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Special Issue Reprint

Plant Cell Wall Proteins
and Development

Edited by
Elisabeth Jamet and Christophe Dunand

www.mdpi.com/journal/ijms
Plant Cell Wall Proteins and
Development
<figure><img src="image_2.png" /></figure>
Plant Cell Wall Proteins and
Development

Editors

Elisabeth Jamet

Christophe Dunand

MDPI • Basel • Beijing • Wuhan • Barcelona • Belgrade • Manchester • Tokyo • Cluj • Tianjin

<figure><img src="image_3.png" /></figure>
Editors

Elisabeth Jamet

CNRS (Centre National de la
recherche Scientifique)

France

Christophe Dunand

Université de Toulouse

France

Editorial Office

MDPI

St. Alban-Anlage 66

4052 Basel, Switzerland

This is a reprint of articles from the Special Issue published online in the open access journal *International Journal of Molecular Sciences* (ISSN 1422-0067) (available at: https://www.mdpi.com/journal/ijms/special_issues/plant_cell wall).

For citation purposes, cite each article independently as indicated on the article page online and as indicated below:

Last Name, A.A.; Last Name, B.B.; Last Name, C.C. Article Title. *Journal Name Year, Volume Number, Page Range*.

ISBN 978-3-0365-7590-2 (Hbk)

ISBN 978-3-0365-7591-9 (PDF)

© 2023 by the authors. Articles in this book are Open Access and distributed under the Creative Commons Attribution (CC BY) license, which allows users to download, copy and build upon published articles, as long as the author and publisher are properly credited, which ensures maximum dissemination and a wider impact of our publications.

The book as a whole is distributed by MDPI under the terms and conditions of the Creative Commons license CC BY-NC-ND.
# Contents

<table><tr><td>About the Editors</td><td>vii</td></tr><tr><td>Preface to "Plant Cell Wall Proteins and Development"</td><td>ix</td></tr><tr><td>Elisabeth Jamet and Christophe Dunand<br>Plant Cell Wall Proteins and Development<br>Reprinted from: Int. J. Mol. Sci. 2020, 21, 2731, doi:10.3390/ijms21082731</td><td>1</td></tr><tr><td>Maria Juliana Calderan-Rodrigues, Juliana Guimarães Fonseca, Fabrício Edgar de Moraes, Laís Vaz Setem, Amanda Carmanhanis Begossi and Carlos Alberto Labate<br>Plant Cell Wall Proteomics: A Focus on Monocot Species, Brachypodium distachyon, Saccharum spp. and Oryza sativa<br>Reprinted from: Int. J. Mol. Sci. 2019, 20, 1975, doi:10.3390/ijms20081975</td><td>7</td></tr><tr><td>Mehdi Cherkaoui, Virginie Lollier, Audrey Geairon, Axelle Bouder, Colette Larré, Hélène Rogniaux, et al.<br>Cell Wall Proteome of Wheat Grain Endosperm and Outer Layers at Two Key Stages of Early Development<br>Reprinted from: Int. J. Mol. Sci. 2020, 21, 239, doi:10.3390/ijms21010239</td><td>27</td></tr><tr><td>Alexander Betekhtin, Anna Milewska-Hendel, Joanna Lusinska, Lukasz Chajec, Ewa Kurczynska and Robert Hasterok<br>Organ and Tissue-Specific Localisation of Selected Cell Wall Epitopes in the Zygotic Embryo of Brachypodium distachyon<br>Reprinted from: Int. J. Mol. Sci. 2018, 19, 725, doi:10.3390/ijms19030725</td><td>45</td></tr><tr><td>Xiao Han, Li-Jun Huang, Dan Feng, Wenhan Jiang, Wenzhuo Miu and Ning Li<br>Plasmodesmata-Related Structural and Functional Proteins: The Long Sought-After Secrets of a Cytoplasmic Channel in Plant Cell Walls<br>Reprinted from: Int. J. Mol. Sci. 2019, 20, 2946, doi:10.3390/ijms20122946</td><td>61</td></tr><tr><td>Zdeňka Kubátová, Přemysl Pejchar, Martin Potocký, Juraj Sekereš, Viktor Žárský and Ivan Kulich<br>Arabidopsis Trichome Contains Two Plasma Membrane Domains with Different Lipid Compositions Which Attract Distinct EXO70 Subunits<br>Reprinted from: Int. J. Mol. Sci. 2019, 20, 3803, doi:10.3390/ijms20153803</td><td>77</td></tr><tr><td>Artur Pinski, Alexander Betekhtin, Katarzyna Sala, Kamila Godel-Jedrychowska, Ewa Kurczynska and Robert Hasterok<br>Hydroxyproline-Rich Glycoproteins as Markers of Temperature Stress in the Leaves of Brachypodium distachyon<br>Reprinted from: Int. J. Mol. Sci. 2019, 20, 2571, doi:10.3390/ijms20102571</td><td>91</td></tr><tr><td>Li Song, Babu Valliyodan, Silvas Prince, Jinrong Wan and Henry T. Nguyen<br>Characterization of the XTH Gene Family: New Insight to the Roles in Soybean Flooding Tolerance<br>Reprinted from: Int. J. Mol. Sci. 2018, 19, 2705, doi:10.3390/ijms19092705</td><td>113</td></tr><tr><td>Weiwei Wu, Shengnan Zhu, Qianqian Chen, Yan Lin, Jiang Tian and Cuiyue Liang<br>Cell Wall Proteins Play Critical Roles in Plant Adaptation to Phosphorus Deficiency<br>Reprinted from: Int. J. Mol. Sci. 2019, 20, 5259, doi:10.3390/ijms20215259</td><td>133</td></tr></table>
# About the Editors

## Elisabeth Jamet

Elisabeth Jamet gained a Ph.D. in 1985 at Strasbourg University (France) on so-called plant-pathogenesis-related proteins. She was the first to be able to purify these proteins which were considered at that time as degradation products due to their great stability. She was hired as a permanent researcher at the National Center for Scientific Research (CNRS) in 1986 at the Institute of Plant Molecular Biology (IBMP) in Strasbourg. She then developed a research project dealing with plant cell dedifferentiation through protoplasts. She focused on the role of the cell wall in protoplast recovery, a critical step to acquire the capability to divide, form a callus, and regenerate a plant. In 2001, as a research director at CNRS, she moved to the Plant Science Research Laboratory (LRSV) close to Toulouse (France) where she developed her own team focusing on the role of cell wall proteins in plant development. She has greatly contributed to the establishment of diverse strategies to characterize the plant cell wall proteome, including experimental approaches and bioinformatics treatment of mass spectrometry data. In parallel, she has contributed to the unraveling of the function of several families of cell wall proteins, notably the PAC (proline-rich, arabinogalactan proteins, conserved cysteines) domain proteins. She is presently the co-leader of the Dynamics and Evolution of Plant Cell Walls team at LRSV. She has co-authored 100 articles in international journals, and she is an editor of *Frontiers in Plant Science*, the *International Journal of Molecular Science*, and *Cells*. She has been/is a member of different scientific evaluation committees at national and international levels. She was the head of LRSV between 2011 and 2015, and she presently exercises responsibilities at Toulouse University.

## Christophe Dunand

Christophe Dunand gained a Ph.D. in 1997 at Grenoble University (France) in plant biochemistry. He has demonstrated the binding and biological effects of oligosaccharides on the enzymatic activity of plant protoplast extracts. After gaining his Ph.D., he completed three post-doctoral internships. First, he completed 1 year at the University J. Fourier (Grenoble) on Arabidopsis floral development and then 18 months at the University of Connecticut on the study of cell walls. In 2000, he joined the Plant Physiology Laboratory of Geneva University as an assistant professor. Until 2008, he strongly developed bioinformatics and molecular biology approaches dedicated to plant peroxidase genes. He was hired as a full professor at the University of Toulouse in 2008 at the Plant Science Research Laboratory (LRSV) close to Toulouse (France). He then developed his own team to continue studies on plant peroxidase functions and evolution. His research interests mainly focus on the model plant Arabidopsis, and his research has taken a more integrative direction with the use of less typical models and approaches such as hepatic or aquatic plants. He is presently the co-leader of the Dynamics and Evolution of Plant Cell Walls team at LRSV. He has co-authored 90 articles in international journals, and he is an editor of *Frontiers in Plant Science*, the *International Journal of Molecular Science*, and *Cells*.
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Preface to "Plant Cell Wall Proteins and
Development"

This book focuses on plant cell wall proteins and their roles during development and in response to abiotic stresses. Plant cell walls are mainly composed of polymers, such as polysaccharides or lignins. Although proteins are minor components, they play crucial roles in signaling and in the dynamics of the cell wall architecture, enabling growth, differentiation, and the adaptation of plants to their changing environment. This collection of research reports and reviews published in a Special Issue of *IJMS* includes a broad range of topics and experimental approaches which illustrate recent advances in cell wall biology.

Elisabeth Jamet and Christophe Dunand

Editors
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Editorial

# Plant Cell Wall Proteins and Development

Elisabeth Jamet * and Christophe Dunand *

Laboratoire de Recherche en Sciences Végétales, Université de Toulouse, CNRS, UPS,
31320 Auzeville Tolosane, France

* Correspondence: jamet@lrsv.ups-tlse.fr (E.J.); dunand@lrsv.ups-tlse.fr (C.D.)

Received: 9 April 2020; Accepted: 13 April 2020; Published: 15 April 2020

Plant cell walls surround cells and provide both external protection and a means of cell-to-cell communication. They mainly comprise polymers like polysaccharides (cellulose, hemicelluloses, and pectins) and lignin in lignified secondary walls and a small amount of cell wall proteins (CWPs) [1,2]. CWPs are major players of cell wall remodeling and signaling. Cell wall proteomics, as well as numerous genetic or biochemical studies, have revealed the high diversity of CWPs, among which proteins acting on polysaccharides, proteases, oxido-reductases, lipid-related proteins, and structural proteins ([3–7]). CWPs may have enzymatic activities such as cutting/ligating polymers or processing/degrading proteins [8]. They may also contribute to the supra-molecular assembly of cell walls via protein/protein or protein/polysaccharide interactions [9–11]. Thanks to these biochemical activities, they contribute to the dynamics and functionality of cell walls. Even though much research has already been pursued to shed light on the many roles of CWPs, many functions still remain to be discovered, especially for proteins identified in cell wall proteomes with yet unknown function.

This Special Issue "Plant Cell Wall Proteins and Development" has welcomed a selection of articles in the field of cell wall biology, which were focused on cell wall proteins and their roles during development. Eight experimental articles, nine up-to-date review articles, as well as a concept article, have been published. We wish to thank all the authors for their great contribution to this unique collection of articles as well as the International Journal of Molecular Science supporting team.

The content of this Special Issue embraces several topics, all of them stressing the roles of cell wall proteins: cell wall proteomics studies on monocot species [7,12]; the role of cell wall proteins during plant development [13–15] or in response to environmental stresses [16–19]; overviews on several cell wall protein families either from green microalgae [20] or from plants, i.e., fasciclin arabinogalactan proteins (FLAs) [21,22], membrane-bound class III peroxidases (Class III Prxs) [23], pectin methylesterases inhibitors [24], DUF642 (Domain of Unknown Function 642) proteins [25], and Proline-rich, Arabinogalactan proteins, conserved Cysteines (PAC) domain-proteins [26]; and the role of fasciclin arabinogalactan proteins (FLAs) in Ca²⁺ signaling during plant morphogenesis [27,28].

For two decades, cell wall proteomics has become a powerful experimental approach and has revealed the diversity of the cell wall protein families. *Arabidopsis thaliana* has been the most studied plant species, and almost half of its expected cell wall proteome has been described so far (see WallProtDB, http://www.polebio.lrsv.ups-tlse.fr/WallProtDB/). The monocotyledon species have been studied more recently thanks to the sequencing of additional genomes like those of *Oryza sativa* [29], *Brachypodium distachyon* [30], and *Triticum aestivum* [31] as well as the availability of transcriptomics data as for *Saccharum* spp [32]. Calderan-Rodrigues et al. [7] provide a comparison of monocotyledon and dicotyledon cell wall proteomes and have discussed the specificities of the former. Such specificities were related to the differences between the composition and structure of monocotyledon and dicotyledon cell walls [1,33]. Also, Cherkaoui et al. [12] report on the comparison between cell wall proteomes of the endosperm, and the outer layers of the wheat grain. They reveal a strong metabolic activity in the cell wall during endosperm differentiation, whereas the accumulation of proteins was more important at an earlier stage of development in the outer layers.
As mentioned above, the cell wall composition and structure varies during development, and these changes can allow further differentiation processes. Betekhtin et al. [13] provide a fine mapping of cell wall epitopes in zygotic embryos of *B. distachyon* at a mature stage of development, including antibodies recognizing extensins and arabinogalactan proteins (AGPs), which are structural proteins involved in the cell wall architecture and proteins assumed to be involved in signaling, respectively. The plasma membrane is the interface between the cytoplasm and the cell wall. Its composition can vary locally in the domains characterized by particular lipid compositions. Kubátová et al. [15] show that two plasma membrane domains with a distinct lipid composition are located close to the Ortmannian ring, a cell wall domain-specific to trichomes. These plasma membrane domains are generated thanks to exocysts complex containing EXO70 subunits recognizing the target membrane. Cell-to-cell communication can be ensured through plasmodesmata [34]. Han et al. [14] provide a review article focusing on the cytoskeleton and on plasmodesmata-associated cell wall proteins like callose synthase and callose hydrolase, which are involved in the regulation of plasmodesmata closure.

Environmental cues induce modifications of the cell wall. In particular, nutrient availability can regulate cell wall composition. The absorption of nutrients by roots occurs through the apoplastic pathway. This pathway is blocked by the deposition of lignin and later of suberin at the level of the Casparian strips around endodermis cells in differentiated roots. In their review article, Ogden et al. [19] focus on the changes observed in the modulation of the suberization of the root endodermal walls in response to nutrient availability, showing that the plasticity of suberin accumulation is an adaptive response. They also show that the availability of nitrate or phosphorus modulates the development of lateral roots and/or of root hairs and has a direct effect on the transcription of genes encoding proteins involved in the biosynthesis of cell wall components or regulating the oxidative status in the cell wall. Wu et al. [18] focus on a few cell wall proteins playing critical roles during phosphorus deficiency such as expansins, Pro-rich proteins, oxidoreductases, and purple acid phosphatases. Abiotic stresses like flooding or temperature can also induce changes in the cell wall. Song et al. [17] show that xyloglucan endotransglycosylases/hydrolases (XTHs), which are hemicelluloses remodeling enzymes in muro, play roles in the regulation of stress responses to flooding. Indeed, the overexpression of the *A. thaliana* *AtXTH31* gene in soybean plants leads to increasing of resistance to flooding. Pinski et al. [16] observe changes in the accumulation of extensin and AGP epitopes in *B. distachyon* leaves exposed to cold and hot temperature stresses.

Cell wall proteins are mostly encoded by multigene families, which can comprise a large number of members like class III Prxs [35] or pectin methyl esterase inhibitors [36] (73 and 71 members in *A. thaliana*, respectively). Each member has its own regulatory pathway during development or upon stress, and even if the proteins of a give family share the same functional domains, subtle differences can confer different biological activities. As an example, AtPrx36 plays a particular role in mucilage release because of the timely regulation of expression of its gene during seed development, and of its anchoring in a cell wall microdomain [37]. Most cell wall protein families are conserved in the green lineage. This is illustrated in four articles of this Special Issue. Guerriero et al. [20] describe a family of green microalgal cellulases. Seifert et al. [21] show the conservation of the fasciclin 1 domain (FAS1) in all the kingdoms of life, suggesting a role in the mechanisms mediating interactions between the cells and their environment. He et al. [14] describe the evolution of FLAs which are possibly involved in signaling. Nguyen-Kim et al. [26] explore the PAC domain-proteins family possibly forming non-covalent networks with polysaccharides and O-glycoproteins.

Since cell wall proteins families contain many members, it is interesting to consider each of them independently to fully uncover their roles in cell wall biology. Three review articles present such overviews. Lüthje and Martinez-Cortes [23] describe the sub-family of membrane-bound class III Prxs which are located at the plasma membrane or in the tonoplast and are assumed to play roles in membrane protection or repair. Wormit and Usadel [24] give an overview of the roles of pectin methylesterase inhibitors (PMEIs). These proteins participate in the regulation of the degree of methylesterification of the pectic homogalacturonans, which in turn contributes to cell adhesion,
cell wall porosity, and plasticity. Finally, Cruz-Valderrama et al. [25] propose a role for the DUF642 protein family in development and in response to environmental stresses by modulating directly, or indirectly, the degree of methylation of homogalacturonans. These proteins were first described as abundant proteins in cell wall proteomes [38] and were until recently considered proteins with unknown function.

This Special issue was also open to new concepts. Two articles by Lamport et al. [27,28] propose new roles for the arabinogalactan protein (AGP) family in root and shoot morphogenesis, as well as in phyllotaxis patterning. Such molecules are actually proteoglycans with a proportion of glycans of up to 90% [39], which are assumed to play roles in signaling. However, the molecular mechanisms underlying this function were not deciphered until recently when its role as an extracellular calcium capacitor was proposed [40].

Altogether, we believe that this Special Issue will provide a collection of articles allowing both experts and newcomers in the field to get a valuable update on plant cell wall biology. A combination of research articles, reviews, and concept articles allows a survey of several topics of interest today regarding the many roles of cell wall proteins.

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© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).
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Review

# Plant Cell Wall Proteomics: A Focus on Monocot Species, *Brachypodium distachyon*, *Saccharum* spp. and *Oryza* sativa

Maria Juliana Calderan-Rodrigues *, Juliana Guimarães Fonseca, Fabrício Edgar de Moraes, Laís Vaz Setem, Amanda Carmanhanis Begossi and Carlos Alberto Labate

Department of Genetics, Max Feffer Laboratory of Plant Genetics, "Luiz de Queiroz" College of Agriculture, University of São Paulo, CP 83, 13400-970 Piracicaba, SP, Brazil; j.g.fonseca@usp.br (J.G.F.); fabricioedgar.m@gmail.com (F.E.d.M.); lala_arnz@hotmail.com (L.V.S.); amanda.lpp@hotmail.com (A.C.B.); calabate@usp.br (C.A.L.)

* Correspondence: mariajuliana@usp.br; Tel.: +55-019-3429-43-17

Received: 24 January 2019; Accepted: 7 March 2019; Published: 23 April 2019

**Abstract:** Plant cell walls mostly comprise polysaccharides and proteins. The composition of monocots' primary cell walls differs from that of dicots walls with respect to the type of hemicelluloses, the reduction of pectin abundance and the presence of aromatic molecules. Cell wall proteins (CWPs) differ among plant species, and their distribution within functional classes varies according to cell types, organs, developmental stages and/or environmental conditions. In this review, we go deeper into the findings of cell wall proteomics in monocot species and make a comparative analysis of the CWPs identified, considering their predicted functions, the organs analyzed, the plant developmental stage and their possible use as targets for biofuel production. *Arabidopsis thaliana* CWPs were considered as a reference to allow comparisons among different monocots, i.e., *Brachypodium distachyon*, *Saccharum* spp. and *Oryza sativa*. Altogether, 1159 CWPs have been acknowledged, and specificities and similarities are discussed. In particular, a search for *A. thaliana* homologs of CWPs identified so far in monocots allows the definition of monocot CWPs characteristics. Finally, the analysis of monocot CWPs appears to be a powerful tool for identifying candidate proteins of interest for tailoring cell walls to increase biomass yield of transformation for second-generation biofuels production.

**Keywords:** plant cell wall; proteome; monocot; stiff brome; rice; sugarcane; *Brachypodium distachyon*; *Saccharum* spp.; *Oryza sativa*

## 1. Introduction

The plant cell wall confines the cell volume and serves as protection against stresses, being responsible for the plant shape, enabling trees to be several meters high. In addition to these functions, the cell wall is the most external part of the cell, and as such, interacts with the apoplast, which is also essential in virtually all cell processes, including division, expansion, differentiation [1], growth and signaling [2]. This versatility implies that the mechanisms involved in a great deal of the cell wall functions have not been completely depicted.

The plant cell wall is a dynamic structure that undergoes changes during development [3]. The cell wall is mainly composed of wall polysaccharides, such as cellulose, hemicellulose, pectin, and proteins [4]. In some cell types, lignin and other compounds might be found, as well. Carbohydrates account for around 90% of the cell wall mass, and proteins around 10% for dicots [5] and 1% for monocots [6]. Cell wall proteins (CWPs) are understood to be proteins directed towards the secretory pathway, such as structural proteins linked to the wall and those secreted into the apoplast and extracellularly [7].
Some characteristics are commonly found in classical CWPs: an N-terminal cleavage signal-peptide, responsible for guiding these proteins to the endoplasmic reticulum (ER) [8]; lack of the ER-retention C-terminal motif KDEL/HDEL, which avoids the secretion of proteins [9,10]; and the absence of a transmembrane domain that retains proteins in the plasma membrane [11]. The secretory pathway directs proteins from ER to the Golgi apparatus, where they are packed into vesicles and guided to the cell wall and the extracellular matrix.

CWPs encompass hundreds of molecules presenting distinctive roles and are related to the modification of the cell wall components, signaling, interaction with the apoplast and the plasma membrane, and with virtually all the cell wall-related processes [12,13]. CWPs are usually divided into categories with respect to their functionality, such as proteins acting on carbohydrates, oxido-reductases, proteins related to lipid metabolism, and proteases, which represent the most abundant classes [7].

Besides being divided regarding their functional roles, CWPs can also be classified according to their type of link to the cell wall, such as structural proteins, which are strongly bound to the wall components, and loosely and weakly wall-bound proteins [13]. Loosely bound proteins are not linked to the cell wall polysaccharides and are able to move unconnected in the intercellular space. Weakly bound proteins interact with the matrix by Van der Waals, hydrogen bonds or ionic forces, and are related to remodeling, signaling, interactions with the plasma membrane and defense. Strongly bound proteins can be connected by covalent bonds [14].

To increase the comprehension of cell wall-related processes, several reports have characterized plant CWPs. To accomplish this task, specific methods of CWP extraction and data analysis had to be established, since these proteins are usually lost or in a low abundance in total proteins extraction surveys [15]. The constitution and structure of the CWPs collection are diverse due to the developmental stage, organ, tissue and species of the analyzed material; thus, it is possible to make a comparison among these different treatments and relate them physiologically [7]. Initial studies have been made on *Arabidopsis thaliana* [11,16–22]. Since then, CWPs from several other species have been identified by mass spectrometry. In *A. thaliana*, about one half of the predicted existing CWPs were identified, being the most studied plant at present, with 935 proteins already identified [11,16–22]. In addition to providing an overview of proteins present in a biological context, the identification of CWPs in *A. thaliana* enabled the selection of candidates to have their structure and function further investigated.

Over the course of time, several other studies have been performed with model plants and crops: Brassica [23,24], Zea mays [25], Cicer arietinum [26], Vitis vinifera [27], Oriza sativa (rice) [28–31], Nicotiana tabacum [32], Brachypodium distachyon (stiff brome) [33–36], Saccharum spp. (sugarcane) [37–40], among others. Due to this functional diversity, CWPs data present differences between monocots and dicots related to the cell wall structure and composition [41], probably due to both metabolisms' specificities. As other C3-dicots, *A. thaliana* has type I walls [42], whereas grasses, such as *Saccharum* spp., *B. distachyon*, *Sorghum bicolor* and *Setaria viridis*, present C4 metabolism and type II cell walls. *O. sativa* is a C3-monocot that also possesses type II walls [42]. Characteristic differences can be predicted when comparing CWPs collection from dicots and monocots, as their cell wall structures and components are diverse. For instance, the cell wall proteome of grasses generally consists of a higher proportion of oxido-reductases, and it has been suggested that this is due to the presence of aromatic compounds in the type II primary cell walls [33]. In summary, type II walls are characterized by a low proportion of pectins and xyloglucans, and high content of glucuronoarabinoxylans and mixed linked β-D-glucan, resulting in absent woody stems and branches [42,43], in comparison to dicots. Type II cell walls also contain ferulic and coumaric acids and more complex arabinoxylans in secondary walls. Moreover, the arabinosyl side chains inside arabinoxylans may be cross-linked with lignin through feruloyl esters [6,42]. One hypothesis for the selective advantage for the appearance of type II wall plants is that they are not as high and as reinforced as, for example, tall trees, so they focus their energy on rapid growth and more efficient reproductive strategies directed to occupy habitats that are not very suitable for trees [43].
Many of the monocots have C4 metabolism, presenting a higher photosynthetic efficiency, making them commercially valuable plants. As an example, *Saccharum* spp. has been used from more than three decades ago up to the present day as a raw material for the production of food, energy and co-products in Brazil [44]. Moreover, the study of CWPs in crops presents extra challenges compared to model plants, as less genetic and molecular data is usually available.

There is a need for more information about cell wall structure, components and their roles. The identification of CWPs allows new targets for further investigation and elucidation of pathways in order to better understand cell wall functions. Knowing that more than 10% of the plant's genome is related to the cell wall biogenesis [45], and as there are fewer studies on CWPs in monocots than dicots, it is important to gather and analyze CWP data from monocots, which would provide new insights about their unique metabolism and the specificities of their cell walls.

In this review, we compiled the information on monocots cell wall proteomics till date, except for studies using special treatments or stress-related data. Thus, *B. distachyon*, *Saccharum* spp. and *O. sativa* were chosen for this review. For each species, we searched for the *A. thaliana* sequences with the highest identities (BLASTp) in order to enable a comparison among all of them. Similar and identical *A. thaliana* sequences were found in monocots, and similarity in functional classes could also be established, as well as some specificities for each species. In this review, when it is mentioned that one *A. thaliana* CWPs was identified in the repertoire of the monocots, it means that the monocot CWP sequence matched that *A. thaliana* protein after BLASTp analysis.

## 2. Methods of Monocots CWPs Extraction and Analysis

Different methods of extraction of CWPs have been developed over the years. One of the biggest challenges is to isolate the cell wall with minimum contamination by intracellular or membrane proteins. Thus, the subcellular fractionation before proteomic analysis can be a useful strategy for acquiring a representative extract of CWPs and reducing contamination by proteins from other organelles. Protocols involving tissue grinding and centrifugation to generate a density gradient enable the separation of fractions that are highly enriched with specific cell compartments. However, such methods will also lead to plasma membrane disruption, which may result in subsequent contamination of cell wall subfraction. On the other hand, methodologies that do not break the cell structure can be less efficient, as the extraction buffers need to come into contact with the CWPs in order to extract them [17]. Thereby, methods of CWPs extraction can be divided into destructive and non-destructive techniques.

Regarding CWP extraction, the situation is not just one-method-fits-all, but rather the species, the organ and the targeted subset of proteins have to be considered in order to choose the most adequate protocol(s). The destructive techniques used for monocot CWP extraction utilize tissue grinding followed by a growing gradient of sucrose with a low ionic strength buffer to allow the sedimentation and isolation of the cell wall while preserving the ionic bonds. This gradient is able to eliminate organelles and other molecules less dense than the cell wall polysaccharides. Then, the last step is dedicated to washing away intracellular proteins that have remained trapped in the cell wall matrix through extensive washing on a polymer net. After cell wall isolation, the destructive protein extraction usually relies on salts such as calcium and lithium chloride to successfully liberate the wall-bound proteins [46]. Calcium chloride ($CaCl_2$) has the ability to collect CWPs because acidic and neutral carbohydrates strongly chelate calcium, and thus proteins weakly bound to the walls' polysaccharides can be solubilized by $CaCl_2$ through a competition mechanism [47]. In addition, lithium chloride ($LiCl$) is used to extract mostly hydroxyproline-rich glycoproteins [17]. The non-destructive CWPs extraction technique used for the monocots here revised is based on vacuum infiltration of the plant samples with solutions containing the same salts used for the destructive method mentioned above, followed by centrifugation of these samples to release the extracted CWPs [17].

Regarding mass spectrometry (MS), different strategies have been used for the study of monocot CWPs. These include previous separation by 1D-polyacrylamide gel electrophoresis (1D-PAGE) prior to tryptic digestion or shotgun analysis by liquid chromatography-tandem mass spectrometry
(LC-MS/MS). Following the identification of proteins through MS and bioinformatics, data analysis is essential for subcellular and functional predictions. Table 1 summarizes the plant samples, isolation of CWPs and MS techniques used in monocot CWP studies.

*B. distachyon* is a model species for temperate grasses with a fully sequenced genome, in addition to being easily grown and closely related to biomass crops. In this species, nine different plant samples have been analyzed, including young and mature leaves, apical and basal internodes, seeds and seedlings at different developmental stages [33–36]. These studies could point to specificities in different organs and developmental stages analyzed, especially in glycoside hydrolases and oxido-reductases. Fewer proteins were extracted from mature organs in comparison to young ones, which was attributed to a higher level of cell wall polymers cross-linking [33]. *Saccharum* spp. is a crop of special economic interest, as it is one of the major sources of sugar and bioethanol. For this crop, ten different plant samples were surveyed, including suspension cells culture, young and mature leaves, and basal and apical internodes at different developmental stages [37–40,48]. *O. sativa* was one of the first plant species with a sequenced genome and it is an economically relevant crop worldwide. *O. sativa* CWP surveys used five plant samples, such as suspension cells culture, culture media, roots and leaves [28–31]. In the next sections, when a CWP is mentioned as identified or not in monocots or *A. thaliana*, it means that this protein was or was not identified by MS in the mentioned CWP studies, respectively. When discussing the gene sequences corresponding to these CWPs or functional studies on them, this information is specified in the text and referenced properly.

### 3. Functional Class Distribution in Monocots

The number of non-redundant proteins was collected from each species and sorted into functional classes [28–31,33–40,48]. Although the results are not equally comparable, since different organs and several methods of CWP extraction and MS analysis were used, we were able to retrieve information regarding the functional classes and unique proteins in each species. Currently, the highest number of identified CWPs in monocots is from *B. distachyon*, comprising 594 proteins. *Saccharum* spp. is the second, with 283, and then *O. sativa*, with 270 identified CWPs. Altogether, 1159 CWPS proteins were identified, corresponding to 466 *A. thaliana* sequences. These proteins were divided into nine functional classes, according to Jamet et al. [7]: proteins acting on carbohydrates (PACs), oxidoreductases (ORs), proteases (Ps), proteins related to lipid metabolism (LMs), proteins possibly involved in signaling (Ss), proteins with predicted interaction domains (IDs), miscellaneous proteins (Ms), proteins of unknown function (UFs) and structural proteins (SPs). The functional classification of the CWPs from the monocots can be seen in Figure 1.
<table><caption>**Table 1.** Summary of the plant samples, extraction and mass spectrometry methods used in monocot CWP studies.</caption><thead><tr><th>Species</th><th>Plant Sample</th><th>Extraction Method</th><th>Pipeline of MS Analyses</th><th>Total of CWPs Identified</th></tr></thead><tbody><tr><td rowspan="2">B. distachyon</td><td>young and mature leaves, apical and basal internodes [33]</td><td>destructive technique followed by salt-based extraction</td><td>separation through 1D-PAGE and LC-MS/MS</td><td rowspan="2">594</td></tr><tr><td>young seedlings [35]</td><td>destructive technique followed by SDS and phenol-based extraction, and enrichment by ConA affinity chromatography</td><td>2D-LC-MS/MS</td></tr><tr><td rowspan="2">Saccharum spp.</td><td>seeds [36]</td><td>destructive technique followed by salt-based extraction</td><td>separation through 1D-PAGE and LC-MS/MS</td><td rowspan="2">594</td></tr><tr><td>seeds (9, 13 and 19 days after flowering (DAF)) [34]</td><td>destructive technique followed by salt-based extraction</td><td>separation through 1D-PAGE and LC-MS/MS</td></tr><tr><td rowspan="4">O. sativa</td><td>7 days cells suspension cultures [37]</td><td>destructive technique followed by salt-based extraction</td><td>2D-LC-MS/MS</td><td rowspan="4">283</td></tr><tr><td>2 month-old internodes [38]</td><td>destructive and non-destructive techniques followed by salt-based extractions</td><td>2D-LC-MS/MS</td></tr><tr><td>4 month-old young and mature leaves and apical and basal internodes [39]</td><td>destructive technique followed by salt-based extraction</td><td>2D-LC-MS/MS</td></tr><tr><td>7 month-old young and mature leaves [40,48]</td><td>destructive and non-destructive techniques followed by salt-based extractions</td><td>2D-LC-MS/MS</td></tr><tr><td>7 month-old apical and basal internodes [40,48]</td><td>non-destructive technique followed by salt-based extraction</td><td>2D-LC-MS/MS</td><td>283</td></tr><tr><td rowspan="4">5 days cultures medium of cell suspension cultures [28]</td><td>non-destructive technique followed by TCAAEB-based extraction</td><td>separation through 2D-PAGE and LC-MS/MS</td><td rowspan="4">270</td></tr><tr><td>non-destructive technique followed by salt-based extraction</td><td>separation through 1D-PAGE and 2D-LC-MS/MS</td></tr><tr><td>non-destructive technique followed by Tween-20, CTAB and TCAAEB-based extraction</td><td>separation through 2D-PAGE and LC-MS/MS</td></tr><tr><td>non-destructive technique followed by salt-based extraction</td><td>separation through 1D-PAGE and 2D-LC-MS/MS</td></tr><tr><td>3 week-old 4th leaf [28]</td><td>non-destructive technique followed by salt-based extraction</td><td>separation through 1D-PAGE and 2D-LC-MS/MS</td><td>270</td></tr><tr><td>2–3 weeks culture medium of cell suspension cultures [30]</td><td>non-destructive technique followed by salt-based extraction</td><td>separation through 2D-PAGE and MALDI-TOF/TOF MS</td><td>270</td></tr><tr><td>roots [31]</td><td>non-destructive technique followed by salt-based extraction</td><td>separation through 2D-PAGE and MALDI-TOF/TOF MS</td><td>270</td></tr></tbody></table>
<figure><img src="image_10.png" /><figcaption>**Figure 1.** Percentage of CWP functional classes in the species *A. thaliana* (a), *B. distachyon* (b), *Saccharum* spp. (c) and *O. sativa* (d). Plant drawings are not to scale. Functional class abbreviations: proteins acting on carbohydrates (PACs), oxidoreductases (ORs), proteases (Ps), proteins related to lipid metabolism (LMs), proteins possibly involved in signaling (Ss), proteins with predicted interaction domains (IDs), miscellaneous proteins (Ms), proteins of unknown function (UFs) and structural proteins (SPs).</figcaption></figure>

To allow some comparison among the different CWP on monocots, we performed BLASTP with *A. thaliana* protein sequences against *B. distachyon*, *O. sativa*, and *Saccharum* spp. sequences (using default parameters) and selected the first-ranked *A. thaliana* sequence. These monocot species were selected since they have a comparable number of CWP already identified by mass spectrometry with no particular treatment. WallProtDB [48] was the database used to retrieve these data. Only proteins predicted to be secreted were considered.

Taking the mentioned data together [28–31,33–40,48], as can be seen in Figure 1, the proportion of PACs inside the CWP were similar among the three species, and ranged from around 20 (*Saccharum* spp.) to 30% (*O. sativa*), which is comparable to *A. thaliana* (~24%). For *O. sativa* and *B. distachyon*, the percentages of ORs were similar to *A. thaliana*—about 12 to 13%. Conversely, *Saccharum* spp. presented the highest proportion of ORs—20%. The LM percentages in all three species varied from around 9 to 11%, which was also similar to that of *A. thaliana*. Except for *B. distachyon*, the percentages of Ps in monocots were slightly lower than in *A. thaliana*. IDs were much lower in all the monocots in comparison with *A. thaliana*. In grasses, the percentage of SP was much lower than in *A. thaliana*. The percentages of Ms and UFs were higher and lower in *Saccharum* spp. in comparison with the other species, respectively.

### 3.1. Proteins Acting on Carbohydrates

Regarding PACs, from the 297 identified in cell wall proteomes, 111 are non-redundant *A. thaliana* sequences identified through BLASTp. Of these, 79 were also identified in *A. thaliana* CWP studies. Among the 297, several members of Glycosyl Hydrolase (GH) families 1 and 17 were identified. Other families, such as GH5, 13, 16, 32, 35 and 38, were also present. GHs are proteins involved in cell wall carbohydrates remodeling and can be regulated during development. These families are used in enzymatic cocktails for biomass degradation in second-generation ethanol production [49]. Literature data suggests that grasses present fewer GH1, GH16, GH28 and GH35 members than dicots and more
GH5, GH13, GH18 and GH51 members [33,50], which is consistent with our comparative analysis on monocots vs. *A. thaliana*, in general.

The GH1 AT1G61820 is a β-glucosidase named BGLU46, identified in *A. thaliana* and *B. distachyon* CWP studies. It has been suggested that coniferin is the substrate for BGLU46 and BGLU45, and that monolignol glucosides are a source of storage monolignols instead of direct precursors of lignin in Angiosperms. These storage monolignols could be metabolized under stress conditions, which would lead to lignin synthesis de novo [51].

The proportion of GH3 was slightly higher in monocots. Some *Saccharum* spp. GH3 have been found to be possible β-xylosidases, which catalyze the hydrolysis of xylose from xylo-oligosaccharides. In barley, a type II-wall C3, a binding site for (1 → 3, 1 → 4)-β-D-glucans was identified [52].

GH5 are more numerous in monocots (Os05g0244500, Os10g0370500, Bradi2g31690, Bradi5g13550 and Bradi5g13560), and this data may correlate with the fact that (1,3) (1,4)-β-D-glucans are their putative substrates [53]. *O. sativa* studies identified several GH13 (Os08g0473600, Os09g0457800, Os08g0473900, Os09g0457400, Os06g0713800, Os02g0765600 and Os09g0457600), all α-amylases that are capable of hydrolizing 1,4-α-glucosidic linkages [49], which is consistent with *O. sativa* endogenous metabolism. Starch is a plant carbohydrate often linked to storage organs, and its breakdown is mediated by amylases. In grains, there is an extracellular matrix enriched in starch, which is degraded by secreted enzymes, the alpha-amylases [4].

*Saccharum* spp. GH17, such as GH1, are probably β-glucosidases [49]. The proportion of GH17 was also higher in monocots (see Supplementary Table S1 for accession numbers). The presence of GH17 in monocots is not surprising, since, as mentioned, type II cell walls present mixed (1,3)(1,4)-β-D-glucans as the principal hemicellulose [54], which is the substrate for GH17 that displays glucan-1,3-β-glucosidase activity. These enzymes are used in enzymatic cocktails for biomass deconstruction and are considered one of the most efficient enzymes in breaking glycosidic bonds in hemicelluloses [55].

GH18 substrates are not currently known, but they could be xylanase inhibitors that promote cell wall extension or chitinases involved in cell signaling or pathogen response [56-58]. The proteins Bradi3g26840, Bradi3g26850, Os10g0416100 and SCQGRT3044B10 are all GH18 that did not match any *A. thaliana* sequence record. Interestingly, these *B. distachyon* GH18 (Bradi3g26840 and Bradi3g26850) were phylogenetically grouped into a separate clade in the GH18 family, and indeed, *A. thaliana* did not present any GH inside this clade [50]. Thus, because of their high specificity, the GH18 enzymes are excellent candidates for further investigation in order to unravel the structure of monocot cell walls. Additionally, as grasses are used as raw materials for biofuels production, the identification of GH functions could be valuable in solving bottlenecks related to biomass deconstruction.

*AtCWIN1* (AT3G13790) is a cell wall invertase from the GH32 family, and was identified in all CWP studies, except in *O. sativa*. *AtCWIN1* regulates carbon partitioning by cleaving apoplastic sucrose and helping in the process of carbon import into the cell, which is the role of membrane sugar transporters [59].

GH families 16 and 35 have more members in the *A. thaliana* proteome, and a possible explanation is that their substrates are xyloglucans and galactans, the last related to pectins. Galactan is involved in xyloglucan structure and mediates the interaction between xyloglucan and cellulose, in *A. thaliana*. Interestingly, the *AtBGAL1* (AT3G13750) from GH35 family, identified in *A. thaliana* and *B. distachyon*, act with BGAL3, identified in *A.thaliana* only, during cell elongation [60]. Whether this enzyme was poorly identified in monocots because of the low content of xyloglucans needs further investigation. The principal xyloglucan β-galactosidase in *A. thaliana*, *AtBGAL10*, was also identified in all cell wall proteomes, with the exception of *O. sativa*.

More members of GH51 were found in *B. distachyon* (Bradi4g26270, Bradi1g63990, Bradi4g43710 and Bradi1g57017) than in *A. thaliana*. These GH51 are probably α-L-arabinofuranosidases [49], a type of enzyme used for monocots biomass deconstruction, as they are rich in arabinoxylans. α-L-arabinofuranosidases act on hydrolysis of α-L-arabinofuranoside in α-L-arabinosides, together with hemicellulases, resulting in hemicellulose hydrolysis [49]. It is important to mention that more
studies related to substrate specificities and protein structure are necessary to establish the function of
these proteins.

In addition to cellulose and hemicellulose, pectin is one of the main constituents of the primary cell wall. Wall porosity, charge density and microfibril spacing are some of the functional roles of pectin [61]. Consistently with the fact that type II-wall plants have lower pectin content, Pectin Methyl Esterases (PMEs) and Pectate Lyase-like proteins were more represented in *A. thaliana*. After the transport from the Golgi apparatus to the cell wall, pectin is partially deesterified by PMEs, exposing a carboxyl group on galacturonosyl residues and allowing the pectin to be stiffened by ionic crossbonding with calcium ions [62]. The degree of methylation impacts on the wall stiffening and access to enzymes [63].

3.2. *Oxidoreductases*

ORs mostly comprise several class III peroxidases (Prxs), multicopper oxidases, plastocyanins, berberine-bridge enzymes (BBEs) and blue copper-binding proteins. Prxs, part of large multigenic families, can either oxidize phenolic compounds, and consume hydrogen peroxide or generate reactive oxygen species [64]. They have been involved in several functional roles, such as cell elongation, lignin metabolism, stress responses and germination (reviewed by [64]). As Prxs are versatile proteins, they can both promote cell wall expansion or the crosslinking of its components, favoring cell wall strengthening [65]; it is difficult to establish a correlation between their higher or lower proportion and their metabolic function. Among the ORs, monocots show a slightly higher percentage of Prxs, which may be related to the fact that Poaceae presents additional groups of paralogous Prxs genes in comparison to *A. thaliana* and other dicots [66]. Sugarcane commercial varieties are highly polyploid and aneuploid plants, usually resulting from the interspecific hybridization of *Saccharum officinarum* and *S. spontaneum* [67]. The higher proportion of ORs in *Saccharum* spp., mostly Prxs, compared to the other monocots may be due the high level of ploidy of this crop, but this observation is speculative. Different Prxs were identified when using destructive and non-destructive CWP extraction in young *Saccharum* spp. culms [38], which was suggested to be due to a differential level of pectin-binding capacity, as Prx with a Ca<sup>+2</sup>-pectate binding domain would be more difficult to extract using the infiltration technique. Interestingly, *AtPrx34* (AT3G49120) and 36 (AT3G50990) were only identified in the *A. thaliana* cell wall proteome, which is consistent with the fact that the first has a putative binding site to the calcium-mediated conformation of a pectin structure [68], and the second is a promoter of pectin solubilization [69], and thus all these Prxs that have some level of relation with pectin are expected to be less numerous in type II walls.

Associated with lignin biosynthesis, *AtPrx52* (AT5G05340) [70] and the monocot homologs (Os01g0205900, Bradi3g09120, SCQSST3114C09) were present in all analyzed cell wall proteomes. Similarly, the proteins related to lignification *AtPrx72* [71] (Os01g0327100, Bradi2g40590 and SCEPRZ1011A06) and *AtPrx64* [72] (Bradi2g37060 and SCJFLR1035D05) were identified in the monocots. *AtPrx16* (Os04g0656800, Bradi1g33740 and SCJFLR1035D02) and *AtPrx53* (Os10g0109300 and Bradi1g68900) were both identified among *A. thaliana* and the monocots' CWP s. The first is related to germination [73] and the second to cell elongation inhibition and cell wall strengthening [74].

*AtPrx17* was also linked to lignin content when induced by the transcription factor AGAMOUS-LIKE15, which controls the lignification of tissues and changes the cell wall properties [75]. This could point to a conserved pathway in plants, as *AtPrx17* was identified in *A. thaliana* and in monocots' CWP studies. Also identified in the four cell wall proteomes, *AtPrx39* was linked to higher production of reactive oxygen species that led to cold tolerance [76], evidencing the multiple roles of peroxidases in plant development. Because of the numerous functions of Prxs, more targeted studies are needed to determine the reason of their high variety in the plant cell wall, both in dicots and monocots.

BBEs catalyze the formation of berberine bridges, but in plants their function is vastly unexplored.
In *A. thaliana*, it has previously been shown that some BBEs can be identified as monolignol
oxidoreductases, and are related to lignin formation [77]. A much higher proportion of BBEs is
found in *A. thaliana*. Phylogenetic analysis shows that not only does *A. thaliana* have more BBE members, but these enzymes also present several types of active sites, and few of type IV, which is exactly the one that is found most in grasses. In the course of evolution, in addition to expanding the amount of BBEs, it seems that the number of BBEs with active site type IV decreased and type I increased. Four BBEs only identified in *A. thaliana*—AT1G01980, AT4G20840, AT1G11770 and AT4G20830, the last with a type I active site—are able to inactivate oligogalacturonides, which is suggested to strengthen the immune response to fungal polygalacturonases [78], pointing to an evolved mechanism. Previously, it has been shown that BBEs in monocots seem to lack the catalytic and substrate coordination motifs linked to monolignol oxidoreductase activity, which was linked to lignin formation [77]. There are few functional studies in plant BBEs with type IV active sites, but fungal BBEs with this site have been related to oligosaccharide oxidation and plant immune response [79].

### 3.3. Proteins Related to Lipid Metabolism

Under this category, we highlight the Lipid Transfer Proteins (LTPs), Glycerophosphodiester Phosphodiesterases (GDPD)/GPDP-Like (GDPDL) and lipases GDSL. LTPs and lipases are proportionally more and less numerous in the monocot cell wall proteome, respectively [28–31,33–40,48]. LTPs are encoded by large multigenic families, which are considered to be essential to land colonization by plants, and are among the most abundant secreted proteins, but their exact in vivo role is still unclear. It has been suggested that LTPs mediate the transference and adhesion of molecules required for the composition of lipid barriers that are water-resistant, such as cutin, suberin and wax (reviewed by Edqvist et al. [80]). In the leaves of C4-metabolism plants, such as *Saccharum* spp. and *B. distachyon*, suberin surrounds the plasma membrane of bundle sheath cells, inhibiting CO₂ diffusion [4], which could be a possible explanation for increased LTPs in these species. Accordingly, in *Saccharum* spp., some LTPs were only identified in leaves. LTPs were also associated with lipid deposition for cell expansion, as their transcripts were differentially expressed in maize elongating internodes in comparison to non-elongating ones [81]. Curiously, homologs of LTP12 were identified in *B. distachyon* (Bradi4g25750) and *O. sativa* (Os12g0115100), but not in *A. thaliana* (AT3G51590), where it was thought to be pollen-specific [82]. Monocot homologs of AT5G01870 were unique to the suspension cell culture or young plants, which could be an indication that it is related to growing tissues. In *B. distachyon* and *Saccharum* spp., AT5G01870 homologs seem to be organ-specific, and are found only in leaves.

AtLTP3 was identified in all four cell wall proteomes analyzed. This protein negatively regulates plant defense mechanisms through the regulation of the antagonism between abscisic and salicylic acids, as it is induced by the former. LTP3 is proposed to be a disease-related marker, and it is also thought that LTP3 and 4 show some level of redundancy in plant immunity [83], but curiously, LTP4 was not identified in the monocots CWP5, and whether this overlay occurs in grasses could be an interesting research topic. Identified in all four species, the AtGDPDL3 (AT4G26690) is linked with lipid rafts in root-hair tip growth, suggesting that root hairs could be used as a model to study lipid rafts in plant development [84].

### 3.4. Proteases

Essentially, Ps break peptide bonds and control several relevant plant processes, such as protein transport, activity and half-lives [85], being generally divided into aspartyl (Asp), serine, cysteine, metallo and threonine proteases. The proportions of Asp Ps seem to be slightly higher in monocots. Phylogenetic analyses indicate that S8, C1A and A1 plant proteases functions were established even before the evolutive divergence of monocots and eudicots. This is corroborated by the conservation patterns of intron/exon arrangements and phylogeny analysis from monocots and dicots [85]. Accordingly, most of the proteases identified in the CWP studies from the monocots showed the highest identities with accessions also identified in the *A. thaliana* cell wall proteome (see Supplementary Tables S1–S4). However, only targeted functional analysis would reveal whether Ps display specific activities in monocots.
The senescence-associated subtilisin (AT3G14067) is a serine protease identified in all four species cell wall proteomes (Bradi2g51440/Bradi3g57140/Bradi3g57130, SCJFRT2057F03/SCRFHR1007E04, Os02g0779200), which confirms the fact that orthologs were identified in monocots crops [86]. Subtilisin was associated with the regulation of abscisic acid (ABA) signaling and drought tolerance [87], probably through a conserved mechanism between dicots and monocots, given the relevant role it plays.

The cysteine protease papain-like AtSAG2 (also named AALP) (AT5G60360) has been associated with senescence and necrotic cell death [88], as its expression increases along with leaf development. Reasonably, AtSAG2 was mostly identified in mature organs in *A. thaliana* and in the monocot cell wall proteomes, pointing to its use as a senescence marker in monocots.

### 3.5. Proteins with Interacting Domains

IDs encompass Pectin Methyl Esterase Inhibitors (PMEIs), proteins with leucine-rich repeat (LRR) and Lysm domains, protease inhibitors such as cystatins, Bowman-Birk inhibitors, lectins and jacalins. PMEIs were proportionally less numerous in monocots, as might be expected for pectin-poor type II walls. Interestingly, Hocq et al. [89] showed that AtPME19 is a strong inhibitor of AtPME3, which were both identified in *A. thaliana* and *B. distachyon* cell wall proteomes. It has been suggested that PMEIs diverge with respect to their ability to bind PME at different pHs, which results in varied modulation of the pectin structure. Additionally, they act in pairs formed by the enzyme plus its inhibitor, an interaction mode more tightly controlled by the structural determinants from the inhibitor than the enzyme [89].

*O. sativa* showed a higher proportion of proteins with Lysm domains, which was associated with both plant immunity and symbiotic interactions in this species [90]. All Bowman-Birk serine protease inhibitors (BBIs) were only identified in members of Poaceae and Fabaceae families [91], and thus they have only been found in *Saccharum* spp. (SCJFLR1013A04 and SCRUFL3062D08), *B. distachyon* (Bradi2g24810, Bradi1g03510, Bradi2g01920 and Bradi2g24820) and *O. sativa* (Os01g0132000) cell wall proteomes. Protease inhibitors regulate protease activities, and BBIs, in particular, display an essential role in defense mechanisms directed towards protection against pathogens and pests [91]. Another type of protease inhibitor, *AtCys6* homologs were identified in *B. distachyon* (Bradi2g52670) and *Saccharum* spp. (SCEPLR1051C09), which are associated with abiotic stresses and nucleic acid degradation [92].

Overall, the LRR-containing domain is conserved throughout evolution in the plants, displaying activity in the innate immune system through the sensing of pathogen-associated molecular patterns [93]. *AtPGIP1* (an LRR-domain protein) and its monocots corresponding accessions were identified in cell wall proteomes. *AtPGIP1* was associated with reduced damage caused by infection of a root nematode by inducing plant camalexin and indole-glucosinolate pathways [94]. The levels of the transcripts of another protein classified as ID, *AtCys-5*, are increased upon nematode infection [95] and ABA [96]. This cystatin was also identified in all four species' CWP surveys. Perhaps both proteins could be part of the conserved defense mechanisms against nematodes in dicots and grasses.

### 3.6. Proteins Possibly Related to Signaling

This class of CWP s is composed by fasciclin-like arabinogalactans (FLAs), leucine-rich repeat receptor protein kinases (LRR-RKs) (actually trans-membrane proteins), and COBRA-like proteins (COBLs), among others. The proportion of proteins from the S class is similar in all four species, being higher in *B. distachyon* (see Figure 1). In proportion, FLAs seem to be more numerous in the cell wall proteomes of monocots. However, phylogenetic analyses showed that *A. thaliana* has 21 and *O. sativa* 15 FLA genes with conserved functions [97]. FLAs are related to cell-to-cell adhesion, mechanical strength for secondary cell walls and cellulose biosynthesis [98], in addition to elasticity [99]. Identified in all four species, FLA1 is supposed to act on the lateral root and shoot formation in tissue culture [100], and two *B. distachyon* FLAs (Bradi2g00220 and Bradi4g33490) were only identified in internodes, indicating that they could display organ-specific activities. In contrast, COBLs were only
identified in *A. thaliana* CWPs, with the exception of one COBL in *O. sativa* (Os10g0497700). These are glycosylphosphatidylinositol-anchored specific plant proteins, and are associated with cell expansion and cellulose level of crystallinity, predominantly in elongating tissues [101]. More members of COBL are found in dicots, as several duplications occurred after the separation of dicots and monocots during evolution [102]. Recently, it has been demonstrated that a *Sorghum bicolor* COBL protein is linked to cellulose biosynthesis in the secondary wall, affecting plant mechanical strength [103], and this provided evidence of a cellulose-related role for this protein family in grasses. Overall, the LRR domain-containing proteins are conserved in both dicots and monocots [104].

### 3.7. Miscellaneous

Proteins with diverse functions are grouped under this category. Invariably, in several studies on CWPs, these consist of dirigent proteins, germins, thaumatins, gibberellic acid-stimulated proteins, purple acid phosphatases, phosphate-induced (phi) proteins, aldose epimerases, carbonic anhydrases, metallophosphoesterases, ribonucleases, pathogenesis-related proteins, low-molecular-weight cysteine-rich proteins and strictosidine synthases. Identified in all four species, AtPAP10 has been proven to be transcriptionally regulated by MYB-CC factors, which control plant responses to inorganic phosphate starvation [105]. A germin protein (AT1G72610, and the corresponding Os08g0460000 and Bradi3g37680) seems to be leaf-specific, as it was mostly identified in this organ. Another protein from the same family (AT1G18970 and the corresponding Os03g0804500) was found to be auxin-responsive [106].

Dirigent proteins have more members in *B. distachyon* and *Saccharum* spp. CWP surveys. The family of dirigent proteins is possibly linked to lignin polymerization [107], and its higher content may be associated with the presence of aromatic molecules such as ferulic acids in some type II primary walls [4].

Nucleoside phosphatases were only identified in *Saccharum* spp. One of them, SCCCRZ1C01H06, is an apyrase associated with calcium signaling and has been suggested to be a messenger for sucrose accumulation [108], which is consistent with this species' high sugar content.

### 3.8. Structural Proteins

The reduced number of SPs in CWP studies on monocots is probably due their low levels in type II walls [6]. They are usually covalently linked to the wall, and thus present extra difficulty with respect to extraction. Extensins (EXTs), Proline- and Glycine-Rich Proteins are the most represented structural protein families in *A. thaliana* CWPs. In monocots, 10 leucine-rich repeat extensins (LRXs) have been identified altogether. EXTs are basic cell wall glycoproteins, rich in hydroxyproline residues with alternating hydrophilic and hydrophobic motifs [109], and have been associated with cell wall strengthening after different stresses and with pectin to create a coacervate that may serve as a template for cell wall deposition [110], reinforcing their role in type I cell expansion. Intriguingly, they are one of the most relevant families of CWPs in C3. In addition, specifically in *B. distachyon*, EXTs are essential to embryo regeneration and germination [111]. Three (Bradi2g05080, Bradi3g03370 and Bradi2g42477) of the four LRXs found in *B. distachyon* CWP studies [33–36] were also identified in the bioinformatic work of Liu et al. [112]. In a previous analysis, eight LRX genes were identified in *O. sativa* genome and were thought to form two distinct clades for vegetative and reproductive organs, which could reflect adaptations to different cell wall types. In addition to their role in cell expansion, it is suggested that LRXs might act on cell differentiation [113].

### 3.9. Proteins of Unknown Function

UFs mostly consist of several proteins with Domains of Unknown Function (DUFs). DUF642, for example, is present in the three monocot species plus *A. thaliana* (AT3G08030, Bradi1g04670, Os03g0807700, SCCCCL4009G04, AT5G11420, Os01g0611000, SCCCLB1001G04, AT5G25460 and AT4G32460). According to this categorization, it would only be speculative to propose for them
a functional role, but this family is thought to be conserved, and has been proposed to be a new
family of carbohydrate-binding proteins [114]. Some of these proteins have shown their ability to
bind cellulose [114], have been shown to interact with PME [115], with auxin flux and hypocotyl
elongation [116] and germination [117].

Several CWPs classified as UFs possess the BURP domain. Identified in *O. sativa* and *B. distachyon* (Os01g0733500 and Bradi2g49000), AT5G25610 has high sequence identity with *GhRDL1*. *GhRDL1* interacts with a cotton α-expansin, and together they promote plant growth when overexpressed simultaneously [118]. The great number of UFs identified in the four species, and the fact that several UFs are suggested to have roles linked to the cell wall polysaccharides, reinforce the need for more functional studies on them to bring more information regarding the biology of monocot cell walls.

4. Applicative Aspects of Research on CWP s

Glycoside hydrolases are good candidates to be used in order to increase plant biomass or decrease recalcitrance destined to bioenergy production. Considering the enzymatic cocktails made from enzymes of microorganisms and the data on CWP s identified in monocots, the focus may be on GH families 1, 3, 17, 27, 35 and 51 [49]. The expression of a bacterial GH5 in A. thaliana led to a less recalcitrant wall without harming plant growth. It is suggested that the β-1,4 linkages of cellulose could be cut in the wall in an appropriate time and result in beneficial effects only [119]. As grasses present more GH5 and are the material used for biofuel production, they could be enzymes to watch with respect to their manipulation in the course of plant development.

As Prxs might be involved with lignification, they need to be mentioned when discussing ways to improve biomass and to facilitate its conversion. In this sense, *OsPrx38* overexpression in *A. thaliana* increased biomass and seed yield under arsenic stress [120], which could be an alternative pathway to be engineered.

The overexpression of a plantacyanin (AT2G02850, an OR), identified in the cell wall proteomes of all species analyzed here, led to decreased plant biomass and seed yield in *A. thaliana*. As this gene is regulated through a microRNA (miR408) whose overexpression results in augmented biomass, this could be further studied [121].

In a previous work, Endo et al. [122] fused the promoter of an LTP gene (AT3G18280), identified in *A. thaliana* and monocot CWP studies, named *AtTED4*, with several other genes. *AtTED4* was able to regulate the transition from the immature to the mature developmental stage, as expected for an early xylem-specific promoter, which could be used to target specialized genes in biomass engineering.

5. Conclusions

Although a comparison among a variety of studies, and modes of extraction, analysis and mass spectrometry techniques is still far from a complete understanding, this review compiled the most extensive studies on monocot CWP s, gathering their common characteristics and comparing them to *A. thaliana* data. As not all studies have used quantitative analysis, the abundance of these proteins should be considered in a more detailed way in the future before drawing conclusions. The different GH families and the low number of both pectin-related CWPs and SPs were related to the specific type II-wall characteristics, such as the presence of mixed β-D-glucans, and lower contents of pectin and structural proteins. Furthermore, specificities were indicated, such as the monocot proteins from the GH18 family, and some questions remain unanswered, such as the roles of the Prx, Bowman-Birk inhibitor and dirigent proteins in type II cell wall processes.

Despite the particularities displayed by type II cell walls, the nucleotide sequences of genes encoding some of the proteins that are lacking in grasses are conserved among dicots and monocots [123]. This observation points to the fact that grasses have the genetic capacity to produce xyloglucan, but it remains inactive [43]. Furthermore, we need to increase our knowledge of CWPs from both dicots and monocots to provide novel insights into the specialization of type II cell walls and their adaptive advantages. This would aid in genetically tailoring plants to improve efficiency and biomass for the
production of commercially important products such as second-generation biofuels. In addition to the gains brought by the first surveys on the CWPs of monocots, data mining and integration using several omics and protein-protein interaction studies are needed to establish the roles of CWPs.

**Supplementary Materials:** Supplementary materials can be found at http://www.mdpi.com/1422-0067/20/8/1975/s1. Table S1. List of CWPs identified in *A. thaliana* (retrieved from WALLPROTDB [48]) and the accessions from *B. distachyon*, *Saccharum* spp. and *O. sativa* identified through BLASTp analysis; Table S2. List of CWPs identified in *B. distachyon* (retrieved from WALLPROTDB [48]) and the *A. thaliana* accessions identified through BLASTp analysis; Table S3. List of CWPs identified in *Saccharum* spp. (retrieved from WALLPROTDB [48]) and the *A. thaliana* accessions identified through BLASTp analysis; Table S4. List of CWPs identified in *O. sativa* (retrieved from WALLPROTDB [48]) and the *A. thaliana* accessions identified through BLASTp analysis. The published and unpublished proteomics data were retrieved from WALLPROTDB [48]. Phytozome nomenclature [124] was used for *O. sativa*.

**Funding:** Financial support was provided by a BIOEN/FAPESP PRONEX project (Process number: 2008/56100-5) as well as by INCT—Instituto Nacional de Ciência e Tecnologia do Bioetanol, CNPq (Process number: 142784/2007-9) and FAPESP (Process number: 2007/59327-8).

**Conflicts of Interest:** The authors declare no conflict of interest.

**Abbreviations**

<table><tr><td>1D-PAGE</td><td>one-dimensional polyacrylamide gel electrophoresis</td></tr><tr><td>2D</td><td>two-dimensional</td></tr><tr><td>2D-PAGE</td><td>two-dimensional polyacrylamide gel electrophoresis</td></tr><tr><td>ABA</td><td>abscisic acid</td></tr><tr><td>Asp</td><td>aspartyl</td></tr><tr><td>BBE</td><td>berberine-bridge enzyme</td></tr><tr><td>BBI</td><td>bowman-birk serine protease inhibitor</td></tr><tr><td>CaCl2</td><td>calcium chloride</td></tr><tr><td>CWP</td><td>cell wall protein</td></tr><tr><td>COBL</td><td>COBRA-like protein</td></tr><tr><td>DAF</td><td>days after flowering</td></tr><tr><td>DUF</td><td>protein with domains of unknown function</td></tr><tr><td>ER</td><td>endoplasmic reticulum</td></tr><tr><td>EXT</td><td>extensin</td></tr><tr><td>FLA</td><td>fasciclin-like arabinogalactan</td></tr><tr><td>GDPD</td><td>glycerophosphodiester phosphodiesterase</td></tr><tr><td>GDPDL</td><td>glycerophosphodiester phosphodiesterase-like</td></tr><tr><td>GH</td><td>glycosyl hydrolase</td></tr><tr><td>ID</td><td>protein with predicted interaction domains</td></tr><tr><td>LiCl</td><td>lithium chloride</td></tr><tr><td>LC-MS/MS</td><td>liquid chromatography-tandem mass spectrometry</td></tr><tr><td>LM</td><td>protein related to lipid metabolism</td></tr><tr><td>LRR</td><td>leucine-rich repeat</td></tr><tr><td>LRR-RK</td><td>leucine-rich repeat receptor kinase</td></tr><tr><td>LRX</td><td>leucine-rich repeat extensin</td></tr><tr><td>LTP</td><td>lipid transfer protein</td></tr><tr><td>M</td><td>miscellaneous protein</td></tr><tr><td>MS</td><td>mass spectrometry</td></tr><tr><td>OR</td><td>oxidoreductase</td></tr><tr><td>P</td><td>protease</td></tr><tr><td>PAC</td><td>proteins acting on carbohydrate</td></tr><tr><td>PME</td><td>pectin methyl esterase</td></tr><tr><td>PMEI</td><td>pectin methyl esterase inhibitor</td></tr><tr><td>Prx</td><td>class III peroxidase</td></tr><tr><td>S</td><td>protein possibly involved in signaling</td></tr><tr><td>SP</td><td>structural protein</td></tr><tr><td>UF</td><td>proteins of unknown function</td></tr></table>
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Article

Cell Wall Proteome of Wheat Grain Endosperm and
Outer Layers at Two Key Stages of Early Development

Mehdi Cherkaoui<fnref n="1" />, Virginie Lollier<fnref n="1" />, Audrey Geiron<fnref n="1" />, Axelle Bouder<fnref n="1" />, Colette Larré<fnref n="1" />,
Hélène Rogniaux<fnref n="1" />, Elisabeth Jamet<fnref n="2" />, Fabienne Guillon<fnref n="1" /> and Mathilde Francin-Allami <fnref n="1" />,*

Received: 15 November 2019; Accepted: 23 December 2019; Published: 29 December 2019

**Abstract:** The cell wall is an important compartment in grain cells that fulfills both structural and functional roles. It has a dynamic structure that is constantly modified during development and in response to biotic and abiotic stresses. Non-structural cell wall proteins (CWPs) are key players in the remodeling of the cell wall during events that punctuate the plant life. Here, a subcellular and quantitative proteomic approach was carried out to identify CWPs possibly involved in changes in cell wall metabolism at two key stages of wheat grain development: the end of the cellularization step and the beginning of storage accumulation. Endosperm and outer layers of wheat grain were analyzed separately as they have different origins (maternal and seed) and functions in grains. Altogether, 734 proteins with predicted signal peptides were identified (CWPs). Functional annotation of CWPs pointed out a large number of proteins potentially involved in cell wall polysaccharide remodeling. In the grain outer layers, numerous proteins involved in cutin formation or lignin polymerization were found, while an unexpected abundance of proteins annotated as plant invertase/pectin methyl esterase inhibitors were identified in the endosperm. In addition, numerous CWPs were accumulating in the endosperm at the grain filling stage, thus revealing strong metabolic activities in the cell wall during endosperm cell differentiation, while protein accumulation was more intense at the earlier stage of development in outer layers. Altogether, our work gives important information on cell wall metabolism during early grain development in both parts of the grain, namely the endosperm and outer layers. The wheat cell wall proteome is the largest cell wall proteome of a monocot species found so far.

**Keywords:** cell wall; grain; remodeling; polysaccharide; proteome; wheat

**1. Introduction**

The wheat grain is a caryopsis composed of an embryo separated from the starchy endosperm and aleurone cells by the scutellum. The progeny is surrounded by the maternal tissues, which are composed of several outer layers [1]. They consist of the pericarp, the testa, and the nucellar epidermis [2]. The pericarp includes epidermal cells, parenchyma cells, and cross and tube cells. Cytological and physiological features of wheat endosperm and pericarp were characterized all along the grain development [3–5].

At the early developmental stage, the outer layers constitute most of the grain volume [3]. By around 150 growing degree days (GDD), the layers of the outer integument are the first to degenerate, whereas endosperm cells continue to divide. The beginning of grain filling occurs around 220 GDD. The aleurone layer becomes recognizable. Endosperm cells stop dividing and the grains

<footnote n="1">¹ INRAE, UR BIA, F-44316 Nantes, France; mehdicherkaoui04@gmail.com (M.C.); virginie.lollier@inra.fr (V.L.); audrey.geairon@inra.fr (A.G.); axelle.bouder@inra.fr (A.B.); colette.larre@inra.fr (C.L.); helene.rogniaux@inra.fr (H.R.); fabienne.guillon@inra.fr (F.G.)</footnote>

<footnote n="2">² Laboratoire de Recherche en Sciences Végétales, Université de Toulouse, CNRS, UPS, 31326 Castanet Tolosan, France; jamet@lrsv.ups-tlse.fr</footnote>

<footnote>* Correspondence: mathilde.francin-allami@inra.fr</footnote>
reach their maximum fresh mass at around 440 GDD. Concomitantly with the fusion of the pericarp with the maternal epidermis, the expanding endosperm compresses the outer layers. At grain maturity, except for the aleurone cells and embryo, all tissues degenerate, and only cell walls remain.

Cell walls play key roles during grain development. They have various compositions depending on the cell type and developmental stages. Due to their importance in nutrient transport, as well as cell size, shape, and stiffness, cell walls are essential for the determination of the final grain size and the nutritional quality of cereal grain [6–8]. They account for 3% of the endosperm, and around 70% of the outer layers in the mature wheat grain [9]. The cell walls are the primary contributors to cereal dietary fiber with important implications for human nutrition and disease prevention [10]. Cell walls also have important effects on cereal processes, such as milling, brewing, and breadmaking quality [11]. They are mostly composed of polysaccharides, together with a smaller amount of proteins, lipids, and phenolic compounds that participate in the cell wall metabolism.

Cell walls of developing endosperm have recently been described in detail [12–17]. Despite their important roles, both in grain physiology and from a nutritional point of view, cell walls of the outer layers of cereal grain have been poorly studied so far. Callose was the first polysaccharide to be detected during the development of wheat grain [18]. Arabinoxylans (AXs) and mixed β-glucans (MLGs) are the main polysaccharides of wheat grain, and of grasses more broadly. The remaining polysaccharides consist of cellulose, mannans, xyloglucans, and pectins [14,15,18]. MLGs are deposited early during the cellularization step and throughout the grain development. AXs are detected at the beginning of the differentiation in the endosperm cells. Spatial and temporal variations of the AX structure were observed in wheat grain. This structural heterogeneity is mainly explained by the variation in the substitution degree of AXs by arabinose, but also by ferulic acid, and to a lesser extent, p-coumaric acid, which binds to the arabinose residues [12,19]. Although less abundant than in dicot plants, cellulose was observed not only in the outer layers, but also in endosperm cells in a proportion that was not expected [20]. Recently, pectins were detected in the endosperm and the outer layers of wheat grain [14]. An impressive deposition of homogalacturonans (HGs) in the testa was reported at 250 GDD, with a decrease in their methylation degree during grain development. Rhamnogalacturonans of type I (RGI) were mainly detected in the endosperm and mostly in aleurone cells. Spatial and temporal variations of the RGI structure have been observed [14].

In addition to variability of the polysaccharide distribution and structures, the cell walls of the outer layers contain significant amounts of hydrophobic and polyphenolic polymers that compose the lignin and/or cutins. Lignification occurs earlier than previously reported in the outer layers and long before the grain reaches its final size [2]. Several cuticles are formed early in the development and persist at grain maturity. They are considered to be physically associated with the cell walls and have some overlapping functions. These cuticles are mainly composed of cutins and soluble waxes. Polysaccharides are also embedded into the cuticular layers [21]. Recently, xyloglucans were detected in the cuticle above the testa in the grain of the wild grass *Brachypodium distachyon* [22].

As described above, the composition and structure of cell wall polysaccharides vary during grain development and according to grain tissue to answer to the physiological needs of plants. Cell wall polysaccharide modifications imply numerous modifying enzymes, among which, glycosyl hydrolases (GHs) act on polysaccharides and oxidoreductases. They are mainly located in the cell wall and participate in the cell elongation, cuticle deposition, and cell wall structural variation that occurs during grain development. They may be transported by vesicles from the cytoplasm to the extracellular compartment [23]. The identification and functional understanding of these protein actors are essential to gain access to cell wall dynamics. To date, only a few experiments have been reported on grass cell wall proteomes: *Brachypodium distachyon* [24–26], *Saccharum* spp (sugarcane) [27–29], *Oryza sativa* (rice) [30–33], and *Triticum aestivum* (wheat) [34]. Transcriptomic data on the developing wheat grain allowed for the identification of GH genes [15], and recently, we reported the first data on the cell wall proteome of endosperm and the outer layers of wheat grain at a given developmental stage [34].
More than 600 cell wall proteins (CWPs) were identified, half of which were present in both tissues, suggesting common and specific tissue remodeling activities.

In this study, we investigated the cell wall proteome targeting both the endosperm and outer layers of the wheat grain at two key developmental stages in order to report on the cell wall dynamics during grain development. We identified proteins annotated as CWPs that are potentially involved in cell wall polysaccharide remodeling and cell wall assembly. A quantitative analysis allowed for describing the cell wall proteome content during the grain developmental stage for two tissues: the endosperm and the outer layers.

## 2. Results

### 2.1. Developing Grain Stages

The aim of this work was to identify CWPs in order to better understand the cell wall dynamics during the development of wheat grains. For this, the experiments were conducted from grains harvested at two developmental stages. The first one corresponded to an early stage of development (150 GDD, Figure 1), where the cellularization step was almost achieved but endosperm cells continued to divide. Aleurone and transfer cells were not yet differentiated. The outer layers derived from maternal tissues were already well differentiated. The outer layers were composed of the pericarp, the testa, and the nucellus epidermis (Figure 1).

The second developmental stage corresponded to the beginning of the accumulation of storage products. At this stage, around 250 GDD, endosperm was well differentiated. Aleurone and transfer cells were easily detected. In addition, the outer tissues progressively began to degenerate (Figure 1).

<figure><img src="image_13.png" /><figcaption>**Figure 1.** Tissue differentiation and development of wheat grain. Wheat grain cross-sections at (**left**) 150 and (**right**) 250 growing degree days (GDD) using bright-field micrographs of toluidine blue-stained sections. pe: pericarp, ep: external pericarp, pa: pericarp parenchyma, ne: nucellus epidermis, t: testa, al: aleurone layer, en: endosperm. Scale bars: 250 µm.</figcaption></figure>

### 2.2. Protein Extraction and Proteomic Analysis

For our analysis, the endosperm and outer layers were manually separated from grains at 150 and 250 GDD. Then proteins were extracted from three independent biological replicates per condition. Our procedure led to an enrichment of CWPs in the extracts. The electrophoretic profiles of replicates were similar and large number of bands was observed between 10 and 120 kDa (Figure S1). For each replicate, around 0.5 mg and 0.8 mg of proteins were obtained from 500 mg of fresh mass of endosperm and outer layers materials, respectively. Fifty micrograms of proteins were separated using 1DE (one dimension electrophoresis) and each lane was cut into five slices prior to being trypsin-digested and analyzed using LC-MS/MS. Only proteins identified as having at least two specific peptides were conserved. Interrogation of the UniProt databank restricted to *T. aestivum* led to the
identification of 2454 proteins (Table S2). The principal component analysis (PCA) indicated a good clustering of biological replicates and a significant discrimination between the four experimental conditions (Figure 2). The first two components explained 59.5% of the total variance. The different replicates clustered quite separately according to the tissue along the first principal component, and the development stage along the second principal component.

The predicted subcellular localization and functional annotation were achieved using the *ProtAnnDB* pipeline. A dataset of 734 total CWP s was selected, as described in Material and Methods (Table S2), which represents about 30% of all the 2454 identified proteins. In the outer layers, the CWP enrichment was 39%, while in the endosperm it was 22%. CWP s identified from the developing grains of *B. distachyon* represented around 40% of the total number of identified proteins, i.e., in the same range as what was obtained for the outer layers of the wheat grain [25]. Given that the complete genome annotation of wheat has currently not been achieved, a particularly large number of proteins were annotated as “uncharacterized protein”. By searching predicted functional domains using the *ProtAnnDB* pipeline, the annotation of our data was refined, leading to the functional annotation of a significant number of additional proteins, thus drastically decreasing the amount of “uncharacterized proteins”.

In addition, a statistical analysis of the quantitative data obtained for CWP s was performed in order to consider the variation of CWP abundance between the two developmental stages and the two parts of the grain. The relative quantification of proteins was performed from the signal intensity of their respective peptides in mass spectroscopy (MS), using a label-free and non-targeted approach. Quantification was achieved from a minimum of two peptides per protein. A total of 243 proteins was shown to present variation in their abundance according to parts of the grain and/or developmental stages (Table S3).

<figure><img src="image_14.png" /><figcaption>**Figure 2.** Principal component analysis (PCA) performed using the relative amounts of cell wall proteins (CWP s) for both parts of the wheat grain and at the two developmental stages (150 and 250 GDD). Individual maps of the biological replicates (red: endosperm samples at 150 GDD, green: endosperm samples at 250 GDD, dark blue: outer layers samples at 150 GDD, light blue: endosperm samples at 250 GDD) along principal components 1 and 2.</figcaption></figure>

### 2.3. *Validation of Cell Wall Localization for Three of the Identified Proteins*

In order to validate our proteomic analysis and bioinformatics treatment, we decided to check the cell wall localization for three randomly selected CWP s. These proteins were respectively annotated
as a pectin methylesterase (PME), a PME inhibitor (PMEI), and a glycoside hydrolase belonging to family 152 (GH152). We therefore generated C-terminal green fluorescent protein (GFP) fusions under the control of the cauliflower mosaic virus (CaMV) 35S constitutive promoter using an appropriate binary vector. The sequences used for the constructs encoded the full length CWPs, including their signal peptides. The constructs were transiently expressed in *N. tabacum* leaves after *A. tumefaciens* infiltration. The subcellular localization of the fusion proteins was analyzed using confocal laser scanning microscopy. The fluorescence of the protein fusions PME::GFP, GH152::GFP, and PMEI::GFP was detected at the periphery of the cells; this was consistent with their localization in the cell wall. A plasmolysis of the infiltrated leaf pieces was performed to separate the plasma membranes from the cell walls. The results indicated that the three fusion proteins were present in the cell wall and in the apoplastic compartment, as observed with the apoplastic marker Sec-mRFP (Figure 3).

<figure><img src="image_15.png" /><figcaption>**Figure 3.** Subcellular localization of a PMEI (A0A1D6BK49), a GH152 (A0A1D6DD47), and a PME (A0A1D5V0T8) fused to a green fluorescent protein (GFP) in plasmolyzed *N. tabacum* leaf epidermal cells. Confocal images of plasmolyzed cells expressing the apoplastic marker Sec-mRFP, PMEI::GFP, GH152::GFP, and PME::GFP in leaf epidermal cells 2–3 days after agroinfiltration. Scale bars: 20 µm.</figcaption></figure>

## 2.4. Distribution of the CWPs in Functional Classes

The CWPs are distributed into eight of the nine functional classes, as defined by the *ProtAnnDB* tool [35] (Figure 4A). No protein was annotated as a structural protein that has been currently observed in cell wall proteomes of monocots [36]. The most effective functional class was that of “proteins acting on cell wall polysaccharides” (PACs) with around a quarter of the CWPs. Inside this functional class, we mainly found GHs, which was consistent with previous cell wall proteome studies [25,26,28,37] (Table S2). PACs also contained expansins, carbohydrate esterases, and to a lesser extent, pectate lyases, PNGases (*N*-glycosidases), and COBRA-like proteins. Altogether, 136 GHs were identified. They belong to 21 different GH families (http://www.cazy.org/). They are thought to play various functions in cell expansion, signaling, plant defense, and the hydrolysis of glycans from glycoproteins and polysaccharides [38]. GHs were fairly distributed between the developmental stages and grain parts. Among them, the best-represented families were GH3 and GH17, as previously found in *B. distachyon* grains [24]. The classes of “proteases” and “proteins with interacting domains” were also well represented. The functional class of “signaling proteins” accounted for 4% of the total CWPs, as currently found in other cell wall proteome of monocots (3.4% for overall cell wall proteomes of monocots [39]; 6% if only the cell wall proteome of *B. distachyon* grains is considered [25]). Among the proteins of yet unknown function, DUF642 proteins were found as in all the previously studied cell wall proteomes (see WallProtDB).
<figure><img src="image_16.png" /><figcaption>**Figure 4.** Distribution of CWP s and variation of their abundance according to the different conditions. (A) Distribution of CWP s into functional classes according to their predicted functions. PACs: proteins acting on cell wall polysaccharides. (B) Heatmap of the relative amounts of CWP s identified in the four conditions (in the endosperm (EN), in the outer layers (OL), and at 150 and 250 GDD).</figcaption></figure>

## 2.5. Distribution of CWP s and Their Abundance Variation According to Tissue

Among the CWP s, almost two times more proteins were identified in the outer layers compared to the endosperm (Figure 5). More than 300 CWP s were shared between the two parts of the grain.

Among the CWP s identified in the endosperm, few of them were specific to this tissue. Several of them were annotated as proteins with leucine-rich repeats (LRR proteins), class III peroxidases, multicopper oxidases, or plant invertases/PME inhibitors, and others corresponded to the COBRA-like, DUF296, DUF239, and antimicrobial peptide (AMP) protein families. No GH family was found exclusively in the endosperm. In contrast, the outer layers accounted for numerous specific CWP s (Figure 5, Table S2) with proteins belonging to families of pectate lyases, acetyl xylan esterases (AXEs), GH10, GH13, GH32, laccases, and some DUF proteins (DUF642, 1929, 568, and 760).

Although numerous protein families were identified, both in the endosperm and outer layers, some of them differed by the number of their members between these two tissues. Some protein families were best represented in one of these two grain parts. This was the case for the fasciclin-like arabinogalactan proteins (FLAs), GDSL lipases/esterases, GH31, GH35, class III peroxidases, purple acid phosphatases (PAPs), and pectin acetylesterases (PAEs), which were more represented in the outer layers. Due to the relatively low number of CWP s identified in the endosperm compared to the outer layers, it was more difficult to reveal the protein families best represented in this tissue. However, our results clearly indicated that proteins with a protease-associated (PA) domain, α-amylase inhibitors, GH28, GH29, and plant invertases/PME inhibitors were well represented in the endosperm (Table S2).

Considering the developmental stages, only a third of the CWP s were shared by the endosperm and outer layers (Figure 5). This was partly due to the big difference in the numbers of CWP s identified in the endosperm and outer layers. The classes of "oxidoreductases," "proteins related to the lipid metabolism," and "proteins possibly involved in signaling proteins" were more numerous in the outer layers compared to the endosperm (Figure S2). Inversely, the class of "proteins with interacting domains" was best represented in the endosperm, with a significant proportion of CWP s annotated as plant invertase/PME inhibitors (Figure S2, Table S2).
From a statistical point of view, variations of the CWPs' abundance were observed between the endosperm and outer layers at a given developmental stage (150 GDD or 250 GDD) (Table S3, Figure S2). Among the CWPs shared by the endosperm and outer layers at 150 GDD, 60 were accumulated at a higher level in the outer layers, and only 28 in the endosperm. At 250 GDD, the numbers of CWPs accumulated at a high level in the outer layers and endosperm were closer (57 CWPs in outer layers versus 47 in endosperm), revealing a strong cell wall dynamic in both parts of the grain. More dirigent proteins, oxidoreductases, and thaumatins were accumulated at a higher level in the outer layers than in the endosperm, especially at 150 GDD. Some GHs (GH1, GH17, and GH79) were also accumulated in the outer layers, whatever the developmental stage. In the endosperm, plant invertases/PME inhibitors were found to have greater levels of accumulation compared to the outer layers, as well as some protease inhibitors and GH3.

<figure><img src="image_17.png" /><figcaption>**Figure 5.** Venn diagrams showing the distribution of CWPs according to the wheat developmental stages and grain parts.</figcaption></figure>

## 2.6. Distribution of CWPs and Their Abundance Variation According to the Developmental Stage

Few differences were noticed between the total number of CWPs identified at 150 GDD and 250 GDD (676 vs. 640 CWPs) (Figure 5, Table S2). A large majority of them were shared by the two stages. Only a few differences were observed for each protein family between the two developmental stages. However, a slightly higher number of expansins, lipid transfert proteins (LTPs), invertase/PME inhibitors, class III peroxidases, and plastocyanins was noticed at 150 GDD compared to 250 GDD. On the other side, arabinofuranosidases (GH51 and GH146), proteins with predicted protease-associated (PA) or cupin domains, and some proteases (subtilases) were slightly more represented at 250 GDD compared to 150 GDD. Whereas the GH10, DUF760, and DUF296 families were only identified at 150 GDD, lectins and COBRA proteins were only found at 250 GDD (Table S2).

Among the CWPs identified at 150 GDD, around one hundred were specific to this stage, with more proteins identified specifically in the outer layers compared to those identified in the endosperm (Figure 5, Table S2). While some invertase/PME inhibitors were specific to the endosperm at 150 GDD, several AXEs, class III peroxidases, and DUF538 proteins were specifically found in the outer layers at this earlier developmental stage. At 250 GDD, we identified 58 CWPs specific to this stage, with twice as many identified proteins in the outer layers compared to the endosperm, two arabinofuranosidases and two subtilases among them (Table S2).

Considering the endosperm, more than twice as many CWPs were significantly accumulated at higher level at 250 GDD compared to 150 GDD (52 vs. 21). The opposite was observed in the outer layers (Table S3, Figure S3). This could reflect an intensive cell wall metabolism in the outer layers at 150 GDD, whereas the cell wall metabolism in the endosperm seemed to be more dynamic at 250 GDD.
In this way, numerous proteases, but also FLAs, leucine rich repeat-containing proteins (LRR-proteins), and GH9 were accumulated at 250 GDD in the endosperm. Inversely, in the outer layers, proteins annotated as AXEs, GDSL lipases/esterases, or proteases were particularly abundant at 150 GDD (Table S3).

### 3. Discussion

In this work, we investigated the cell wall proteome of two distinct parts of the wheat grain. The outer layers and endosperm are composed of distinct cell layers that have different functions in grain. Thus, the cell wall structures are very different and we were expecting to obtain distinct CWP profiles between these two parts of grain. Two early developmental stages, 150 GDD (end of cellularization step) and 250 GDD (differentiation step and beginning of storage accumulation) were considered since clear differences at the histological and cell levels (Figure 1) and in the cell wall composition have been described between these two stages [12,14,17]. Although numerous cell wall events occur after 250 GDD, the accumulation of storage proteins, such as glutenins and gliadins in wheat endosperm, makes wall preparation and proteomic analysis more difficult. Indeed, due to the amount of storage proteins being particularly high from 300 GDD, the detection of CWPs, which are by weight minor cell wall components, would be very hard.

Although the outer layers can be easily detached from the endosperm of early harvested wheat via manual dissection [40], we cannot totally exclude protein cross-contaminations between these two tissues. However, the PCA indicated a strong discrimination between the endosperm and outer layers samples collected at 150 or 250 GDD, as well as a low variability between replicates (Figure 2). Because most of the polysaccharides were deposited both in the endosperm and outer layers, and since several cell wall remodeling events occurred in both grain tissues, numerous CWPs were expected to be identified in both of them. Forty-six percent of the identified CWPs were indeed found in both parts of grain.

To analyze the cell wall proteome, we performed cell wall enrichment and the proteins were extracted with CaCl₂ and LiCl buffers using a protocol successfully used on different plant species and tissues [25,26,28,29,37,41,42]. In this work, we identified a total of 2454 proteins, 734 of them being predicted as secreted proteins present in the cell wall or the apoplast, and called CWPs. CWPs were more numerous in the outer layers compared to the endosperm. The outer layers contained several cell layers, whereas endosperm tissues only contained starchy endosperm and aleurone cells. This could explain why more CWPs were identified in the outer layers, reflecting the presence of more different cell types. In addition, the outer layers contained more cell walls than the endosperm, leading to a higher protein amount. Also, we cannot exclude a better extractability of proteins from the outer layers than from endosperm. Conversely, almost the same number of CWPs were identified at both developmental stages of grains, with around 90% being shared proteins. These results, reinforced by the PCA data, suggest that in our experiments, the tissue effect was stronger than the developmental stage effect. However, many cell wall metabolism events differed from 150 to 250 GDD, such as the aleurone cell differentiation or the start of the pericarp degeneration, which may involve different CWPs [3,5]. Additionally, these distinct metabolic events were certainly reflected by differences in protein accumulation (Table S3). Many CWPs were more abundant in the endosperm at 250 GDD (in particular expansins, GH9, or GH28), while accumulation of a large number of CWPs was observed at 150 GDD in the outer layers (with numerous GDSL lipases/esterases, or oxidoreductases, such as class III peroxidases) (Table S3, Figure S3).

The distribution of wheat grain CWPs into functional classes slightly differed from those of the other cell wall proteomes. It appeared that the cell wall proteomes of grass grains (wheat and *B. distachyon*) contained more proteases and less oxidoreductases than the cell wall proteomes already described (Figure 4, [24–46]). The cell wall proteome of wheat grain also stood out as having numerous proteins with predicted interaction domains compared to other cell wall proteomes, including that of the *B. distachyon* grain [24,25]. Among proteins with interaction domains, more than 50 CWPs
belonged to the superfamily of the "plant invertase/PME inhibitors". Although found in all conditions, these proteins were more represented in the endosperm, and at 150 GDD. The transient expression of a fusion protein plant invertase/PME inhibitor::GFP in tobacco leaves confirmed its presence in the cell wall and in the apoplast (Figure 3). Wheat has a complex genome, with six copies of each chromosome and many nearly identical sequences scattered throughout, which could account for the large number of protein isoforms. As a matter of fact, many of the identified plant invertase/PME inhibitors present a strong homology. In wheat grain, there is a considerable number of plant invertase/PME inhibitors compared to other plant cell wall proteomes. In the cell wall proteomes of *A. thaliana* and *B. distachyon* (compilated data from WallProtDB), only 12 and 7 plant invertase/PME inhibitors were identified, respectively. In *B. distachyon*, they were all found in the developing grain [25]. In rice, several of the 54 predicted invertase/PME inhibitors are expressed specifically in developing endosperm [44]. PMEIs inhibit the demethylesterification of HGs by PMEs [45]. In wheat, PMEIs were suggested to play a role in plant defense mechanisms by decreasing the cell wall porosity and/or by conferring resistance against several pathogens [46,47]. Recently, two invertase inhibitor genes were identified in sugarcane, one of them encoding a protein (ShINH1) localized in the apoplast of the root, flower, stalk, and leaf. Heterologous expression of *ShINH1* confirmed its possible function as an invertase inhibitor [48]. In the same way, an apoplastic invertase inhibitor was shown to affect seed mass in soybean by regulating extracellular invertase during seed maturation [49]. Invertase inhibitor ZM-INVINH1 has been reported to bind cell wall invertase during grain development in maize [50]. Invertases belong to the GH32, GH68, and GH100 families. In our study, five GH32 that were predicted to be secreted were only found in the outer layers, which is consistent with a preferential presence of such proteins in the endosperm. On the other hand, the high amount of the plant invertase/PME inhibitors raises the possibility that they could regulate other carbohydrate metabolizing enzymes, as previously suggested [44].

Additional protein families were also well represented in the endosperm, such as GHs belonging to the GH9, GH28, and GH38 families, respectively annotated as endo-β-1,4-glucanases, polygalacturonases, and α-mannosidases. GH28 proteins were potentially active on HGs. Although present at the both developmental stages, some were more abundant at 250 GDD in the endosperm. It was already surprising to identify numerous GH28s in the cell wall proteome of the *B. distachyon* grain, at three developmental stages (9, 13, and 19 days after flowering), pectin also being poorly abundant in this plant species [25]. Although cell walls of grasses contain low level of pectins (HGs and rhamnogalacturonans), the presence of such enzymes is consistent with the remodeling of early synthesized pectins in the developing grain [14,17]. Two putative α-mannosidases belonging to the GH38 family were previously identified in the developing grain of *B. distachyon*, but no data are available concerning their fine localization [25]. Many plant α-mannosidases have been isolated and characterized from seeds and fruits. More generally located in the Golgi or endoplasmic reticulum, they are known to be involved in the maturation of N-glycans, and they appear to play a critical role in plant development [51,52]. Finally, the quantitative data indicated that GH9 were more abundant at 250 GDD than at 150 GDD in the endosperm. These enzymes are usually able to hydrolyze artificial soluble cellulose derivatives, such as carboxymethyl cellulose (CMC) or hydroxyethyl cellulose (HEC), and biochemical analyses have revealed their specificity for different substrates in vitro [53]. More generally, a large proportion of the endosperm CWPs were more abundant at 250 GDD than at 150 GDD, suggesting a more intensive cell wall metabolic activity in this tissue at this developmental stage of grain. This could be explained by the differentiation of the aleurone cells in addition to the expansion of the starchy endosperm cells at the beginning of storage accumulation.

Our work revealed significantly different cell wall proteomes between the endosperm and outer layers of the wheat grain, which could be largely explained by the differences of the cell wall structures between these two parts of grain. Thus, numerous protein families, highly represented in the cell wall proteome of outer layers, are known to act not only on cell wall polysaccharides but also on hydroxycinnamic acids, lignin, or cuticle formation, the last two being characteristic
of the outer layers. This was the case with the GDSL lipases/esterases being highly numerous at 150 GDD. These proteins are members of a large family that display numerous functions, such as cutin synthase, cutin hydrolase, deacetylation of polysaccharides, polysaccharide hydrolysis, fucosidase, and esterification of secondary metabolites [54]. They certainly collaborate to implement the cuticle during grain development, including the association of cutin with polysaccharides and the rearrangement of cutin and polysaccharides [54]. In wheat, cuticles were observed early during grain development [14], which is consistent with the presence of GDSL lipases/esterases at 150 GDD in outer layers. The accumulation of lipid transfer proteins (LTPs) in the outer layer matches with their role in the synthesis of cuticular waxes [21,55]. However, although less numerous, their presence in the endosperm suggest another function, such as antimicrobial defense or signaling during pathogen attacks [55]. Beside the cuticle, other non-polysaccharidic components are present in cell walls, such as lignin or hydroxycinnamic acids. Feruloylated AXs and lignins were indeed detected early in developing grain [14]. Current evidence suggests that two enzyme families, class III peroxidases and laccases, are involved in lignin polymerization [56]. These proteins were identified in the outer layers of developing grain in wheat. One laccase was identified only in the outer layers and numerous class III peroxidases were found mainly in the outer layers at 150 GDD with several of them more abundant at 150 GDD than at 250 GDD. Moreover, class III peroxidases could also be involved in AX crosslinking by ferulic acid, as previously shown in maize [57].

The outer layers and endosperm exhibit distinct functions that might explain differences of the cell wall proteomes. For instance, we found eight members of the GH19 family with four of them only identified in outer layers. In addition, the GH19 found in the two parts of the grain were always more abundant in the outer layers. These proteins are involved in the biotic stress response and they are known to be defense weapons against fungi by exhibiting chitinase activity [58]. These findings correlate with the protective role of outer layers throughout grain development. Among the protein families of the PAC class, many have a higher population in the outer layers. This is the case for the DUF642 proteins, which were identified only in the outer layers at both developmental stages. The DUF642 proteins belong to a well-conserved family. They were shown to be involved in plant development [59]. They may modulate the activity of PMEs, which are more abundant in the outer layers than in the endosperm, suggesting that their function could be related to the regulation of HG modifications by interacting with the catalytic domain of PME. Their interaction with cellulose was also demonstrated in vitro but no functional studies yet confirm these interactions [60]. Pectin acetysterases (PAEs) and AXEs were almost only found in outer layers. Acetylated AXs were detected in young wheat grain (245 GDD), but not at a mature stage (700 GDD) [61]. AXEs could participate in the deacetylation of AXs during the grain development. The deacetylation of AXs could improve the rigidity of the cell wall, as deacetylation of xylans facilitates their bonding to cellulose [62].

Other protein families, even though not specific to one developmental stage, are best represented at 150 GDD and in the outer layers, particularly the expansin, DUF538 proteins, and plastocyanins. In wheat, expansin genes were reported to be expressed at a high level during early grain expansion, with a peak of *TaExp6* transcript accumulation in the pericarp at 141 GDD, and a higher level of expression in the pericarp and endosperm at 187 GDD [7]. These results are consistent with the identification of more expansins at the cellularization step, and the presence in both the endosperm and outer layers. Moreover, the role of expansins in grain is reinforced by a recent study suggesting that expansin genes could be specifically involved in grain enlargement and may control seed growth in sunflower at the early phases of development [63]. As in wheat, several DUF538 were identified in the developing grain of *B. distachyon* [25]. Several functions have been suggested for proteins containing this domain of yet unknown function, such as their involvement in the photosynthetic system of plants by degrading chlorophyll molecules to antioxidant compounds under stress induction [64,65]. Initially characterized as chloroplastic proteins, some DUF538 proteins were shown to be in other cell compartments. Recently, a maize DUF538 protein, Glossy6, was demonstrated to be involved in cuticular wax accumulation and drought tolerance [66]. Proteins annotated as plastocyanin-like
proteins, also named plantacyanins, were also preferentially found at 150 GDD and in the outer layers. They are blue copper-binding proteins belonging to the oxidoreductase class, identified in numerous cell wall proteomes, such as in stems, leaves, and the developing grain of *B. distachyon* [24,26]. Recently it was shown that the blue copper proteins from the phytocyanin family were involved in the control of panicle branching and grain weight in rice through their regulation by the microRNA miR408 [67].

Proteases are well represented in the early developmental stages, and especially at 250 GDD. Endoprotease activities were characterized in developing wheat grain [68]. Serine protease activities were characterized at early stages of grain development, with metalloproteases and Asp proteases appearing later. We identified proteases belonging to the Asp-protease, Cys-protease, or serine carboxypeptidase families, both in outer layers and the endosperm. In the endosperm, several of them were more abundant at 250 GDD than at 150 GDD, suggesting stronger metabolic activities at 250 GDD in the endosperm cell wall. Endoproteases could participate in cell wall dynamics through the maturation of cell wall remodeling enzymes, their degradation, or the release of signaling peptides. In the case of subtilases, forming a large Ser protease family, it was suggested they were involved in the regulation of PMEs in *Arabidopsis* seeds [69,70]. A subtilase specifically found in the endosperm of *Medicago truncatula* or *Pisum sativum* seeds during development was shown to influence seed weight [71]. It was also suggested that endoproteases may be involved in the degeneration of the pericarp from the differentiation stage to the grain maturation, whereas in the endosperm, they may be accumulated for protein metabolism (processing or turn-over) or further degradation of storage proteins [68].

# 4. Material and Methods

## 4.1. Plant Material

*Triticum aestivum* (cv Recital) was grown under natural day-length conditions in a greenhouse (INRA Le Rheu, France). Two months of vernalization were applied in a growth chamber at 8 °C before the wheat seedlings were transplanted into individual pots containing a standard potting mixture (peat RHP15 Klassman, K Klassman France, Bourgoin Jallieu, France). An Osmocote (R) Exact Tablet containing nitrogen (15%), phosphate (9%), potassium hydroxide (9%), and magnesium (3%) (Scotts International B. V., Waardenburg, The Netherlands) was added. The plants were watered daily. To harvest grains at defined developmental stages, individual ears were tagged at flowering. The temperature was recorded every day in order to follow the plant development on the basis of the cumulated temperature in growing Celsius degrees days (GDD). Plants were harvested at 150 and 250 GDD after flowering. Grains were manually dissected to separate the outer layers and endosperm and immediately stored at -80 °C.

*Nicotiana tabacum* plants were grown under a 16-h light/8-h dark photoperiod at a temperature of 24 °C day/18 °C night for five weeks in a growth chamber prior to *Agrobacterium tumefaciens* infiltration.

## 4.2. Cell Wall Enrichment and Protein Extraction

Cell wall fractionation and CWPs extraction were performed as previously described [34]. Briefly, grain samples (500 mg) were ground and centrifuged in increasing sucrose concentrations to isolate cell walls. Proteins were extracted via successive washes in a 5 mM acetate buffer at pH 4.6, supplemented with 0.2 M CaCl₂ or 2 M LiCl. Three biological replicates were performed for each condition (developmental stages and parts of the grain).

## 4.3. Mass Spectrometry Analysis

Proteins (50 µg) were briefly separated in precast gels in order to reduce the mix complexity. Each track was cut into five fragments, which were reduced using dithiothreitol (DTT), alkylated by iodoacetamide and trypsinolyzed as previously described [34], prior to mass spectrometry (MS) analysis.
The trypsin-digested samples (6 µL) were injected into the LC-MS/MS system. The MS analyses were performed using an LTQ-Orbitrap VELOS mass spectrometer (i.e., a linear ion trap quadrupole mass filter associated with an Orbitrap™ analyzer, Thermo-Fisher Scientific, Bremen, Germany) coupled to a nanoscale LC system (Ultimate U3000 RSLC system, Thermo-Fisher Scientific). Chromatographic separation was performed on a reversed-phase capillary column (Acclaim® PepMapTM C18 2 µm 100 Å, 75 µm i.d. × 50 cm long, Thermo-Fisher Scientific, Waltham, MA, USA) at 60 °C using a linear gradient performed between the following mobile phases: (A) 97.9% water, 2% acetonitrile (ACN), 0.1% trifluoroacetic acid (TFA); and (B) 90% ACN, 0.08% formic acid. The other parameters were as described previously [72]. Analyses were performed using a typical survey method, in which full MS scans were acquired at 60,000 resolution (full width at half-maximum, FWHM) using the Orbitrap analyzer (m/z 300–2000), while the CID (collision induced dissociation) spectra for the eight most intense ions were recorded in the LTQ.

### 4.4. Bioinformatics Treatment, Functional Annotation, and Label-Free Quantification

The LC-MS/MS raw data were processed into mzXML files and were further searched against the Uniprot database restricted to *T. aestivum* (release May 2017, 136,892 accessions) and a contaminant database including human keratin and trypsin by using the X!Tandem pipeline software (version 3.4.3 "Elastine durcie", http://pappso.inra.fr/bioinfo/xtandempipeline/), which encapsulates the X!Tandem program and supplies a graphical interface to manage results [73].

Enzymatic cleavage was declared as a tryptic digestion with one possible miscleavage event. The carbamidomethylation of Cys residues, as well as the possible oxidation of Met residues, were considered in the modification parameters of the software. Precursor mass and fragment mass tolerance were set at 5 ppm and 0.5 Da, respectively. The results obtained for the peptide identification were validated by filtering the peptides with an e-value below $10^{-2}$. Proteins were identified using at least two specific peptides and an e-value score of $10^{-4}$.

The ProtAnnDB pipeline (http://www.polebio.lrsv.ups-tlse.fr/ProtAnnDB/) was used to annotate the identified proteins [35]. To predict the subcellular localization of the proteins, three software programs were used: TargetP and SignalP, which are both included in the pipeline, and Phobius (http://phobius.sbc.su.se/). Proteins were considered to be secreted when they had no ER retention signal and displayed a predicted signal peptide from their sequence by at least two out of the three software programs. In order to identify CWPs and annotate the dataset, three functional domains prediction software programs were used: Pfam, InterPro, and Prosite, which are all included in ProtAnnDB. The information provided by these tools allowed for the identification of CWPs in our dataset and classified them into nine functional classes. The proteomic data of the present work was included in the Pride database (http://www.proteomexchange.org/; dataset identifier: PXD016258).

Spectra with confident identifications were used for label-free quantification based on extracted ion chromatograms (XICs) computed using the MassChroQ tool (MassChroQ Gui, version 2.2.2) [74]. The resulting table of peptide XICs was then processed by the Rstudio statistical environment (version 1.0.143) (Boston, MA, USA). We first applied a filter to the outlying retention times of identified peptides to prevent possible false identifications. In this scope, the standard deviation of retention times for each of them was computed through the whole dataset and a 25 s threshold was applied. In a second step, we normalized the computed XICs on runs twice to remove any engine effect with the help of the experimental TIC, and on biological replicates in order to compare lanes. More precisely, we applied a simple correction factor on each computed XIC to place 1D lanes in the same range of values. The third step filtered the peptide population. Indeed, we chose to keep only peptides that were specific to one protein, assuming that the abundances of peptides should be correlated to the abundance of the related protein. Additionally, these peptides had to be identified in two of three biological replicates within at least one condition (developmental stages or grain parts). Finally, the relative abundance of a protein in a biological replicate was computed by summing the XIC values of the selected peptides.
4.5. Statistical Analysis

Statistical analysis was performed on log₁₀ transformed abundance values. To manage missing
values, we replaced them with a value mimicking a minimum mass spectrometer detection threshold
and by picking at random such a value in a range between the minimum value of the XIC dataset and
half of this minimum.

The ade4 package (version 1.7-6, http://pbil.univ-lyon1.fr/ade4/home.php?lang=eng) was used to perform PCA analysis on scaled data. In order to determine the CWPs showing a significant effect of developmental stages or grain parts on their abundance, one-way ANOVA was used. Protein abundances were considered to be significantly different for *p*-values below 0.05 and a data subset of “significant proteins” was created from this analysis.

We used this subset, restricted to CWPs to focus on, to build heat maps with the gplots package on covariance matrix (version 3.0.1) (Figure 4B, Figure S3).

4.6. Subcellular Localization

Total RNA was extracted from wheat grains harvested from 150 to 250 GDD using an RNA
kit (Qiagen, Courtaboeuf, France). RNA samples were then treated twice with a DNase buffer and
purified using the RNeasy MinElute Cleanup kit (Qiagen, Dusseldorf, Germany) by following the
manufacturer's instructions. Reverse transcription was carried out with 2 µg of total RNA, anchored
oligo-dT, and the Transcription First Strand cDNA synthesis kit (Roche Applied Science, Mannheim,
Germany). The binary vector pvKH18En6 was used to generate the constructs encoding the green
fluorescent protein (GFP) C-terminal fusion of the three proteins of interest. The full length cDNAs
encoding a pectin methylesterase (PME, A0A1D5V0T8), a PME inhibitor (PMEI, A0A1D6BK49), and a
glycoside hydrolase belonging to family 152 (GH152, A0A1D6DD47) were amplified via PCR using
specific oligonucleotide primers (Table S1). The corresponding amplicons were purified and cloned into
the pvKH18En6 binary vector [75] using the XbaI and BamHI restriction sites leading to the upstream
fusion to GFP. The construct Sec-mRFP was used as an apoplactic marker [76]. Transformation of
*A. tumefaciens* and the transient expression of the constructs in tobacco leaf epidermal cells were
performed as previously described [77]. Transformed *A. tumefaciens* cultures were resuspended in an
infiltration buffer at OD<sub>600nm</sub> = 0.05. Fluorescence was examined with an inverted Nikon A1 confocal
laser-scanning microscope (Nikon Europe, Amsterdam, The Netherlands) at three days after infiltration.
Infiltrated leaf samples were placed in a solution containing 30% glycerol 5 min before imaging with
the confocal microscope to plasmolyze the cells.

4.7. Histochemistry

Wheat grains at the developmental stages of approximately 150 GDD and 250 GDD were fixed
overnight in 3% (w/v) paraformaldehyde in a 0.1 M phosphate buffer at pH7.4, dehydrated in ethanol
series, and embedded with LR-White resin as previously described [14]. Transverse semi-thin sections
(1-µm thickness) of grains were cut with an ultracut (UC7, Leica, Nussloch, Germany) and stained
with Toluidine Blue O (1% in 2.5% Na₂CO₃ for 5 min, then washed in water). Stained sections were
observed using a Multizoom Macroscope (AZ100M, Nikon Europe, Amsterdam, The Netherlands)
under bright-field conditions.

5. Conclusions

Our quantitative proteomic work provides an overview of CWPs of early-developing grains of
wheat, both in the endosperm and the outer layers. It provides important knowledge for a better
understanding of cell wall deposition and remodeling during grain development that appears essential
to controlling cereal quality and yield. Certainly due to the different tissues that composed the
outer layers, more CWPs were identified, reflecting more numerous protein activities than in the
endosperm. Additionally, a higher level of accumulation of a significant number of CWPs was found
in the endosperm at 250 GDD, revealing strong metabolic activities in endosperm cell walls at this developmental stage. Altogether, many CWPs could be involved in the cell wall polysaccharide remodeling, as well as in lignin and cutin formation in the outer layers. In addition, an unexpected number of proteins annotated as plant invertase/PME inhibitors were found in the endosperm. Further characterization is required to confirm their precise function during grain development. To date, this is the most complete cell wall proteome from a monocot species, and the first one to be performed on separated tissues of cereal grains.

**Supplementary Materials:** Supplementary materials can be found at http://www.mdpi.com/1422-0067/21/1/239/s1.
Table S1: Sequences of oligonucleotide primers used for gene amplification. Table S2: (A) Proteins extracted from cell walls of the developing wheat grain (endosperm and outer layers) at two developmental stages and predicted to be secreted (the so-called CWPs). (B) Proteins extracted from the cell walls of the developing wheat grain (endosperm and outer layers) at two developmental stages and predicted to be intracellular. Table S3: Statistical analysis performed on the quantitative data corresponding to wheat CWPs identified at two developmental stages, both in the endosperm and outer layers. S3a. CWPs differentially accumulated in different grain parts at 150 GDD. S3b. CWPs differentially accumulated in different grain parts at 250 GDD. S3c. CWPs differentially accumulated in the endosperm at 150 and 250 GDD. S3d. CWPs differentially accumulated in the outer layers at 150 and 250 GDD. Figure S1: 1DE profiles of wheat grain proteins extracted from enriched cell wall fractions. Ten micrograms of proteins from each sample (three biological replicates (1-3) of proteins extracted from the outer layers (OL) or endosperm (E) at 150 GDD and 250 GDD have been separated by 1DE and stained with Coomassie Blue. Molecular mass markers (M) are in kDa. Figure S2: Distribution into functional classes of CWPs identified in the endosperm and the outer layers at 150 GDD and 250 GDD according to their predicted function. PACs: Proteins acting on cell wall polysaccharides. Figure S3: Heat maps of relative amounts of the CWPs identified in the endosperm or in the outer layers, and at one of two early developmental stages. En = endosperm samples; OL = outer layers samples; 150 = 150 GDD samples; 250 = 250 GDD samples.

**Author Contributions:** M.F.-A. coordinated the project and wrote and reviewed the manuscript; M.F.-A., F.G., C.L., and H.R. conceived and designed the experiments and edited the manuscript; M.C. and A.G. performed the experiments; A.B. performed the validation analysis; V.L. and M.C. conducted the bioinformatics and performed the related experiments; M.C. and M.F.-A. analyzed the data; E.J. reviewed and supervised the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Acknowledgments:** The authors are thankful to INRA (National Institute of Agronomic Research) for supporting their research work. EJ wishes to thank the CNRS and the Paul Sabatier-Toulouse 3 University for supporting her research. Wheat plants were cultivated at INRA of Rheu. LC-MS/MS analyses were performed on the Platform BIBS (Biology and Structural Biochemistry) in Nantes. The authors wish to thank the following contributors: Jean-Christophe Helleisen (INRA of Rheu) for growing wheat plants; Hélène San Clemente (LRSV, UPS) for her contribution to the functional annotation of proteins; and Camille Alvarado for providing images of toluidine blue-stained cross-sections of developing wheat grains.

**Conflicts of Interest:** The authors declare no conflict of interest.

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Article

# Organ and Tissue-Specific Localisation of Selected Cell Wall Epitopes in the Zygotic Embryo of *Brachypodium distachyon*

Alexander Betekhtin ¹,*, Anna Milewska-Hendel ², Joanna Lusinska ¹, Lukasz Chajec ³, Ewa Kurczynska ² and Robert Hasterok ¹

¹ Department of Plant Anatomy and Cytology, Faculty of Biology and Environmental Protection, University of Silesia in Katowice, 28 Jagiellonska Street, 40-032 Katowice, Poland; joanna.lusinska@wp.pl (J.L.); robert.hasterok@us.edu.pl (R.H.)

² Department of Cell Biology, Faculty of Biology and Environmental Protection, University of Silesia in Katowice, 28 Jagiellonska Street, 40-032 Katowice, Poland; anna.milewska@us.edu.pl (A.M.-H.); ewa.kurczynska@us.edu.pl (E.K.)

³ Department of Animal Histology and Embryology, Faculty of Biology and Environmental Protection, University of Silesia in Katowice, 28 Jagiellonska Street, 40-032 Katowice, Poland; lukasz.chajec@us.edu.pl

* Correspondence: alexander.betekhtin@us.edu.pl; Tel.: +48-32-2009-484

Received: 26 January 2018; Accepted: 1 March 2018; Published: 3 March 2018

**Abstract:** The plant cell wall shows a great diversity regarding its chemical composition, which may vary significantly even during different developmental stages. In this study, we analysed the distribution of several cell wall epitopes in embryos of *Brachypodium distachyon* (Brachypodium). We also described the variations in the nucleus shape and the number of nucleoli that occurred in some embryo cells. The use of transmission electron microscopy, and histological and immunolocalisation techniques permitted the distribution of selected arabinogalactan proteins, extensins, pectins, and hemicelluloses on the embryo surface, internal cell compartments, and in the context of the cell wall ultrastructure to be demonstrated. We revealed that the majority of arabinogalactan proteins and extensins were distributed on the cell surface and that pectins were the main component of the seed coat and other parts, such as the mesocotyl cell walls and the radicula. Hemicelluloses were localised in the cell wall and outside of the radicula protodermis, respectively. The specific arrangement of those components may indicate their significance during embryo development and seed germination, thus suggesting the importance of their protective functions. Despite the differences in the cell wall composition, we found that some of the antibodies can be used as markers to identify specific cells and the parts of the developing Brachypodium embryo.

**Keywords:** Brachypodium; cell wall; nucleus; zygotic embryo

## 1. Introduction

The cell wall is a protective layer that is located around the cell membrane, which is found in plant, fungi, bacteria, and archaea cells. The structure and functions of the cell wall were interconnected during the process of evolution in spite of the fact that its chemical structure, and to a lesser extent, its functions are diverse in different groups of organisms. In plants, its most fundamental function is to provide structural support and protection to the cell with some additional specific functions that may occur during plant development and cell differentiation [1].

The plant cell wall is a dynamic and highly specialised network that is formed by a heterogeneous mixture of cellulose, hemicelluloses, pectins and, to some extent, proteins and phenolic compounds. The cell wall that is formed during cell division is called the primary cell wall. In many plants, as/after
the cell completes its growth, additional layers of cellulose fibres are deposited on the inner surface of the primary cell wall, which produces a secondary cell wall. The cell wall composition in vascular plants is approximately 30% cellulose, 30% hemicellulose, and 35% pectins, and 1–5% structural proteins, on a dry weight basis [2,3]. However, the precise proportions of these compounds may differ significantly at different stages of plant development. For example, grass coleoptiles consist of 60–70% hemicelluloses, 20–25% celluloses, and 10% pectin substances and the grass endosperm cell wall may contain up to as much as 85% hemicelluloses. By contrast, the secondary cell walls are generally more cellulose-rich [4–7].

*Brachypodium distachyon* (Brachypodium) belongs to the Pooideae subfamily and is a well-established model species for the grasses. It has several features and advantages that make it useful for gaining a better understanding of the genetic, cellular and molecular biology of temperate climate zone cereals and forage crops [8]. There are many studies, which are often connected with the chemical composition of the Brachypodium cell wall [9–12]. A comparative study of the primary cell wall in the seedlings of Brachypodium, barley and wheat demonstrated similar relative levels and developmental changes of hemicelluloses [10]. Analyses of the Brachypodium proteome facilitate better understanding of the enzymes that are involved in cell wall remodelling during seed development; such research is of great importance for gaining better understanding of these processes in grasses and for finding the key components that are responsible for the size and weight of grass grains [9]. However, there is a dearth of information about the localisation of specific cell wall components at different stages of Brachypodium development.

Here, we characterise the chemical composition of the cell walls in Brachypodium embryos and describe the differences in the number of nucleoli that were observed in the cell nuclei in different parts of an embryo. We used light and transmission electron microscopy (TEM), histological and immunolocalisation techniques to analyse the distribution of selected pectins, arabinogalactan proteins (AGP), extensins, and hemicelluloses in the cell walls, internal cell compartments, and on the embryo surface.

## 2. Results and Discussion

### 2.1. *The Morphological and Histological Features of Brachypodium Embryos*

In their study, Wolny et al. [13] demonstrated that Brachypodium embryos are small in size, which makes their initial examination possible only by the use of a dissecting microscope. In this study, we distinguished the main parts of the embryo, such as scutellum, V scale, coleoptile, first and second leaf, shoot apex, mesocotyl, epiblast, radicula, root cap, and coleorhiza (Figure 1). The coleoptile and coleorhiza are two organs that are found exclusively in grass species [14]. A comparison of the cell nuclei in different parts of Brachypodium embryos demonstrated that the majority contained only one nucleolus (Figure 1). However, some cells of the shoot apex, mesocotyl, radicula and root cap were characterised by the presence of a round nuclei that contained two nucleoli (Figure 1; nucleoli indicated by red arrows). TEM analysis of the selected embryo parts confirmed these observations and demonstrated the presence of a centrally positioned nucleus with one or two large nucleoli as well as a high nucleus:cytoplasm ratio (Figure 2a,b). The cytoplasm of these cells was dense and contained lipid droplets and starch granules around the nucleus. Interestingly, we also found cells in the embryo with nuclei that were extended in their shape but that also contained two nucleoli (Figure 2c). The architecture of these cells is typical for the initial vascular tissue [15]. According to Verdeil et al. [16], pluripotent plant stem cells, which are located within the root and shoot meristems, are isodiametric, have a dense cytoplasm, a high nucleus:cytoplasm ratio, a fragmented vacuome, contain granules of starch, and have a spherically-shaped nucleus with one or two nucleoli. Both of the meristematic cells of the oil palm (*Elaeis guineensis*) [17], maize (*Zea mays*) [18], and onion (*Allium cepa*) [19] have similar characteristics to those described by Verdeil et al. [16], which appear to be universal for monocots.
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**Figure 1.** Schematic representation of Brachypodium embryo. Asterisks in different colours represent nuclei from histological sections of specified parts of the embryo. Red arrows mark two nucleoli. Purple arrows show respective parts of the embryo: V scale, coleoptile, first leaf, second leaf, shoot apex, mesocotyl, epiblast, coleorhiza, radicula, and root cap. Bars: 5 µm.

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**Figure 2.** TEM (transmission electron microscopy) of radicula nuclei (**a–c**). Abbreviations: CW—cell wall, LD—lipid droplets, *N*—nucleus, *Nu*—nucleolus, *PI*—plastid. Bars: (a) 3 µm; (b) 1.5 µm; and (c) 2.5 µm.

## 2.2. AGP, Extensin, Pectin, and Hemicellulose Epitopes in Various Tissues and Organs of Brachypodium Embryos

AGPs distribution is dynamically regulated during the ontogenesis of a plant [20]. Although we demonstrated that AGPs have a diverse localisation in Brachypodium embryos, most of them
are found on the cell surface. The MAC207 antibody had an affinity to all parts of the embryo and was localised in the internal cell compartments (Figure 3a-a"). JIM8, JIM13, and JIM16 were detected mainly in outer pereclinal walls of surface embryo tissues (Figure 3b-b",c-c",d-d" and Figure 4a-a",c-c"). The JIM13 (Figure 3d-d") and JIM16 (Figure 4c-c") epitopes were components of the seed coat. Furthermore, the JIM16 epitope was localised in the internal cell compartments (Figure 4a-a",b-b"). Compared to most of the analysed AGPs, the LM2 epitopes were found only in the cell wall compartments (Figure 4d-d",e-e"). The possible functions of the AGPs on the embryo surface are to protect it from infections, contribute to plant-microbe interactions, seed germination and other important processes during plant development [20]. In their study of transcriptome and metabolome changes in *Arabidopsis thaliana* (Arabidopsis) that were connected with the seed dormancy and germination, Joosen et al. [21,22] showed that AGPs were more connected with the embryo cell walls. It is also known that AGPs are crucial in preventing infections in *Brassica napus* and *Pisum sativum* via the encystment of the *Aphanomces euteiches* zoospores, thereby inducing plant germination [23]. Van Hengel et al. [24] demonstrated that AtAGP30 is a non-classical AGP core protein in Arabidopsis, which had a root-tip specific expression in seedlings. This may imply its importance in root development or growth.

<figure><img src="image_22.png" /><figcaption>**Figure 3.** Immunolocalisation of arabinogalactan proteins in Brachypodium embryos. (a—a′) MAC207 in coleorhiza; (b—b′) JIM8 signal in coleoptile; (c—c″,d—d″) JIM13 in the entire embryo and the coleoptile with the seed coat, respectively. Abbreviations: CF—calcofluor, Cr—coleorhiza, Col—coleoptile, TSC—the seed coat. Bars: (a—a",b—b",d—d") 10 µm; and (c—c") 100 µm.</figcaption></figure>
<figure><img src="image_23.png" /><figcaption>**Figure 4.** Immunolocalisation of arabinogalactan proteins in Brachypodium embryos. (a—a",b—b",c—c") JIM16 with coleorhiza with radicula, radicula with root cap and coleoptile with the seed coat, respectively; Both (d—d") shoot apex and (e—e") radicula with LM2. Abbreviations: CF—calcofluor, Cr—coleorhiza, Col—coleoptile, Rad—radicula, RC—root cap, SA—shoot apex, and TSC—the seed coat. Bars: 10 µm.</figcaption></figure>

Extensins are one of the major plant cell wall protein families found to be secreted into the cell wall. They are glycoproteins, in which about one-third of all amino acids is hydroxyproline with four monosaccharide arabinose residues attached. These compounds seem to play a strengthening and/or defensive role during the response to disease and wounding [25–27]. We observed that most of the Brachypodium extensins in this study had a surface localisation, which was similar to that of the AGPs. The signals of the LM1 antibodies were localised on the surface of the coleoptile and first leaf (Figure 5a—a",b—b"). Furthermore, a high level of LM1 binding was observed in the internal compartments of the coleoptile epidermal cells (Figure 5b—b"; red arrows). The cell wall localisation was a specific feature of the JIM11 and JIM12 epitopes in the mesocotyl (Figure 5c—c" and Figure 6c—c"). Signals of the first antibodies were also observed in the cell walls of the radicula (Figure 5d—d"). Additionally, strong fluorescence signals of these antibodies were also found in the inner layer of the seed coat (Figure 5e—e"). The JIM12 extensin epitope had a surface distribution on the coleoptile, but no signals of these antibodies were observed in any part of the second leaf (Figure 6a—a",b—b"). It is still not clear whether extensins have redundant or specific functions in the different cell types of various plant tissues. The inhibition of both extensin and AGP biosynthesis by 3,4-dehydro-L-proline impedes
the development of embryogenic cells and decreases the rate of embryo germination [28]. This research demonstrated that extensins appear to play a vital role in the regeneration and germination of embryos during early plant development via somatic embryogenesis. It is also known that plants increase the amount of extensins when they are under aluminium stress. Sujkowska-Rybkowska et al. [29] suggested the importance of extensins in the aluminium resistance mechanisms and demonstrated an aluminium-induced extensin accumulation in the cell walls. Additionally, some extensins protect the plant cells from pathogen attacks [30]. An analysis of the extensin transcript levels in banana (Musa spp.) demonstrated that these proteins mainly appear in the root cap and meristematic cells after wounding or inoculation with *Fusarium oxysporum* [30]. The level of extensin transcription in intact plants were found to be up-regulated by wounding or inoculation with a pathogen, but down-regulated by a pathogen attack on wounded plants. In general, the connection of extensins with other components of the cell wall, as well as many of their functions during the development of the cell wall architecture are not yet fully understood [26]. It is likely that more extensive studies using mutants with the knocked down genes encoding extensins would shed more light on these matters.

<figure><img src="image_24.png" /><figcaption>**Figure 5.** Immunolocalisation of extensins in Brachypodium embryos. Both (*a—a''*) coleoptile and (*b—b''*) coleoptile with first leaf with LM1; (*c—c''*,*d—d''*,*e—e''*) mesocotyl, radicula and coleorhiza (including the seed coat) with JIM11, respectively. Arrows show outer layers of coleoptile including protodermal cells (*b—b''*). Abbreviations: CF—calcofluor, Col—coleoptile, Cr—coleorhiza, FL—first leaf, Mes—mesocotyl, Rad—radicula, and TSC—the seed coat. Bars: 10 µm.</figcaption></figure>