 258 fossil, Miocene, China, 337 *J. regia*, 270, 389 Juncaceae, 139, 141, 186, 195, 199, 292, 323 Juncaginaceae, 246, 370 *Juncus jacquinii*, 186 *Juniperus communis*, 292 *Jurinea mollis*, 174 *Justicia* sp. *J. brandegeeana*, 261, 296, 424, 425 *J. carnea*, 173, 181, 233, 297 *J. furcata*, 273 *J. macrantha*, 233 *J. menesii*, 238, 280 *J. procumbens*, 233 *J. xylosteoides*, 233

#### **K**

*Kallstroemia maxima*, 270, 327 *Kalmia latifolia*, 419

*Kickxia spuria*, 214, 331 *Knautia* sp. *K. arvensis*, 265 *K. drymeia*, 169, 252, 313 *Kolkwitzia amabilis*, 163 *Kraussia floribunda*, 234

#### **L**

*Lachenalia aloides*, 195, 283, 324 *Lagerstroemia indica*, 316 Lamiaceae, 13, 16, 17, 27–32, 55, 83, 119, 133, 159, 166, 168, 175, 211–213, 217–219, 226, 229, 230, 251, 255, 279, 280, 299–301, 317, 325, 331, 347–349, 354, 376, 380, 387, 393, 395, 396, 398, 401, 411–413 *Lamiastrum galeobdolon*, 219 *Lamium* sp. *L. album*, 347 *L. maculatum*, 17, 226 *L. purpureum*, 331 *Lamprocapnos spectabilis*, 215 Lardizabalaceae, 373, 424 *Larix* sp., 68 fossil, middle Miocene, Austria, 68 *L. decidua*, 323 *Lathyrus* sp. *L. sylvestris*, 235, 240 *L. tuberosus*, 175 *L. vernus*, 234, 353 Lauraceae, 56, 245 *Lavandula angustifolia*, 317 *Lavatera thuringiaca*, 271, 309 Lecythidaceae, 162, 290, 343 *Ledum palustre*, 315, 419 *Legousia speculum-veneris*, 267, 280 Lentibulariaceae, 161, 237–239, 353 *Leontodon saxatilis*, 10, 328 *Leontodon* type, 10 *Leucadendron* sp. *L. brunioides*, 200 *L. discolor*, 201, 359 *Ligustrum* sp. fossil, middle Miocene, Austria, 156 *L. vulgare*, 387, 404 Liliaceae, 70, 71, 194, 196, 197, 217, 252, 253, 281, 282, 324, 351, 355, 357, 371, 410, 433 *Lilium* sp. *L. candidum*, 196, 355 *L. martagon*, 282, 355 Limnanthaceae, 178, 187, 248, 275 *Limnanthes douglasii*, 178, 187, 248, 275 *Limodorum abortivum*, 324 Linaceae, 57, 60, 302, 303, 372 *Linaria vulgaris*, 413 *Linum* sp., 303 *L. bienne*, 372 *L. capitatum*, 303 *L. flavum*, 57, 60, 302 *Liquidambar* sp., fossil, middle Miocene, Austria, 271 *Liriodendron tulipifera*, 284 *Lithospermum officinale*, 346 Loasaceae, 249 *Lomatogonium carinthiacum*, 383 *Lonicera fragrantissima*, 226, 312 *Lopezia racemosa*, 163, 420 *Lophophora williamsii*, 306, 370 Loranthaceae, 68, 72, 75, 98, 163, 186, 290

*Loranthus europaeus*, 163, 186 Lowiaceae, 246, 346 *Ludisia discolor*, 81, 152, 417 *Ludwigia octovalvis*, 140, 224 *Luffa cylindrica*, 177, 348 *Lumnitzera racemosa*, 273 *Lupinus* sp., 215 *L. polyphyllus*, 350 *Luzula* sp. *L. campestris*, 139, 199, 323 *L. luzuloides*, 292 *L. sylvatica*, 195 *Lycium barbarum*, 362 *Lysichiton americanus*, 197 *Lysimachia* sp. *L. lichiangensis*, 173 *L. nemorum*, 342 *L. punctata*, 240 *L. vulgaris*, 248 Lythraceae, 43, 60, 61, 169, 242, 273, 291, 316, 364 *Lythrum* sp. *L. hyssopifolia*, 242 *L. salicaria*, 57, 60, 273

#### **M**

*Macadamia ternifolia*, 163 *Maclura pomifera*, 265, 340 Magnoliaceae, 284 *Mahonia aquifolium*, 390 *Maianthemum stellatum*, 318 Malpighiaceae, 256, 306 *Malpighia glabra*, 256 *Malus* sp. *M. baccata*, 223 *M. sieboldii*, 62, 63 *M. sylvestris*, 364 Malvaceae, 70, 73, 132, 156, 181, 199, 200, 211, 218, 220, 222, 256, 260, 261, 271, 301, 309, 310, 312, 314, 350, 357, 404, 423 fossil, Miocene, China, 333, 338 *Malva* sp. *M. alcea*, 256 *M. moschata*, 256 *M. neglecta*, 312 *Maripa nicaraguensis*, 232 Martyniaceae, 305, 430 *Maxillaria densa*, 150, 347 *Mayna odorata*, 157, 225 *Medinilla scortechinii*, 243 *Megaskepasma erythrochlamys*, 268, 297 *Melampyrum* sp. *M. arvense*, 201, 228 *M. nemorosum*, 29, 30, 412, 425 *M. pratense*, 32, 333, 387 *M. subalpinum*, 214 Melanthiaceae, 194, 246, 262, 331, 332, 351 *Melastoma sanguineum*, 201 Melastomataceae, 201, 243, 410 Meliaceae, 407 *Melilotus officinalis*, 351 *Melittis melissophyllum*, 301 *Mendoncia albida*, 221, 229, 315 *Mentha aquatica*, 119, 213, 387, 396, 413 Menyanthaceae, 359, 365 *Menyanthes trifoliata*, 365 *Mercurialis perennis*, 219, 357 *Meriania selvaflorensis*, 243

*Merinthopodium neuranthum*, 249 *Merremia umbellata*, 230 *Metasequoia glyptostroboides*, 259, 421 *Microrrhinum minus*, 17 *Microstrobus* sp., 50 Mimosaceae, 57, 72, 140, 296, 424, 432 *Mimosa pudica*, 140, 296 *Mimulus* sp., 277 *M. guttatus*, 277, 278 *Moehringia muscosa*, 195 Molluginaceae, 232 *Mollugo verticillata*, 232 *Moltkia petraea*, 216, 239 *Moneses uniflora*, 141, 315, 434 *Monotropa hypopitys*, 176 *Monstera* sp., 15 *M. deliciosa*, 274 Montiaceae, 277, 332 *Montrichardia* sp., 54, 99 *M. arborescens*, 183 Moraceae, 203, 210, 263, 265, 340 *Morina longifolia*, 46, 210 *Myosotis* sp. *M. alpestris*, 186 *M. arvensis*, 342 *M. palustris*, 78 *M. ramosissima*, 242 *M. scorpioides*, 176 *Myriophyllum spicatum*, 210, 266 *Myrrhis odorata*, 236, 359 Myrtaceae, 68, 164, 291, 377, 434 *Myrtus communis*, 291

#### **N**

*Nandina domestica*, 227 *Napoleonaea imperialis*, 343 Nelumbonaceae, 226 *Nelumbo nucifera*, 226 *Nematanthus strigillosus*, 250, 318 *Neoalsomitra sarcophylla*, 364 *Neottia* sp. *N. nidus-avis*, 293, 416 *N. ovata*, 434 *Neottianthe cucullata*, 417 *Nicandra physalodes*, 186 *Nicotiana tabacum*, 162, 237, 359 *Nigella arvensis*, 392, 403, 412, 423 *Nonea pulla*, 166 *Nuphar lutea*, 179, 196, 217, 281, 284, 310, 311 *Nuskoissporites* sp., 43 Nyctaginaceae, 199, 319 Nymphaeaceae, 74, 79, 139, 179, 196, 217, 276, 281, 284, 298, 310, 311, 405 *Nymphaea* sp., 79 *N. alba*, 74, 298 *Nymphoides peltata*, 359 *Nypa* sp., 13 Nyssaceae, fossil, middle Miocene, Austria, 50 *Nyssa* sp., fossil, middle Miocene, Austria, 50

#### **O**

Ochnaceae, 373 *Ocimum basilicum*, 299 *Oculopollis* sp., fossil, Upper Cretaceous, Hungary, 11 *Odontites* sp.

*O. luteus*, 227, 382, 394, 423 *O. vulgaris*, 395, 408 *Oenothera* sp. *O. biennis*, 208, 419 *O. fruticosa*, 264, 336 Oleaceae, 32, 157, 227, 228, 326, 354, 370, 382, 387, 393, 398, 404, 412, 413 fossil, middle Miocene, Austria, 156 fossil, Miocene, China, 371 *Omphalodes* sp. *O. linifolia*, 243 *O. verna*, 249 Onagraceae, 58, 106, 138–140, 163, 165, 170, 185, 187, 208–210, 217, 224, 264, 265, 323, 336, 339, 358, 418–420 fossil, Miocene, China, 341 *Oncidium maizaefolium*, 151 *Onosma visianii*, 178, 291 *Ophiorrhiza* sp., 213 *Opuntia* sp. *O. basilaris*, 167, 255, 328 *O. paraguayensis*, 350 *O. phaeacantha*, 257 *O. polyacantha*, 330 *Orbeanthus hardyi*, 153 Orchidaceae, 33, 57, 76, 81, 137, 147–152, 203, 292, 293, 324, 347, 352, 353, 416, 417, 434 *Orchidantha maxillarioides*, 246, 346 *Orchis* sp. *O. pallens*, 417 *O. ustulata*, 81 *Origanum vulgare*, 230 *Orlaya grandiflora*, 165, 430 *Ornithocephalus myrticola*, 150 *Ornithogalum* sp. *O. narbonense*, 325 *O. nutans*, 399 Orobanchaceae, 26, 29, 30, 32, 172, 185, 201, 204, 214, 225, 227, 228, 275, 276, 290, 332, 333, 341, 371, 382, 384, 387, 395, 397, 408, 412, 423, 425 *Orobanche* sp. *O. hederae*, 13, 26, 204, 384, 397, 425 *O. lutea*, 332 *Orthilia secunda*, 160 *Oryctanthus sp*., 72, 75 *Ouratea hexasperma*, 373 Oxalidaceae, 62 *Oxalis* sp., 62 *O. acetosella*, 182 *Oxytropis jacquinii*, 33, 173

#### **P**

*Pachira* sp. *P. aquatica*, 70, 73, 261, 357 *P. quinata*, 260 *P. sessilis*, 260 *Pachypodium* sp. *P. saundersii*, 432 *P. succulentum*, 393, 402 *Pachysandra terminalis*, 357, 383 *Pachystachys lutea*, 273, 300 Papaveraceae, 156, 162, 171, 184, 194, 209, 215, 225, 232, 255, 257, 290, 305, 375–377, 395 *Papaver rhoeas*, 215 *Paradisea liliastrum*, 283, 349 *Paramoltkia doerfleri*, 268 *Pardoglossum* sp., 242 *Parentucellia latifolia*, 371

*Parnassia palustris*, 171 *Paronychia polygonifolia*, 182 *Parthenocissus* sp., 240 Passifloraceae, 77, 252, 253, 276, *Passiflora* cf. *incarnata*, 73, 77 *P. amethystina*, 276, 320 *P. citrina*, 73, 77, 252 *P. suberosa*, 73, 77, 253 *Patrinia gibbosa*, 311 *Paullinia tomentosa*, 164 *Pavonia multiflora*, 271, 310 *P. elongata*, 225 *P. gyroflexa*, 172, 276 *P. palustris*, 275 *P. portenschlagii*, 185 *P. rostratocapitata*, 275 *P. verticillata*, 290, 341 *P. carnosum*, 367, 404 *P. punctatum*, 162, 368 *P. tetragonum*, 367 *P. polybotrya*, 297 fossil, Miocene, Austria, 354 *P. bistorta*, 260 *P. chinensis*, 320, 327 *Petasites albus*, 312 *Petrea volubilis*, 200, 221, 228 *Petrorhagia prolifera*, 332 *Peucedanum cervaria*, 174, 358 *Pfaffia tuberosa*, 328 *P. campanularia*, 241, 243 *P. tanacetifolia*, 242

320, 322, 352

*Passiflora* sp.

*Pedicularis* sp.

*Pelargonium* sp.

*Peperomia* sp.

*P. rubella*, 296 *Persicaria* sp.

*P. mitis*, 349

*Phacelia* sp.

*Phlox* sp.

*Phaleria capitata*, 258, 355 *Pherosphaera hookeriana*, 193 *Phillyrea angustifolia*, 370 *Phleum pratense*, 157

*P. drummondii*, 300 *P. paniculata*, 181, 300 *Phoebe sheareri*, 245 Phrymaceae, 277, 278 *Phyllanthus* sp., 306

*P. abies*, 191 *P. pungens*, 191 Picrodendraceae, 50

*Pinellia* sp., 15 *P. ternata*, 245, 309

*Pinguicula* sp. *P. alpina*, 353 *P. ehlersiae*, 161, 238

*Phyllanthus x elongatus*, 72, 74 *Physostegia virginiana*, 159, 348 *Phyteuma spicatum*, 266 *Picea* sp., 188, 190, 247

fossil, Miocene, China, 188, 190

199, 211, 247, 323, 385, 433

Pinaceae, 16, 50, 51, 68, 133, 179, 188, 191–193,

fossil, middle Miocene, Austria, 68, 189 fossil, Miocene, China, 188–190, 372

*Pinus* sp., 190, 385 fossil, middle Miocene, Austria, 52, 385 fossil, Miocene, China, 190 *P. cembra*, 188, 247 *P. contorta*, 192 *P. heldreichii*, 192 *P. mugo*, 191, 192, 433 *P. nigra*, 192 *P. strobus*, 133, 179, 211, 247 *Pinus* subgenus *Pinus*, 16, 50, 52, 445 *Pinus* subgenus *Strobus*, 16, 50, 52, 445 Piperaceae, 197, 198, 296, 297, 372 *Piper* sp. *P. auritum*, 198, 372 *P. nigrum*, 197 *Pistacia* sp., 221 *Pistia* sp., 10 *P. stratiotes*, 344, 396 *Pisum sativum*, 352 *Pithecellobium dulce*, 145, 316 Pittosporaceae, 353 *Plagiorhegma dubium*, 369 Plantaginaceae, 16, 17, 80, 178, 203, 213, 214, 217, 227, 254, 270, 308, 331, 332, 337, 340, 361, 374–376, 380, 381, 386, 389, 393–395, 401, 413, 414, 426 *Plantago* sp. *P. lanceolata*, 254, 376, 389 *P. major*, 270, 380, 426 *P. maritima*, 87, 374, 381, 393, 395 *P. media*, 375 *P. reniformis*, 376 *Platycapnos tenuilobus*, 376 *Platycodon grandiflorus*, 175, 230 *Plectranthus* sp. *P. esculentus*, 168, 279, 325 *P. ornatus*, 301 *Pleurothallis loranthophylla*, 151 Plumbaginaceae, 61, 226, 304, 326, 327, 357, 373 *Plumbago auriculata*, 304 Poaceae, 13, 14, 31, 54, 133, 157, 178, 203, 204, 209, 210, 212, 251, 254, 292, 296, 336, 337, 399, 421, 434 *Poa* sp., 14 *P. angustifolia*, 254 *P. pratensis*, 254 Podocarpaceae, 16, 50, 188, 192, 193 fossil, early Miocene, South-Africa, 193 *Podocarpus* sp., 192 *Podophyllum peltatum*, 333 Podostemaceae, 134, 224, 431 *Podostemum munnarense*, 431 *Poikilacanthus macranthus*, 296 Polemoniaceae, 181, 258, 269, 300, 322, 368, 373, 387 *Polemonium caeruleum*, 368, 373 *Polemonium pauciflorum*, 368 Polygalaceae, 238, 239, 279, 280, 352 *Polygala* sp. *P. chamaebuxus*, 238 *P. major*, 239, 352 *P. myrtifolia*, 279, 280 Polygonaceae, 10, 55, 234, 235, 260, 320, 327, 343, 349 fossil, Miocene, Austria, 354 fossil, Quaternary Austria, 241 *Polygonum aviculare*, 10 Polypodiaceae, 44 *Polypodium* sp., 44 *Polyscias filicifolia*, 335

*Poncirus trifoliata*, 237, 351 Pontederiaceae, 287 *Pontederia cordata*, 75 *Populus alba*, 204, 245, 360 Portulacaceae, 255, 311 *Portulaca grandiflora*, 255, 311 Posidoniaceae, 245 *Posidonia* sp., 245 *Potentilla* sp. *P. erecta*, 253 *P. incana*, 251 *P. inclinata*, 165, 361 *Prenanthes purpurea*, 329, 431 Primulaceae, 12, 57, 60, 68, 69, 164, 173, 175, 202, 230, 240, 241, 248, 250, 290, 291, 320, 342, 394, 434 *Primula* sp. *P. denticulata*, 68, 69, 164 *P. farinosa*, 68, 69, 434 *P. veris*, 57, 60, 230 *Proboscidea fragrans*, 305 Proteaceae, 42, 163, 169, 180, 200–202, 321, 359 *Prunella grandiflora*, 301 *Prunus* sp., 33 *P. avium*, 361 *P. laurocerasus*, 365 *Pseuderanthemum alatum*, 243 *Pseudofumaria lutea*, 232, 290, 305 *Pseudotsuga*, 68 *Ptelea trifoliolata*, 332 Pteridaceae, 44 fossil, middle Miocene, Austria, 44 *Pterocarya* sp., fossil, middle Miocene, Austria, 268 *Pulmonaria* sp. *P. angustifolia*, 177 *P. mollis*, 237 *P. officinalis*, 343 *Pulsatilla grandis*, 374

#### **Q**

*Quercus* sp. *Q. petrea*, 43 *Q. robur*, 29, 334, 374, 381, 400, 422, 426 *Quesnelia* sp. *Q. augusto*-*coburgii*, 179 *Q. imbricata*, 185 *Q. lateralis*, 263, 383

#### **R**

Ranunculaceae, 28, 161, 195, 215, 219, 227, 255, 335, 371, 374, 381, 392, 394, 395, 397, 403, 412, 423, 426 *Ranunculus* sp. *R. lanuginosus*, 255 *R. trichophyllus*, 395 Rapateaceae, 275 *Razisea citrina*, 348 Resedaceae, 335 *Reseda luteola*, 335 Rhamnaceae, 249 *Rhamnus cathartica*, 249 *Rhaphidophora africana*, 339, 384 *Rhododendron* sp., 95 fossil, Miocene, North-east China, 95 *R. hippophaeoides*, 406 *R. hirsutum*, 139, 315, 420 *Rhus* sp., 235 *Ribes aureum*, 257

*Roemeria hybrida*, 171, 194 Roridulaceae, 169, 218, 322 *Roridula gorgonias*, 169, 218, 322 Rosaceae, 26, 33, 62, 160, 165, 211, 223, 234, 238, 251, 253, 262, 279, 360–365, 392 fossil, Miocene, China, 338 *Rosa pendulina*, 253 Rubiaceae, 27, 32, 139, 157, 176, 194, 213, 229–231, 234, 236, 280, 336, 373, 411 *Rubus caesius*, 211, 362 *Ruellia* sp., 326 *R. brevifolia*, 350 *R. devosiana*, 320 *R. macrantha*, 182 *R. makoyana*, 319 *R. tuberosa*, 327 *Rumex* sp. fossil, Quaternary Austria, 241 *R. acetosa*, 234, 343 *Ruspolia seticalyx*, 422 Rutaceae, 237, 241, 298, 332, 351, 364, 404 *Ruta graveolens*, 364

#### **S**

*Sagittaria sagittifolia*, 272 Salicaceae, 204, 216, 245, 250, 349, 360, 383, 413 *Salix* sp. *S. alba*, 216 *S. daphnoides*, 349 *S. fragilis*, 383 *S. retusa*, 250 *Salix x fragilis*, 413 *Salvia* sp. *S. argentea*, 168, 299 *S. austriaca*, 55 *S. coccinea*, 159 *S. glutinosa*, 279, 299, 411, 413 *S. hians*, 299 *S. nemorosa*, 27, 380, 411 *S. sclarea*, 175 *S. verticillata*, 28, 380 *Sambucus nigra*, 414 *Sanchezia nobilis*, 51, 54, 263 *Sandersonia vaurantiaca*, 433 *Sanguisorba* sp. *S. cretica*, 262 *S. minor*, 262, 363 *S. officinalis*, 160, 238, 279, 360 *Sanicula europaea*, 161 *Sansevieria* sp. *S. parva*, 179, 292 *S. suffruticosa*, 186 Santalaceae, 70, 72, 172, 187, 202, 298 fossil, Miocene, China, 340 Sapindaceae, 68, 69, 160, 164, 173, 177, 184, 208, 215, 216, 219, 359, 361, 362, 365, 394 fossil, middle Miocene, Austria, 228 *Saponaria officinalis*, 53 *Sarcocapnos enneaphylla*, 171, 232, 377 *Sarcococca pruniformis*, 356 *Sarcopoterium spinosum*, 262 *Sarracenia alata*, 167, 169 Sarraceniaceae, 167, 169 *Saruma henryi*, 157 *Satyria warszewiczii*, 409 *Sauromatum venosum*, 384, 423, 431

Saururaceae, 285 *Saururus cernuus*, 285 Saxifragaceae, 48, 217, 362, 363, 365, 372 *Saxifraga* sp. *S. hostii*, 372 *S. rotundifolia*, 362 *S. scardica*, 48 *S. taygetea*, 363 *S. tridactylites*, 365 *S. vandellii*, 217 *Scabiosa* sp. *S. caucasica*, 46 *S. ochroleuca*, 221 *Scaevola* sp., 62 *S. aemula*, 241, 425 *Scandix pecten*-*veneris*, 62 *Schaueria flavicoma*, 261 *Schinus molle*, 369 *Schoenoplectus lacustris*, 135, 272 *Schoenorchis fragrans*, 151 Schoepfiaceae, 172, 184 *Schoepfia schreberi*, 172, 184 *Schotia brachypetala*, 320 *Scilla bifolia*, 413 *Scirpoides holoschoenus*, 136, 336 *Scirpus sylvaticus*, 135, 272 *Scorzonera* sp. *S. aristata*, 330 *S. austriaca*, 314 Scrophulariaceae, 26, 214, 394 *Scrophularia* sp. *S. nodosa*, 26 *S. vernalis*, 214 *Scutellaria baicalensis*, 17 *Secale cereale*, 209 *Sechium edule*, 231 *Securigera varia*, 358 *Sedum* sp. *S. acre*, 358 *S. rupestre*, 161 *Sempervivum globiferum*, 165, 346 *Sesleria* sp., 14 *S. albicans*, 178, 204 *S. caerulea*, 434 *S. sadleriana*, 212 *Sherardia arvensis*, 231 *Sideritis* sp. *S. montana*, 55 *S. romana*, 166, 229 *S. syriaca*, 255 *Silene* sp. *S. flos*-*cuculi*, 432 *S. latifolia*, 181 *S. nutans*, 171 *S. succulenta*, 53 Simaroubaceae, 367, 382, 394 *Sinapis alba*, 415 Smilacaceae, 203, 334 *Smilax spinosa*, 203, 334 *Smyrnium perfoliatum*, 32, 34, 403 Solanaceae, 55, 162, 184, 186, 237, 249, 333, 359, 362, 368, 369 *Solandra* sp. *S. longiflora*, 55, 368 *S. maxima*, 369 *Solanum torvum*, 333 *Solidago canadensis*, 386

*Sparganium erectum*, 73, 78, 197, 293 *Sparmannia africana*, 194, 218, 301 *Spathicarpa* sp., 384 *S. sagittifolia*, 244 *Spathiphyllum* sp., 244 *S. blandum*, 396 *S. cannifolium*, 345 *S. minor*, 345 Sphagnaceae, fossil, middle Miocene, Austria, 44 *Sphagnum* sp., fossil, middle Miocene, Austria, 44 *Spinizonocolpites* sp., fossil, 13 *Spiranthes spiralis*, 150, 416 *Spirea* sp., 26 *Spirogyra* sp., 49 *Stachys* sp. *S. officinalis*, 395 *S. palustris*, 28, 30, 226 *Stanfieldiella imperforata*, 377 *Stanhopea oculata*, 151 *Stellaria* sp. *S. graminea*, 258, 421 *S. holostea*, 167, 269 *S. media*, 53 *Stenandrium* sp. *S. dulce*, 321 *S. guineense*, 175 *Stephanotis floribunda*, 149, 153 *Steveniella satyrioides*, 148, 152 *Stigmaphyllon lindenianum*, 256 *Stratiotes aloides*, 314 *Streptocalyx poeppigii*, 317 *Strobilanthes roseus*, 326 *Stylochaeton bogneri*, 244 *Succisa pratensis*, 221 *Symphytum* sp. *S. caucasicum*, 239, 337 *S. orientale*, 222 Symplocaceae, fossil, middle Miocene, Austria, 208 *Symplocarpus foetidus*, 197 *Symplocos* sp., fossil, middle Miocene, Austria, 208 *Synandrospadix vermitoxicus*, 244 *Syringa vulgaris*, 398, 413

#### **T**

*Tabernaemontana simulans*, 222 *Taccarum weddellianum*, 46 *Takhtajania perrieri*, 89–92 Talinaceae, 167, 232 *Talinum paniculatum*, 167, 232 *Tanacetum corymbosum*, 312, 388, 391 *Taraxacum* sp., 329 fossil, Quaternary, Austria, 329 *Tasmannia insipida*, 89–92 Taxaceae, 50, 53 Tecophilaeaceae, 376 *Tetragonia tetragonioides*, 341 *Tetramerium nervosum*, 242 *Tetrapollinia caerulescens*, 265, 314 *Teucrium* sp. *T. chamaedrys*, 376 *T. pyrenaicum*, 251 *Thalictrum flavum*, 397 Theaceae, 252 *Thelethylax minutiflora*, 134, 224

*Theobroma cacao*, 350 *Thesium* sp. *T. alpinum*, 70, 72 *T. arvense*, 172, 202 *T. dollineri*, 187 *Thladiantha hookeri*, 319, 353 *Thlaspi montanum*, 326 *Thunbergia* sp. *T. alata*, 277, 433 *T. laurifolia*, 278 Thymelaeaceae, 258, 307, 308, 355 *Thymelaea passerina*, 307, 308 *Thymus* sp. *T. glabrescens*, 27 *T. odoratissimus*, 31, 83 Tiliaceae, 194 *Tilia* sp., 211 fossil, Miocene, China, 333, 338 *T. euchlora*, 199 *T. platyphyllos*, 220, 222, 260, 423 *Tofieldia calyculata*, 75, 287 Tofieldiaceae, 287 *Torilis arvensis*, 174 *Tradescantia* sp. *T. spathacea*, 334 *T. zebrina*, 286 *Tragopogon* sp. *T. dubius*, 214, 329 *T. orientalis*, 132, 133, 329 Trapaceae fossil, late Miocene, Austria, 76 fossil, Miocene, China, 339, 341 *Trapa* sp., fossil, Miocene, China, 339, 341 *Trichosanthes* sp. *T. anguina*, 78, 82 *T. cucumerina*, 210 *Tricolporopollenites wackersdorfensis*, fossil, middle Miocene, Austria, 235 *Tridax procumbens*, 237 *Trifolium* sp. *T. montanum*, 343 *T. rubens*, 349 *Triglochin maritima*, 246, 370 Trigoniaceae, 347 *Trigonia nivea*, 347 *Trillium* sp. *T. chloropetalum*, 246, 332 *T. grandiflorum*, 331 *Trisetum flavescens*, 399, 421 *Tristellateia australasiae*, 256 *Triticum aestivum*, 31, 209, 254 *Trollius europaeus*, 227 Tropaeolaceae, 163, 202, 208, 351 *Tropaeolum* sp. *T. emarginatum*, 163 *T. majus*, 208, 351 *T. moritzianum*, 202 *Trudopollis* sp., fossil, Upper Cretaceous, Hungary, 11 *Tsuga* sp. fossil, middle Miocene, Austria, 189, 385 fossil, Miocene, Austria, 189 *T. canadensis*, 188, 189, 199 *Tsusiophyllum tanakae*, 202 *Tuberaria guttata*, 33

*Tulipa* sp. *T. kaufmanniana*, 71, 410 *T. linifolia*, 410 *T. sylvestris*, 252 *T. turkestanica*, 371 *Turbinicarpus pseudomacrochele*, 232 *Turnera ulmifolia*, 322 *Tussilago farfara*, 240 Typhaceae, 41, 143, 197, 224, 293, 434 *Typha* sp., 10 *T. latifolia*, 41, 143, 224 *T. laxmannii*, 434

#### **U**

*Ulearum sagittatum*, 313 Ulmaceae, 31, 56, 219, 268, 389, 426 fossil, Miocene, Austria, 358 fossil, Miocene, China, 323, 372 *Ulmus* sp., 372 fossil, Miocene, China, 372 *U. laevis*, 56, 389, 426 *U. minor*, 31, 219, 268 Urticaceae, 204, 267, 426 *Urtica dioica*, 204, 426 *Utricularia vulgaris*, 239 *Uvularia grandiflora*, 75, 287

#### **V**

*Vanilla* sp. *V. planifolia*, 203 *V. pompona*, 33 Velloziaceae, 287 *Veratrum* sp. *V. album*, 194, 286, 351 *V. nigrum*, 262 Verbenaceae, 200, 202, 221, 228 *Verbena officinalis*, 202 *Veronica* sp. *V. anagallis-aquatica*, 401 *V. cinerea*, 361 *V. longifolia*, 337 *V. prostrata*, 332 *V. serpyllifolia*, 227 *V. spicata*, 213, 414 *V. wyomingensis*, 217 *Viburnum* sp. *V. lantana*, 304 *V. opulus*, 319 *V. tinus*, 177 *V. utile*, 371 *Vicia faba*, 241 *Victoria regia*, 139, 276, 405

*Vigna speciosa*, 218, 352 *Vinca minor*, 346 Violaceae, 161, 162, 166, 167, 238, 338, 389, 408, 409, 430 *Viola* sp. *V. alba*, 161, 430 *V. arvensis*, 167, 238 *V. calcarata*, 338, 409 *V. riviniana*, 162 *V. tricolor*, 166, 389, 408 *Viscum album*, 298 Vitaceae, 240 *Vitaliana primuliflora*, 175, 241 *Vitex trifolia*, 300 *Vriesea* sp. *V. neoglutinosa*, 170, 283 *V. pabstii*, 58

#### **W**

*Wachendorfia thyrsiflora*, 198, 286, 322 Welwitschiaceae, 345, 432 *Welwitschia mirabilis*, 345, 432 *Werauhia tarmaensis*, 351 *Whitfieldia* sp. *W. elongata*, 271 *W. lateralis*, 263 *W. lateritia*, 172, 185, 347 Winteraceae, 86, 89–92, 139, 293

#### **X**

*Xanthium* sp. *X. saccharatum*, 404 *X. spinosum*, 49 *Xanthosoma ceronii*, 137 Xanthorrhoeaceae, 133, 166, 187, 196, 198, 282, 285, 288, 337 *Xerophyta elegans*, 287 Xyridaceae, 302

#### **Z**

Zamiaceae, 159, 197, 285, 360 *Zamia loddigesii*, 159 *Zamioculcas zamiifolia*, 274, 385 *Zantedeschia* sp. *Z. aethiopica*, 26 *Z. aetiopica*, 32 *Zea mays*, 31, 210, 251 *Zelkova* sp.,fossil, Miocene, China, 358 *Zeylanidium subulatum*, 134, 431 Zingiberaceae, 257, 346, 384, 431 *Zomicarpa riedeliana*, 313 Zygophyllaceae, 270, 327