This review focuses on the responses of the plant cell wall to several abiotic stresses including drought, flooding, heat, cold, salt, heavy metals, light, and air pollutants. The effects of stress on cell wall metabolism are discussed at the physiological (morphogenic), transcriptomic, proteomic and biochemical levels. The analysis of a large set of data shows that the plant response is highly complex. The overall effects of most abiotic stress are often dependent on the plant species, the genotype, the age of the plant, the timing of the stress application, and the intensity of this stress. This shows the difficulty of identifying a common pattern of stress response in cell wall architecture that could enable adaptation and/or resistance to abiotic stress. However, in most cases, two main mechanisms can be highlighted: (i) an increased level in xyloglucan endotransglucosylase/hydrolase (XTH) and expansin proteins, associated with an increase in the degree of rhamnogalacturonan I branching that maintains cell wall plasticity and (ii) an increased cell wall thickening by reinforcement of the secondary wall with hemicellulose and lignin deposition. Taken together, these results show the need to undertake large-scale analyses, using multidisciplinary approaches, to unravel the consequences of stress on the cell wall. This will help identify the key components that could be targeted to improve biomass production under stress conditions.

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Mutants at the PROCUSTE1 ( PRC1 ) locus show decreased cell elongation, specifically in roots and dark-grown hypocotyls. Cell elongation defects are correlated with a cellulose deficiency and the presence of gapped walls. Map-based cloning of PRC1 reveals that it encodes a member ( CesA6 ) of the cellulose synthase catalytic subunit family, of which at least nine other members exist in Arabidopsis. Mutations in another family member, RSW1 ( CesA1 ), cause similar cell wall defects in all cell types, including those in hypocotyls and roots, suggesting that cellulose synthesis in these organs requires the coordinated expression of at least two distinct cellulose synthase isoforms.

Mutants at the PROCUSTE1 (PRC1) locus show decreased cell elongation, specifically in roots and dark-grown hypocotyls. Cell elongation defects are correlated with a cellulose deficiency and the presence of gapped walls. Map-based cloning of PRC1 reveals that it encodes a member (CesA6) of the cellulose synthase catalytic subunit family, of which at least nine other members exist in Arabidopsis. Mutations in another family member, RSW1 (CesA1), cause similar cell wall defects in all cell types, including those in hypocotyls and roots, suggesting that cellulose synthesis in these organs requires the coordinated expression of at least two distinct cellulose synthase isoforms.

Primary structures of the N-glycans of two major pollen allergens (Lol p 11 and Ole e 1) and a major peanut allergen (Ara h 1) were determined. Ole e 1 and Ara h 1 carried high mannose and complex N-glycans, whereas Lol p 11 carried only the complex. The complex structures all had a ␤(1,2)-xylose linked to the core mannose. Substitution of the proximal N-acetylglucosamine with an ␣(1,3)-fucose was observed on Lol p 11 and a minor fraction of Ole e 1 but not on Ara h 1. To elucidate the structural basis for IgE recognition of plant N-glycans, radioallergosorbent test analysis with protease digests of the three allergens and a panel of glycoproteins with known N-glycan structures was performed. It was demonstrated that both ␣(1,3)-fucose and ␤(1,2)-xylose are involved in IgE binding. Surprisingly, xylose-specific IgE antibodies that bound to Lol p 11 and bromelain did not recognize closely related xylose-containing structures on horseradish peroxidase, phytohemeagglutinin, Ole e 1, and Ara h 1. On Lol p 11 and bromelain, the core ␤-mannose is substituted with just an ␣(1,6)-mannose. On the other xylose-containing N-glycans, an additional ␣(1,3)-mannose is present. These observations indicate that IgE binding to xylose is sterically hampered by the presence of an ␣(1,3)-antenna.In the early 1980s, it was reported for the first time that IgE antibodies in sera of pollen allergic patients can be directed to carbohydrate determinants on glycoproteins (1-3). The carbohydrate nature of these epitopes was supported by several characteristic properties, such as their periodate sensitivity and their resistance to heating and protease digestion. IgE antibodies directed to these carbohydrate structures were shown to be extremely cross-reactive not only between different plant-derived glycoproteins but also to glycoproteins from invertebrate animals (e.g. seafood and insect venoms) (2, 4 -7).This high degree of cross-reactivity was explained by the conserved structure of N-glycans from plants and invertebrate animals, sharing several features that are not found in mammalian N-glycans (8). More recently, several research groups have confirmed the role of carbohydrate epitopes in IgE reactivity (9 -22).In plants, the N-glycosylation of proteins starts by the transfer of the oligosaccharide precursor Glc 3 Man 9 GlcNAc 2 in the endoplasmic reticulum (reviewed in Ref. 23). This structure can subsequently be modified by glycosidases and glycosyltransferases during transport of the glycoprotein through the endoplasmic reticulum, the Golgi apparatus, and the vacuole. Depending on the accessibility of the glycan side chain, these enzymes can convert the precursor to high mannose-type Nglycans ranging from Man 9 GlcNAc 2 to Man 5 GlcNAc 2 and then to complex-type N-glycans having an ␣(1,3)-fucose attached to the proximal glucosamine residue and/or a ␤(1,2)-xylose residue attached to the ␤-mannose. These linkages of fucose and xylose are typical for complex N-glycans from plants and invertebrate animals and are not found in mammals. Mo...

N-glycosylation is a major modification of proteins in plant cells. This process starts in the endoplasmic reticulum by the co-translational transfer of a precursor oligosaccharide to specific asparagine residues of the nascent polypeptide chain. Processing of this oligosaccharide into high-mannose-type, paucimannosidic-type, hybrid-type or complex-type N-glycans occurs in the secretory pathway as the glycoprotein moves from the endoplasmic reticulum to its final destination. At the end of their maturation, some plant N-glycans have typical structures that differ from those found in their mammalian counterpart by the absence of sialic acid and the presence of beta(1,2)-xylose and alpha( 1,3)-fucose residues. Glycosidases and glycosyltransferases that respectively catalyse the stepwise trimming and addition of sugar residues are generally considered as working in a co-ordinated and highly ordered fashion to form mature N-glycans. On the basis of this assembly line concept, fast progress is currently made by using N-linked glycan structures as milestones of the intracellular transport of proteins along the plant secretory pathway. Further developments of this approach will need to more precisely define the topological distribution of glycosyltransferases within a plant Golgi stack. In contrast with their acknowledged role in the targeting of lysosomal hydrolases in mammalian cells, N-glycans have no specific function in the transport of glycoproteins into the plant vacuole. However, the presence of N-glycans, regardless of their structures, is necessary for an efficient secretion of plant glycoproteins. In the biotechnology field, transgenic plants are rapidly emerging as an important system for the production of recombinant glycoproteins intended for therapeutic purposes, which is a strong motivation to speed up research in plant glycobiology. In this regard, the potential and limits of plant cells as a factory for the production of mammalian glycoproteins will be illustrated.

The mixed‐linkage (1→3),(1→4)‐β‐d‐glucans are unique to the Poales, the taxonomic order that includes the cereal grasses. (1→3), (1→4)‐β‐Glucans are the principal molecules associated with cellulose microfibrils during cell growth, and they are enzymatically hydrolyzed to a large extent once growth has ceased. They appear again during the developmental of the endosperm cell wall and maternal tissues surrounding them. The roles of (1→3),(1→4)‐β‐glucans in cell wall architecture and in cell growth are beginning to be understood. From biochemical experiments with active synthases in isolated Golgi membranes, the biochemical features and topology of synthesis are found to more closely parallel those of cellulose than those of all other noncellulosic β‐linked polysaccharides. The genes that encode part of the (1→3),(1→4)‐β‐glucan synthases are likely to be among those of the CESA/CSL gene superfamily, but a distinct glycosyl transferase also appears to be integral in the synthetic machinery. Several genes involved in the hydrolysis of (1→3),(1→4)‐β‐glucan have been cloned and sequenced, and the pattern of expression is starting to unveil their function in mobilization of β‐glucan reserve material and in cell growth.

Cellulose microfibrils are para-crystalline arrays of several dozen linear (1→4)-β-d-glucan chains synthesized at the surface of the cell membrane by large, multimeric complexes of synthase proteins. Recombinant catalytic domains of rice (Oryza sativa) CesA8 cellulose synthase form dimers reversibly as the fundamental scaffold units of architecture in the synthase complex. Specificity of binding to UDP and UDP-Glc indicates a properly folded protein, and binding kinetics indicate that each monomer independently synthesizes single glucan chains of cellulose, i.e., two chains per dimer pair. In contrast to structure modeling predictions, solution x-ray scattering studies demonstrate that the monomer is a two-domain, elongated structure, with the smaller domain coupling two monomers into a dimer. The catalytic core of the monomer is accommodated only near its center, with the plant-specific sequences occupying the small domain and an extension distal to the catalytic domain. This configuration is in stark contrast to the domain organization obtained in predicted structures of plant CesA. The arrangement of the catalytic domain within the CesA monomer and dimer provides a foundation for constructing structural models of the synthase complex and defining the relationship between the rosette structure and the cellulose microfibrils they synthesize.