Advances in Biological Chemistry
Vol.06 No.03(2016), Article ID:67461,36 pages
10.4236/abc.2016.63008

Plant Cell Wall, a Challenge for Its Characterisation

Guy Costa*, Idelette Plazanet

Laboratoire de Chimie des Substances Naturelles (EA 1069), Faculté des Sciences et Techniques, Limoges Cedex, France

Copyright © 2016 by authors and Scientific Research Publishing Inc.

This work is licensed under the Creative Commons Attribution International License (CC BY).

http://creativecommons.org/licenses/by/4.0/

Received 16 December 2015; accepted 14 June 2016; published 17 June 2016

ABSTRACT

The plant cell wall is a complex 3D network composed of polysaccharides, lignin and proteins. The knowledge of the structure and content of each cell wall polymer is a prerequisite to understand their functions during plant development and adaptation but also to optimise their industrial applications. The analysis of cell wall compounds is complicated by their multiple molecular interactions. In this review, we present numerous methods to purify, characterise and quantify proteins, polysaccharides and lignin from the wall. Two kinds of approaches are detailed: the first presents in vitro methods which involve the breakdown of the molecular linkages between polymers thanking to chemical, physical and/or enzymatic treatments. The second approach describes in situ methods that allow the cell wall polymer characterisation thanking to many analytical techniques coupled with microscopy. If microscopy is the common point of all of them, their development is associated with improvement of analytical techniques, increasing their power of resolution.

Keywords:

Polysaccharides, Proteins, Lignin, Purification, Spectroscopy, Chromatography, Immunology, Microscopy, Plant Cell Wall

1. Introduction

Plants represent an important form of life on Earth. The organic carbon they produce by the fixation of atmospheric CO2 during photosynthesis is the most abundant biological resource. Cellulosic biomass, a non-food carbon resource, can be found in agricultural and forestry products as well as in by-products (stalks, leaves, and husks, chips and sawdust from lumber mills), municipal waste (household garbage and paper products) and industrial waste (black liquor). Cellulosic matter is made up of acomplex three-dimensional network of cellulose/hemicelluloses embedded in a heterogeneous matrix of pectins and proteins. Secondary metabolites, lignin and mineral ions form the other components of the wall. Primary cell wall is found in all cells from algal to terrestrial plants. The tracheophytes clade exhibits, in some specialised cells such as fibres, vessels, tracheids and sclereids, an additional thicker secondary wall consisting of an interconnected network of cellulose and lignin [1] . As an exoskeleton, the extracellular primary or secondary walls give cells their shape. Moreover, the molecular compositions and the diversity of linkages within cell wall polymers define some new properties for plants such as mechanical and chemical resistance. As a consequence, the fine knowledge of its molecular composition is a prerequisite to understand how the polymers influence physicochemical and biological properties of the wall. However, the complexity of the plant cell walls makes their analysis difficult. Here we propose to review all of our knowledge on the analytical techniques, technologies or approaches developed for plant cell wall analysis.

2. Plant Cell Wall Composition

The most characteristic component found in all plant cell walls is cellulose accounting for 40% to 60% of the weight of the biomass. It consists of a collection of β-(1-4)-linked glucans (Figure 1) that interact through hydrogen bonds forming a semi-crystalline microfibril. In addition to cellulose, plant cell walls contain several other polysaccharides that can be grouped into hemicellulosic and pectic polymers. Depending on plants and tissues analysed, 20% to 40% of the carbon source are hemicelluloses. Hemicelluloses are a family of wall polysaccharides having β-(1-4)-linked backbones of glucose, mannose, or xylose termed as xyloglucans, xylans, mannans, galactomannans, glucomannans, galactoglucomanannans and β-(1-3,1-4)-glucans [2] [3] . Xyloglucans, which are found in primary cell walls of a wide number of higher plants, can be considered as the main hemicellulosic polymer (Figure 1). They have a cellulosic backbone to which α-D-xylose residues are linked at O-6

Figure 1. Schematic representation of some cell wall polysaccharides ( [423] [424] ).

and can be substituted at O-2 with β-galactose or α-L-arabinose. Other polymers complete the panel of the wall hemicellulosic polysaccharides. Xylans form a large group of polysaccharides with the common feature of a backbone of β-(1-4)-linked xylose residues (Figure 1). About 80% of the xylan backbone can be substituted with monomeric side-chains of arabinose or glucuronic acid linked to O-2 and/or O-3 of xylose residues, and also by oligomeric side chains containing arabinose, xylose, and sometimes galactose residues [3] . The arabinose substitution of xylan can be esterified by ferulic and diferulic acids [4] - [6] and can then be covalently linked to lignin monomers via an ether bond [7] [8] . Mannans and galactomannans (Figure 1) are mannose β-(1-4)-linked polysaccharides whereas glucomannans and galactoglucomannans are non-repeating glucose and mannose backbone polymers. Mixed β-glucans (MLG), essentially found in Poales [9] [10] are linear chains of repeat units of β-(1-4/3)-glucose linkages (Figure 1) giving plants specific motives for example G4G4G3G for Equisetum genius where G designs Glc and 3 or 4 the type of linkage [11] . Pectic polysaccharides are a covalently linked galacturonic acid family of polysaccharides including homogalacturonans, rhamnogalacturonans I (RGI) and rhamnogalacturonans II (RGII) [12] [13] . Homogalacturonans are unbranched homopolymer chains of a α-(1,4)-linked D-GalA making up more than 50% of the pectin (Figure 1). To form RGII, clusters of complex side chains are attached onto the O-2 or O-3 position in the galacturonan backbone (Figure 1). These side chains are composed of 12 different glycosyl residues (some of which are rare) linked together by at least 22 different glycosidic bonds. RGI is a branched polymer with a backbone of disaccharides (α-(1,4)-DGalA-α- (1,2)-L-Rha) repeats (Figure 1). The Rha residues in the backbone can be substituted with galactan, arabinan, and/or type II arabinogalactan side chains. Unlike RGII, the structure of the side chains of RGI can vary greatly amongst plants [14] .

Lignin, which makes up 10% to 24% of the weight of biomass, is an aromatic phenylproprionate consisting of covalent linkages of three aromatic alcohol monomers differing in their degree of methoxylation: hydroxycinnamyl, p-coumaryl, coniferyl, and sinapyl alcohols. When monolignols are incorporated into lignin polymers, they become respectively p-hydroxyphenyl H, guaiacyl G, and syringyl S phenylpropanoid units. These units are linked together during a lignification process, thanks to radical coupling reactions. Structure and amount of lignin depends on taxa, cell types and cell wall layers and can be influenced by developmental and environmental phenomena [15] . Lignin contains energy and can be burned to produce steam and electricity for the biomass-to- ethanol process.

Cell wall proteins are ubiquitously found from algal to land plants andform strongly different groups of plant cell proteinsboth by their amino acid content (unusually rich in one or two amino acids, contain highly repetitive sequence domains) and their level of N- or O-glycosylation. The most abundant and studied plant cell wall proteins are the hydroxyproline-rich glycoproteins (HRGPs) or extensins, the arabinogalactan proteins (AGPs), the glycine-rich proteins (GRPs), the proline-rich proteins (PRPs), and chimeric proteins that contain extensin-like domains. Abundance of these proteins in the walls differs depending on the cell type. Thus, each can be assumed to have functions specific to its particular cell type. Extensins are said to be structural cell wall proteins that may also play a significant role in development, wound healing, and plant defence. As extensins, PRPs are thought to be insoluble in the cell wall matrix. PRPs are implicated in various aspects of development (germination to pod formation, early stages of nodulation) and the expression of PRP genes is influenced by wounding, endogenous and fungal elicitors, ethylene, drought, and light. Contrary to the former proteins, AGPs are not covalently linked to the cell wall and therefore do not have a structural function [16] . The majority of the characterised AGPs contains between 1% and 10% (wt/wt) proteins and more than 90% (wt/wt) carbohydrate. Many studies implicate arabinogalactan proteins in several biological processes of cell proliferation and survival, pattern formation and growth, and in plant microbe interaction [17] . GRPs are a class of structural wall protein which seems to play important roles in the development of vascular tissues, nodules, and flowers and during wound healing and freezing tolerance [16] .

3. Plant Cell Wall Component Extraction

In order to analyse ex-situ cell wall components, their dissociation/destructuration/disorganisation from each other is a prerequisite for their analysis. Linkage breakdown between polymers can be done by chemical, enzymatic, physical or by a mixture of two or three of these proceedings. The challenge is then to release all the cell wall components without any change in their chemical composition. The plant cell wall disorganisation greatly depends on the number and type of linkages (covalent, hydrogen, ionic, electrostatic linkage) between all of them. With some few exceptions, polymer isolation, or more precisely polymer enrichment in extracted fractions is usually done by chemical and/or enzymatic methods, which purify specifically one family of cell wall macromolecules. Here, we describe the different approaches used by a large number of research groups for partial purification of plant cell wall polymers, as some new methods using for example ionic liquid as cell wall solvent. For full access to proteome and glycome, sequential extraction proceedings have been developed. These approaches combine the methods described in the following pages and are used to collect enriched fractions of proteins weakly and strongly bound to the matrix, or enriched fractions of pectin and hemicelluloses as described by Vanzin et al. [18] .

3.1. Protein Purification

Two groups of cell wall proteins (CWP) can be identified, the first is proteins formed only with amino acids and the second is glycoproteins, which have in addition, a non-amino acid component, usually a glycosidic fraction (2 to 15 carbohydrates) covalently linked to a hydroxyl and/or an amid function of some particular amino acid [19] - [22] . Arabidopsis thaliana genome analysis predicted that approximately 5000 genes encode proteins and/or glycoproteins targeted at the secretory pathway such as the wall compartment. It is probable that not all of them are CWP. Their amount, location and chemical environment require, for their isolation, the use of particular extractive buffers such as high ionic extractive salt buffers [23] - [25] .

Depending on their linkage strength with the wall, CWP can be divided in labile proteins versus weakly bound proteins. Labile protein extraction can be done directly from liquid media of plants or cell cultures [26] [27] , or by vacuum infiltration [28] . The extraction of weakly bound proteins (Van der Waals interactions, hydrogen bonds, hydrophobic or ionic interactions) can be done by salt solutions (CaCl2, LiCl, NaCl) without detergent. After protein extraction, their precipitation is usually done with trichloroacetic acid (TCA), acetone, chloroform/methanol, ammonium sulphate or combinations of them [29] - [31] . If cell wall glycoproteins are preferred as the whole proteome isolation, they can also be isolated as one fraction of CWP or directly by lectin conjugated resin such as ConA [32] - [35] , with specific resin, e.g. boronic acid-functionalised beads [36] , by capture of hydrazide-based glycoprotein [37] and by β-Glc-Yariv precipitation [38] [39] . Enzymatic and carbohydrate modification allows highly efficient purification of glycoproteins [40] . As for proteins, glycoproteins capture can be performed by different strategies which can be adapted to plant material used to improve the coverage of the glycoproteome, as well as the quantification of glycopeptides [41] . CWP can then be stored at −80˚C before analysis.

3.2. Polysaccharide Purification

Three approaches can be carried out for plant cell wall polysaccharide purification. The first consists of a partial purification of one family of polysaccharides such as pectins, cellulose or hemicelluloses without paying any attention to the other cell wall components. These methods have been developed from a large panel of plants known as high producers of one of the cell wall polysaccharides. The second approach consists of a sequential extraction procedure suitable to collect numerous fractions enriched in one compound. The last uses physical or chemical processes suitable to solubilise cell wall components.

3.2.1. Cellulose Isolation

In literature, cellulose purification is usually obtained by elimination of other cell wall components. The dissolution efficiency of cellulose depends greatly on the ability of solvents to disrupt its interchain hydrogen bonds. Some powerful solvents can dissolve cellulose such as thiourea/H2O, NaOH/urea, NaOH/CS2, alkali/H2O, LiOH/urea/H2O, molten salt like LiSCN∙2H2O and LiClO4∙3H2O, N-methylmorpholine-N-oxide (NMMO), dimethyl sulfoxide (DMSO)/tetrabutyl ammonium fluoride (TBAF), LiCl/1,3-dimethyl-2-imidazolidinone (DMI), LiCl/N-methyl-2-pyrrolidine (NMP) and LiCl/N,N-dimethylacetamide (DMAc) [42] [43] . More recently, ionic liquids such as 1-butyl-3-methylimidazolium acetate ([Bmim]Ac) [43] have been used to dissolve cellulose [44] - [49] . If cellulose is the major component of the cell wall, its isolation/purification requires a preliminary treatment, for example with trifluoroacetic acid (TFA), to hydrolyse non-cellulosic polymers [50] .

3.2.2. Pectins Isolation

Two groups of pectins can be found in the plant cell wall: low methylesterified pectins which are weakly bonded to the matrix [51] and high-methylesterified pectins which can be termed labile pectins. Methylesterified pectins can be easily extracted by cold water [52] [53] , or hot water [54] - [58] . Large amount of pectins can also be isolated by a hot acidified water known as conventional acidic extraction [59] . Pectins can also be extracted from an alcohol-insoluble solid phase as described by Bertin et al. [60] . Weakly bounded pectins, particularly those present in the middle lamella, can form gels due to Ca2+ bridges between two carboxyl groups belonging to two chains in close contact with each other [61] . Thus, Calcium removal by chelating agent can be considered another efficient method for pectin release [62] . Various chelating agents have been used such as ethylene diamine tetra-acetic acid (EDTA) [62] , cyclohexanediamine tetra-acetic acid (CDTA) [63] [64] , ammonium oxalate [54] , sodium carbonate [55] [63] or sodium hexametaphosphate [52] [62] [65] . All methods presented so far, with the exception of CDTA method, undergo thermal degradation of pectin polymers [55] . As a pretreatment, microwave heating at ambient and moderate pressures, ultrasonication and a super-high frequency electromagnetic field may give increased extraction yield and pectin quality [66] - [77] . With these methods, the quality of pectins solubilised is improved due to the reduction of both time and temperature [78] . To have access to a more strongly binding pectin fraction, CDTA treatment can be coupled with sodium carbonate (0.05 M) treatment with or without sodium borohydride, known as a protectant for the carbohydrate’s β-elimination [79] . But hemicellulosic polymers usually contaminate this last pectic fraction.

Enzymatic approach for depectination can be used either to purify non-modified pectins orto obtain degraded pectins (oligosaccharides, deesterified homogalacturonans...), depending on the enzymes activity. Pectinolytic enzymes are classified according to the nature of the substrate (pectin, pectic acid, oligogalacturonate), the degradation mechanism (trans-elimination or hydrolysis) and the type of cleavage (endo and exo) [80] . These enzymes can be divided into two main groups: pectinesterases (PE) or pectin methylesterases (PME) and depolymerases (polygalacturonases and lyases). PE or PME areexpressed in both plant, fungal and bacterial taxa [81] . They catalyse the hydrolysis of methyl esters of highly methylesterified pectin-bonds, causing the release of methanol and the formation of polygalacturonic acid [81] . The depolymerases are hydrolases (polygalacturonases and lyases) that have endo or exo-galacturonases activities. The depolymerases may be subdivided depending on the substrate and the cleavage mechanism of the glycosidic bonds in four categories: polygalacturonase (PG), polymethylgalacturonases (PMG), polygalacturonate lyases (PGL) and polymethylgalacturonate lyases (PMGL). The PG and PMG respectively act on the pectin by a hydrolysis mechanism, while PGL and GPLG act respectively by β-elimination of pectin [82] . Most commercial preparations of pectinases are of fungal origin, Aspergillus niger being the most commonly used source for industrial production of pectinolytic enzymes [83] . Thus, using the combined action of endoPG (Aspergillus niger) and PME (Aspergillus aculeatus), Pauly et al. [84] conducted a partial depectination of pea cell walls. Following the action of an endoPG (Aspergillus aculeatus, enzymes purified, without galactanase, galactosidase, arabinanase, arabinofuranosidase, xylosidase and esterase activity) on highly purified potato pulp, Byg et al. [85] obtained undegraded RGI with good yields of extraction. A mix or a sequential extraction proceeding can be used to recover all or one fraction of cell wall pectin.

3.2.3. Hemicelluloses Isolation

Because hemicelluloses are a family of polymers strongly linked to lignin and cellulose, their extraction is usually done by high concentration of alkali or organic solvents. Alkaline solutions hydrolyse the ester bonds and hydrogen linkages between polysaccharides and non-polysaccharides. That is why, extraction of hemicelluloses is usually done with aqueous alkali solutions such as potassium [86] , sodium [87] , lithium [88] or calcium hydroxide [89] , and hydrogen peroxide [90] . But to minimise the reducing end degradation of hemicellulose, sodium borohydride can be added [91] . KOH has been demonstrated to be the most efficient base to isolate heteroxylans from corn bran [92] whereas NaOH is a better solvent for glucomannans [91] . (Galacto) glucomannan isolation can be facilitated by the addition of borates which form anionic products, having a higher solubility in alkali solutions. By using a gradient of concentration of alkali solution, different hemicellulosic polymers can be isolated [91] . Without prior delignification, hemicelluloses from H. ammodendron and E. angustifolia have been extracted with ethanol/H20 under acidic conditions and hydrogen peroxide under alkaline medium [93] . This treatment allows solubilisation of hemicelluloses and lignin, and authors found no significant degradation or oxidation of hemicelluloses after alkaline peroxide post-treatment. Therefore, they deem it possible that lignin has a protective effect on them. However, not all the hemicellulosic substances were solubilised. According to mentioned authors, the hemicelluloses which remain are probably tightly bound to the cellulose [93] . Delignification and depectination increase the yield of hemicelluloses extracted [55] . Few neutral solvents have been used, among them dimethylsulfoxide (DMSO) [94] . Hemicellulose-DMSO extraction has the advantage of keeping the acetylation of xylans that is saponified by extraction with alkali. This extraction method thus provides low-modified hemicellulose fractions, but has the disadvantage of having relatively low extraction yields (<50%) [91] . Other solvents can also be used to extract hemicelluloses for example water or ethanol. By using pressurised low-polarity water and pressurised aqueous ethanol, Buranov and Maza [95] managed to remove 90% and 80% of hemicelluloses, respectively, from flax shives.

As for pectin, the yield of hemicelluloses extracted with chemical methods can be improved by physical pretreatments such as ultrasound [96] , microwave irradiation [97] or steam pretreatment [98] .
Partial hydrolysis after enzymatic treatment represents another way for direct analysis of soluble hemicellulosic oligosaccharides by mass spectrometry [84] . Two kinds of enzymes can breakdown hemicelluloses: the exohydrolases that release terminal monosaccharide units from the reducing end and the endohydrolases that cleave glycosidic bonds at random or specific positions [99] . Moreover, for each hemicellulosic polymer, enzymes involved in the cleavage of main chains and lateral chains of polysaccharides are necessary [100] - [103] . Hemicellulosic oligosaccharides can also be extracted from the biomass by physical process alone such as stem explosion or microwave treatment [104] .

3.3. Lignin Purification

Two main kinds of method are used to isolate lignin: acidolyse methods [105] - [108] and enzymatic methods [106] [109] - [111] . Even if acidolysis methods are quite fast and allow high pure lignin to be extracted, they lead to structural modifications of these polymers [106] [112] . To limit these alterations, it is possible to decrease the acidity of dioxane solutions for example, although this usually results in a higher carbohydrate contamination [106] [109] . The enzymatic method consists of a biomass treatment with cellulolytic enzymes to hydrolyse polysaccharides and liberate lignin. This method leads to good purification yields and lignin undergoesonly minimal structural modifications [112] . However, isolated ligninis contaminated by carbohydrates (12%) [111] . A combination of these two methods, a cellulolytic enzyme treatment followed by a mild acidolysis, triggers high lignin yields with fewer impurities than the enzymatic process alone [106] [109] [110] [113] .

Using a sequential extraction protocol, Sun et al. [114] purify lignin and hemicelluloses at the same time with mild dioxane, acidic dioxane, DMSO and alkali treatments. They obtained good yields of the two polymers and were able to find important structural features of lignin and hemicelluloses from barley straw. Ionic liquids, such as [Bmim]Cl (1-Butyl-3-methylimidazolium chloride) [115] , [Amim]Cl (1-Alyl-3-methylimidazolium chloride) [116] , [Bmim]MeSO4 (1-Butyl-3-methylimidazolium methylsulfate) [117] , allow the dissolution of biomass and the regeneration of lignin, hemicelluloses and cellulose fractions without prior purification treatments (see “Cell wall destructuration” paragraph).

3.4. Cell Wall Destructuration

Another strategy for plant cell wall composition analysis is the destructuration of the biomass. Nevertheless, wood dissolution is virtually impossible because of high levels of organisation of the cell wall polymers [118] . Chemical and physical proceedings have been developed to breakdown polymer linkages producing full cell wall disorganisation. Free polymers can then be solubilised before their use or analysis.

3.4.1. Chemical Dissociation

The majority of known solvents have been developed for the solubilisation of cellulose, but present little effect on lignocellulosic biomass [119] - [124] . Biofuel development has highlighted ionic liquids as revolutionary solvents for biomass disorganisation [44] . Ionic liquids (ILs), considered as “green solvents”, are salt solutions with a melting point below 100˚C [125] [126] . The interest of ILs is based on their outstanding abilities to dissolve monomeric or polymeric compounds, polar or non-polar organic or inorganic compounds [127] . It has thus been shown that they can dissolve some polysaccharides such as chitin [128] or cellulose [44] . At the same time, Vesa and Reijo [129] were the first to demonstrate that it was possible to dissolve more complex matrices such as wood completely in an IL (1-butyl-3-methyl imidazolium chloride [BmimCl]). The following work has sought to improve the biomass dissolution [130] and to understand the impact of cation and anion on the ability of IL to dissolve large amounts and diversity of lignocellulosic materials [46] [115] [131] - [133] (Figure 2).

Figure 2. Douglas fir wood dissolution in presence of 3 different ionic liquids under constant heating and microwave activation (Plazanet and Costa, not published).

Wood destructuration by [EmimOAc] can be monitored by laser confocal fluorescence as demonstrated for switch grass [134] . Because of the numbers of polymeric linkages between cell wall polysaccharides, different combinations of ILs can be developed. Muhammad et al. [118] suggests that ILs having high hydrogen bond basicities ß are suitable solvents for wood biomass dissolution. One limitation of ILs efficiency is their hygroscopicity requiring the use of dry material [135] . A new solvent, tetra-n-butylphosphonium hydroxide ([P4,4,4,4]OH) was suitable to dissolve cedar wood in aqueous solution at 60˚C while being stirred gently for 24 hours. However, in this method the solution induced a partial lignin degradation [135] .

3.4.2. Physical Dissociation

Physical processes including comminution (mechanical reduction in biomass particulate size), steam explosion (heating by high-pressure) and hydrothermolysis (solvolysis by hot compressed liquid water) have been applied for biomass disruption. But all these techniques strongly destroy the biomass, and they cannot be used for cell wall analysis. These methods, considered as pretreatment are able to break the lignin seal and disrupt the crystalline structure of cellulose [136] . Biomass can also be disrupted by ultrasonic (UAE) and microwave-assisted extraction (MAE) [137] [138] . These two techniques are suitable to extract large amounts of polysaccharides in shorter time, with less solvent than Soxhlet extraction [139] . But parameter control and temperature elevation during the extraction procedure are the two main limits for a large application of UAE and MAE. Some combinational methods have then been tested such as ultrasonic-microwave synergistic extraction (UMSE) [140] and enzymolysis-ultrasonic assisted extraction [141] . They have been developed to increase purity and yield ofextracted polysaccharides [142] .

4. In Vitro Plant Cell Wall Component Analysis

Analytical proceedings are largely dependent on the chemical composition and the amount of the bio-molecules to be characterised.

4.1. UV/Visible Spectrometry Methods

4.1.1. Protein Quantification

Several colorimetric methods exist to determine protein concentration in solution: the Biuret [143] , the Lowry method [144] , the bicinchoninic acid (BCA) assay [145] , the Coomassie Blue G-250 dye-binding assay [146] , the colloidal gold protein assay [147] , two dye-binding protein assay using Pyrogallol Red [148] or Pyrocatechol [149] , and the Pierce 660 nm assay [150] .

4.1.2. Polysaccharide Quantification

Polysaccharides are converted to furfural or a derivative of furfural by strong acidic solutions. Subsequent, complexion of the furfural or its derivative with an appropriate organic developer provides the formation of a chromophore, the absorbance of which, in visible light, allows the quantification of monosaccharides. Organic developers used to quantify total, neutral monosaccharides or uronic acids are mainly indole [151] , orcinol [152] , carbazole [153] [154] , resorcinol [155] , m-hydroxydiphenyl (m-HDP) [156] or phenol [157] . The phenol-sulphuric acid method is said to be the easiest and most reliable method to quantify neutral sugar because of its sensitivity and simplicity [158] . The m-hydroxydiphenyl (m-HDP) method is the most efficient to quantify uronic acids [156] . Due to the interference of uronic acids in the determination of neutral monosaccharides and vice versa, a correction patch should be applied as described by Montreuil and Spik [159] . Some of the methods described above have been miniaturised to gain greater sensitivity and to reduce reagents, biomass and time. Among them, we quote a micro-scale phenol-sulphuric assay [158] and a micro-scale m-HDP assay [160] .

4.1.3. Lignin Quantification

It is also possible to quantify lignin and some secondary metabolite content thanks to the use of UV spectrophotometry techniques. The most common methods to quantify lignin are the acetyl bromide, thioglycolic acid, and Klason. The first two methods are based on solubilisation of lignin and determination of absorbance values at 280 nm whereas the Klason mathod is a gravimetric assay. In the acetyl bromide method, lignin is first solubilised in acetyl bromide before their quantification at 280 nm [161] - [164] . The thioglycolic acid method is based on the formation of thioethers of benzyl alcohol groups found in the lignin, resulting in solubilisation of this polymer under alkaline conditions. The acetyl bromide method is faster, simpler and presents better recovery of lignin than the other methods [165] . Another simple method to quantify lignin from lignocellulosic biomass has been developed by Kline et al. [166] . After the dissolution of the whole biomass in an ionic liquid ([bmim]Cl), absorbance of the solution is read at 440 nm via UV-VIS spectrophotometry. Lignin concentration is then calculated using an extinction coefficient from standard lignin (isolated from biomass bythe Organoslov process) with Beer-Lambert Law. Coupled with multivariate analysis, fluorescence spectroscopy was used to develop models to study different parameters of poplars and Northern red oak such as total lignin, extractive and estimation of total holocellulose content [167] [168] .

4.2. Raman and Infrared Spectroscopies

Raman and infrared (IR) spectroscopes are vibrational spectroscopy techniques used to identify complex plant molecules [169] . The Raman spectrometry induces a change in the polarisability of the molecule whereas the IR transition exhibited a change in the dipole moment of polar components. These two non-destructive techniques are complementary: Raman spectroscopy is best at vibration of non-polar groups whereas IR spectroscopy is best at asymmetric vibration of polar groups. Vibrations which are seen in both spectra serve as evidence in the case of complex samples [170] [171] . Each polymer can be characterised by vibrational transitions of known chemical groups and a vibrational fingerprint of the molecule [170] [171] . A chemical function of a polymer is characterised by one or several wavenumber(s). For cellulose, these vibrational spectrometries allowed the identification of the different types of cellulose [172] - [175] , the determination of its crystallinity [175] - [178] and the chemical groups orientation inside the main chain [179] - [181] . Hemicelluloses [182] - [186] and pectic polysaccharides [187] - [193] have also been structurally characterised from several plant species. Synytsya et al. [193] showed FT-IR spectroscopy was more adapted to study functional groups of pectin (hydroxyls, carboxyls...), whereas FT-Raman spectroscopy was better to analyse the structure of the main chain (i.e. glycosidic bounds). Thus, FT-IR spectroscopy is an interesting tool to determine the degree of methylesterification of pectin [190] [191] . Kacurakova et al. [194] studied model plant cell wall polysaccharides by FT-IR spectroscopy in order to establish model data allowing identification of pectic polysaccharides and hemicelluloses [194] [195] . Raman and IR spectroscopes are also suitable to characterise lignin [196] . Total lignin content can be estimated by FTIR [197] , NIR [198] , UV-Raman [199] and FT-Raman [200] . FT-Raman [201] [202] , UV-Raman [203] [204] and FTIR [205] have been used to determine S/G/H content and to analyse the lignin structure [203] [204] [206] [207] .

4.3. Mass Spectrometry

The mass spectrometry (MS) has often been used instead of conventional detectors coupled to plant cell wall polymer separation. But MS can also be used directly on purified polymers.

4.3.1. Protein Fingerprint

Mass analysis is usually done after electrophoretic or liquid/nano-liquid chromatographic peptide separation. Matrix Assisted Laser Desorption/Ionisation Time Of Flight (MALDI-TOF-MS) and Electrospray Ionisation Time Of Flight (ESI-TOF-MS), that differ by their method of ionising analysts, are the two most conventional mass spectrometry technologies referenced in literature [208] - [212] . MALDI-TOF-MS is mainly done for peptide identification purified in 2D-gel electrophoresis before trypic in-gel digestion [213] . Electrospray, usually coupled with nano-Liquid Chromatography (LC), is suitable for the analysis of large numbers of proteins especially for membrane associated proteins [214] [215] . In some cases, Sodium Dodecyl Sulfate-PolyAcrylamide Gel Electrophoresis (SDS-PAGE) and LC can be combined for proteome analysis on an approach named GeLC-MS [215] . But until now, this approach has not been carried out for CWP. De novo protein sequencing from unknown or poorly known genomes and analysis of peptides containing non-proteinic or modified amino acids have also been developed using peptide fragmentation. Edman techniques (cleavage of the amino-terminal labelled residue) for protein sequence determination have been replaced by MS/MS proceeding: LC-ESI-MS/ MS and more recently MALDI-TOF-MS/MS [216] . Peptide purity for fragmentation is a prerequisite for high quality sequence and explains why LC-MS/MS is preferred to gel-MS/MS. Electron capture dissociation (ECD) and Electron Transfer Dissociation (ETD) are promising new fragmentation methods [217] applied for the identification of AGP31, an A. thaliana cell wall O-glycoprotein. Mass spectrometry can be used for proteomic quantification. Two methods have been developed: label-free and stable isotopes labelling methods [215] . Isobaric tags for relative and absolute quantification (iTRAQ) and isotope-coded protein labelling (ICPL) [218] are applied to plant proteomic [219] [220] after incorporation of stable modified amino acid to the culture media (SILAC) or under 15N isotope flux [221] - [223] .

4.3.2. Polysaccharide Fingerprint

Different types of MS have been used for structural cell wall carbohydrates determination such as MALDI-TOF, ESI [224] - [226] , Ion Trap Mass Spectrometry (ITMS) coupled with either MALDI or ESI [227] and Fourier Transform Ion Cyclotron Resonance Mass Spectrometry (FTICR-MS) [228] . In Oligosaccharide Mass Profiling (OLIMP), oligosaccharides are released by specific glycosylhydrolases. This method combines the advantages of enzymes specificity and the high sensitivity of mass spectroscopy [229] - [231] . Coupled with micro-dissection, the technique is suitable to analyse pectin and xyloglucan structures in one cell type, such as the outer or entire epidermis cell layer, palisade mesophyll cells, and vascular bundles [231] .

4.3.3. Lignin Fingerprint

MS can be used to analyse lignin structure from lignin degradation products or from whole lignin macromolecules. It is possible to degrade lignin by thermal or chemical treatments. For the first, rapid pyrolysis (py) techniques are the most commonly used to produce molecular fragments that can be analysed by MS or GC (Gas chromatography)-MS [232] . Py-GC-MS was widely used to characterise S/G/H ratio of lignin [233] - [235] . This technique needs small amounts of sample and the sample does not require pre-treatment except grinding. For the lignin study, several ionisation modes have been used with in-source py-MS [232] . The chemical methods to degrade lignin include alkaline nitrobenzene oxidation [236] - [238] and cupric oxide [239] which combined with GC-MS give information on the minimal quantities of uncondensed phenylpropanoide units and their relative amount; and the permanganate oxidation [240] , the hydrogenolyse [241] , the thioacidolysis [242] , the acidolysis [243] , the ozonolyse [244] and the Derivatisation Followed by Reductive Cleavage (DFRC) method [245] . The analysis by GC-MS of the degradation products obtained using the latter methods provides structural characterisation of lignin. The thermochemolysis with Tetramethylammonium Hydroxide (TMAH) combined with GC- MS is a rapid method for the characterisation of the S/G/H ratio in plant material [246] . Structural characterisation and molecular weight distribution of lignin can be provided from non-degraded macromolecules by means of soft-ionisation mass spectrometric techniques (mainly MALDI-MS and ESI-MS) [247] - [251] .

4.4. Nuclear Magnetic Resonance (NMR) Spectroscopy

Even if NMR methods are less rapid and not as sensitive as others spectroscopic methods, they can provide useful structural information on the cell wall polymers [252] .

4.4.1. Polysaccharide Fingerprint

NMR methods are too insensitive to use at the cellular level and must be applied from a sample resulting from a homogenising of several cell types [252] . NMR spectroscopy has been used to provide information on the chemical structure of plant cell wall polysaccharides and their interaction [252] - [260] . CP-MAS (Cross Polarisation Magic Angle Spinning) 13C-NMR has been largely used to characterise cellulose crystallinity [261] - [264] . Structural polysaccharide information by NMR can also be available from the dissolution of the whole cell wall without the need to isolate fractions [252] [265] - [273] .

4.4.2. Lignin Fingerprint

The molecular structure of lignin has been widely studied by NMR [274] . NMR techniques can be suitable as a qualitative and quantitative method forcharacterisation of lignin isolated or in the entire dissolved plant cell wall [275] . NMR technique has been used to quantify the S/G/S ratio in different biomass samples [276] [277] , estimate the total lignin content [278] and analyse the lignin structure [279] .

4.5. Electrophoretic Methods

4.5.1. Protein Fingerprints

The size of the putative cell wall proteome and the diversity of CWPs require the use of a combination of separative and analytical methods without any certainty as to the recovery of the whole wall proteome. The CWP analysis is classically done by 1D and 2D-PAGE electrophoresis followed by mass spectroscopy identification. The mass of tryptic digested peptides, given by MALDI-TOF MS or MALDI-TOF/TOF [280] are then mapped against databases prior to protein identification. The high amounts of acidic and basic CWPs like the occurrence of heavily glycosylated CWPs require some particular electrophoretic or chromatographic methods. The separation of basic proteins on 2D-PAGE can be achieved by blocking the cysteine oxidation during protein separation. Cysteine alkylation with iodoacetamide, acrylamide [281] [282] , maleimide [283] or by disulphide exchange with dithiodiethanol [284] greatly increases the protein resolution in 2D-PAGE.

4.5.2. Polysaccharide Fingerprints

Carbohydrate gel electrophoresis method (PACE) with derivatisated reducing end sugar by fluorophore, such as 8-amino-naphtalene-1,3,6-trisulfonic acid (ANTS), has been developed for N-glycan [285] - [287] and cell wall polysaccharide analysis [288] [289] . The fluorophore-assisted carbohydrate electrophoresis (FACE) uses a combination of chemical and specific hydrolases which are able to release monosaccharides before their electrophoretic separation [290] [291] . Derivatisated glycans can also be analysed by capillary electrophoresis (CE) [292] . Unknown glycan migration can then be compared with standard if they are available. Mass spectroscopy (MS) coupled with CE (CE/MS) is no longer dependent on the standards used to identify simultaneously, the oligosaccharide and its molecular design [293] - [296] .

4.5.3. Lignin Fingerprint

A method coupling capillary electrophoresis (CE) with on-column UV-detection has been developed to quantitatively analyse phenolic lignin degradation products [297] . Isotachophoretic separation is another electrophoretic method used to separate biomass hydrolysis products such as phenol compounds [298] .

4.6. Chromatographic Methods

Chromatographic methods, such as liquid and gaz chromatography, are the most conventional analytical processes developed for glycoprotein and glycan analysis. These separative methods can be coupled with a large panel of detectors: UV/visible, fluorescence, Pulsed Amperometric Detection (PAD) and/or mass spectroscopy.

4.6.1. Protein Fingerprints

Because glycoproteins are poorly separated by 2D-PAGE, LC coupled with tandem MS/MS are preferred. Tryptic digestion is coupled with a peptide N-glycosidase from sweet almonds (PNGase A) for N-glycans release before MS analysis [299] . For O-glycoproteins such as cell wall AGP previously purified by β-Glc-Yariv, their separation can be performed by 2D High Pressure Size Exclusion Chromatography (HPSEC) coupled with a double fluorescence and 215 nm UV detection [300] . Collected AGP fractions can be then deglycosylated with hydrogen fluoride or trifluoromethanesulfonic acid [301] followed by tryptic digestion before MS protein analysis.

4.6.2. Polysaccharide Fingerprints

Previously solubilised polysaccharides can be analysed from partial purification and enrichment fractions or directly after biomass disorganisation by chemical and or physical techniques. After their hydrolysis in monosaccharides, High Performance Liquid Chromatography (HPLC), GC or GC-MS can be used to determine the polysaccharide composition. LC-MS is also suitable to analyse oligosaccharides produced after a partial digestion of polysaccharides. The most common approach consists of the analysis of the monosaccharides composition of cell wall polysaccharides after they have hydrolysed or methanolysed [302] - [304] . For proteoglycans, a preliminary chemical deglycosylation-step can be done before analysis (see paragraph on Protein fingerprints). The monosaccharides are released from poly- or oligosaccharides with strong acid solutions such as trifluoricacetic acid usually coupled with high temperature. Such processes have been demonstrated to be efficient for cellulose depolymerisation, but may also lead to degradation of some hemicellulosic polysaccharides, such as xylose [305] . The methanolysis procedure is preferred because of its low level of monosaccharidedegradation [306] . Methanolysis releases stable methyglycosides and methylglycuronosides which can be derivatisated by pertrimethylsililylation [307] acetylation [308] , trifluoroacetylation [309] , reduction/acetylation [310] oxilmation/acetylation [311] and O-methyloximation/acetylation [312] . Volatile compounds can then be separated by GC or GC-MS. Underivatisated monosaccharides can also be separated by HPLC, but the sensitivity and the efficiency of the separation is lower than with the GC technique.

In some case, oligosaccharides and deglycosylated N-glycans can be directly identified by High-Performance Anion-Exchange Pulsed-Amperometric Detection (HPAE-PAD) and LC-MS [313] - [315] . Thin layer chromatography (TLC) is also a technique used to rapidly separate and estimate the content of mono- or oligosaccharide mixtures [316] .

4.6.3. Lignin Fingerprint

GC has been used to determine the S/G/H ratio of lignin after their oxidation by cupric oxide [296] [297] [317] [318] , permanganate [297] or alkaline nitrobenzene oxidation [237] [238] [319] . Coupled with MS, GC was applied to study lignin monomers obtained after thioacidolysis [242] or acidolysis [243] . It is also possible to analyse acidolysis products without derivatisation with HPLC, but with a lower resolution [243] . High-performance size exclusion liquid chromatography coupled with different detectors (multi-angle laser light scattering (MALLS), quasi-elastic light scattering (QELS), interferometry refractometry (RI) and UV detection) can be used to determine the molecular weight distribution of purified lignin [320] .

4.7. Immunological and Immunological-Like Methods

Immunological techniques are used for polysaccharides, glycoproteins and protein identification and quantification. Generally monoclonal antibodies (mAbs) have been developed from neo-glycoprotein immunisation procedures or from retrospective characterisation after immunisation with cell-wall material [321] [322] . A lot of mAbs recognise epitopes of RGI [323] - [329] , homogalacturonan [330] [331] , xyloglucan [332] - [334] , xylogalacturonan [335] , xylan [336] , even epitopes of wall proteins such as AGP [337] - [340] and extensins [341] . This collection of mAbs has been completed by Pattathil et al. [342] with the creation of 130 new glycan-directed mAbs. Cell-wall-directed mAbs available are inventoried in CarboSource (http://www.ccrc.uga.edu/~carbosource/CSS_home.html), PlantProbes (http://www.plantprobes.net), Biosupplies Australia (http://www.biosupplies.com.au) and the Wall-BioNet (http://glycomics.ccrc.uga.edu/wall2/antibodies/antibodyHome.html). Once the mAbs recognising the epitope of the polymer of interest is bound, the surplus of mAb is washed, and a secondary antibody hybridised on the first one. The secondary antibody can be tagged with an enzyme, a radioisotope, a fluorochrome or a paramagnetic bead [343] .

Carbohydrate-Binding Modules (CBMs) are discrete protein modules present in microbial or plant carbohydrolases (and few from non-hydrolytic proteins) and have a wide range of binding specificities toward cell wall carbohydrates [344] - [346] . Coupled directly with fluorophore (FITC) or with His-tag, CBMs can recognise cellulose [335] - [339] [347] , xylans [348] - [350] , and mannans [351] [352] . McCartney et al. [353] used CBMs containing His-tags in a three-stage procedure, with an anti-His antibody produced in mice (secondary stage) recognised by an anti-mouse FITC (tertiary stage). Because they deemed the available anti-His FITC antibodies less effective, they decided against a two-step procedure. Compared to antibodies, CBMs have several advantages. Firstly, because of their high specificity and small size compared to antibodies, CBMs can access their targetmore easily [351] . Secondly, gene or protein sequences of CBMs can be obtained, and the engineering of CBM specificities is possible once the protein structure is known [354] . Extracted carbohydrates can be spotted on several supports like membranes (Dot-blots; [355] ), silica plates [356] or in different ELISA (Enzyme- Linked Immunosorbent Assay) plates [342] [357] before their analysis. However, a limit of these techniques is the efficiency of hybridisation between mAb or CBM with their target due to masked epitopes by other cell wall polymers or substituents, or by a polymer aggregation [311] [330] [333] . Moller et al. [358] have developed a technique named CoMPP for Comprehensive Microarray Polymer Profiling, in which glycans are sequentially extracted from plant tissues to generate microarrays. Microarrays are then probed with mAbs or CBMs specific to cell wall polymers. This technique allows them to compare Arabidopsis mutants providing a global snapshot of their cell wall composition. Immunological and immunolical-like methods can also be carried out on plant biomass dissolved in ionic liquid to rapidly obtain its main composition [359] .

5. In Situ Plant Cell Wall Components Analysis

As prospective approaches, plant cell wall components can be directly identified and quantified inside the cell wall by specialised microscopic techniques. Specific dye reagents, protein-polymer recognition (antibodies, lectin, etc...), fluorescent and vibrational excitation and more recently mass spectrometry are susceptible to be used at a resolution level of μm or lower. For all these approaches, sample preparation, microscopic resolution and cartography reconstitution are the principal limitations for their large scale use. Probably one of the strongest difficulties is the sample preparation (quality and fitness of the section, flatness of the sample, component accessibility formeasuring).

5.1. Staining Microscopy

Tissue sections from 10 to 50 μm canusually be stained, in 10 min, by toluidine blue O [360] (Figure 3(c)) as a basic for cell labelling. To have an idea about the relative proportion of cellulose and lignin, wood sections can be sustained by the simultaneous action of safranin and alcian blue according to Tolivia and Tolivia [361] (Figure 3(b)). For finer knowledge of lignin composition and distribution in secondary cell walls, some specific dyes such as phloroglucinol-acid [362] (Figure 3(c)) and Maüler eagent [363] can be applied (Figure 3(d)).

5.2. Fluorescent Microscopy

Fluorescent and confocal fluorescent microscopes are usually used for the determination of the spatial distribution of target compounds by a large panel of chromophores. Safranine, known as a suitable lignin stain, produces a green/yellow fluorescence in the secondary cell wall and a red/orange fluorescence in the middle lamella (ML) region [364] . Other fluorescent stains generally used to localise lignin (even if there are non-specific) are

Figure 3. Transversal section of in vitro poplar wood. (a) Poplar wood organisation; (b) FASGA staining; (c) Toluidine staining; (d) Phloroglucinol staining; (e) Maüle staining (Costa, not published).

acriflavin [365] , basic fuchsin [366] and berberine sulphate [367] . Contrary to lignin, cellulose is not an intrinsically fluorescent molecule. However, even after the removal of lignin, fluorescence can still be observed in different cellulose samples [368] , probably because of proteins and residual lignin [369] . Fluorescent markers, such as 7GFE, calcofluor, S4B have been tested [370] [371] for their potential to allow imaging of cellulose distribution in the wall of Arabidopsis seedlings. Hoch et al. [372] have used pontamine Fast Scarlet 4B (S4B) to determine the cellulose distribution.

Wall polysaccharides can also be studied thanks to glycan-directed probes such as antibodies, carbohydrate-binding molecules (CBM) and lectins coupled with fluorophore (GFP, CFP, YFP, FICT) (Figure 4(a)). These techniques allow us to study the cell wall microstructure and polymer localisation in situ within complex plant tissues [373] - [377] . Multiple fluorophore dyes can also be used for simultaneous multiple compound identification. This approach is limited by the number of chromophores available and by the autofluorescence of some natural plant cell wall components. For fluorescence, mAbs, lectins and or CBMs are the same as those described earlier (compare the paragraph on Immunological and immunological-like methods).

5.3. Multi-Photonic Microscopy

Multiphoton laser scanning microscopy is another approach for three-dimensional fluorescence imaging. Two-photon and multi-photon excitation reduce light scattering, autofluorescence, photo bleaching and photo damage of living cells better than wide-field fluorescence or confocal microscopy [378] - [382] . Two-photon excitation in the NIR spectral region (740 - 1200 nm) has been applied for root tip analysis [383] , embryos [384] ,

Figure 4. Transversal section of in vitro poplar wood. ((a), (b)) JIM8 immunolabelling observed by fluorescence (a) and by TEM (b); ((c), (d)) LM11 immunolabelling observed by fluorescence (c) and by TEM (d) (Costa, not published).

chloroplasts [385] and for intra-tissue nanodissection of cell walls [386] . A modified periodic acid schiff treatment using propidium iodide as cell wall stain gives a fluorophore suitable for monitoring plant cell wall evolution [387] . Monolignols, suchas ferulic acids that are a key molecular bridge between hemicellulose and lignin, are also two-photon excitable fluorophores [388] . However, until now, no application of multi-photon laser scanning microscopy has been published in literature.

5.4. Electronic Microscopy

Two electronic microscopes have usually been applied for plant cell wall component identification: Scanning Electron Microscopy (SEM) and Transmission Electron Microscopy (TEM). If SEM gives information about the cell wall surface, it can be coupled withelementary analysis for the identification of some particular incrustation inside the wall. For high-resolution SEM the sample must be made conductive, which is normally accomplished by coating biological specimens with vaporised metal or carbon. It is believed that these sample preparation procedures used for the plant materials can damage and ultimately change the native structure of the plant cell wall. With the recent introduction of high-pressure freezing and freeze substitution methods for preserving cells for infrastructural analysis, electron microscopy has become a valuable tool for correlative imaging [389] . However, there are still limitations with these imaging techniques and often a reduction of resolution is the resulting compromise. For in situ plant cell wall component identification, immunostaining of ultrathin sections of wood were incubated with a similar set of antibodies as those described previously (see paragraph on immunological and immunological-like methods). Sample preparation involves tissue embedding into a resin (LR- white for example). The secondary antibody is tagged with electron-opaque particles (Figure 4(b)).

5.5. Infrared and Raman Micro-Spectroscopies

IR and Raman micro-spectroscopy are suitable for analysis at the single cell wall level [390] [391] (Figure 5). FT-IR microspectroscopic imaging or FT-IR microscopy is able to locate plant cell wall polymers [392] - [394] . In some applications such as biorefinery, FT-IR microscopy is able to monitor the enzymatic hydrolysis of wood polymers [393] [395] and the biomass destructuration during steam treatment [396] . Hemicellulose and pectin images can be obtained and (acyl) ester modifications inside hemicellulosic components can be quantified as demonstrated by the work of Gou et al. [397] . Polarised FTIR microscopy can also give some data concerning the angle of carbohydrates as demonstrated by Chang et al. [398] during the deposition of G-layer in Populus tremula x P. alba, clone INRA 717-1B4. FT-IR microscopy is a non-destructive method to visualise the com-

Figure 5. Transvers al section of Douglas-fir softwood. (a) Infrared microscopy picture; (b) infrared spectra extracted from the infrared microscopy picture; (c) Raman microscopy picture obtained from the 1600 cm-1 absorbance band of the spectra ; (d) Raman spectra (Plazanet and Costa, not published).

ponent distribution, which has a resolving power of about one µm sufficient for structural and chemical in situ investigation especially for monitoring plant cell wall evolution [399] (Figure 5(a)).

Because some chemical functions do not have IR detectable vibrational wavelengths, Confocal Raman Microscopy (CRM) represents a complemenraty approach for non-destructive plant cell wall component analysis [400] . Arabidospsis cell walls [401] and spruce wood biomass [400] [402] have been visualised in situ with CRM. This technique can be applied to compare wild type and transgenic poplars [403] , and normal and compressed wood of pines [404] . As for FTIR microscopy, CRM is a valuable approach to assess the in situ effect of biomass modification such as wood degradation for bioraffinery [399] [405] [406] . Limits of this approach are the autofluorescence produced during the laser excitation, and the time necessary for image acquisition. New laser sources and bioinformatic progress are the key to an easier application of this non-destructive method [175] [407] . A considerable advantage of CRM is its resolution level to less than 10 nm (Figure 5(b)).

5.6. Mass Spectra Microscopy

Mass spectrometry imaging for plant cell wall analysis can be divided into two groups: TOF-SIMS and MALDI- MS. Secondary ion mass spectrometry (SIMS) exhibits a sub-µm resolution and has first been used for the imaging of elements such as Na+, K+, Ca2+ in plant cells [408] [409] . Cyro-TOF-SIMS and nano SIMS are two evolutions of SIMS techniques making analysis of high water content materials at lower than ~50 nm level possible [410] [411] . TOF-SIMS has also been used to analyse and conduct direct-mapping of organic compounds [412] . Recently, TOF-SIMS has been applied to investigate wood components, for example lignin and polysaccharides [413] [414] and for monolignol distribution on transversal stem sections of two geniuses of eucalyptus [415] . TOF-SIMS has also been used to investigate heartwood extractable components from Cryptomeria japonica such as ferruginol [416] [417] . Because polysaccharides have highly complex glucidic repeat units, MALDI is preferred to SIMS for their identification. Recently, MALDI-TOF MS imaging has been developed for cellulose and hemicellulose determination on poplar wood transversal section [418] .

5.7. NMR Spectroscopy

NMR spectroscopy is a non-invasive and non-destructive technique able to collect information in the native cell wall. Using in-vivo solid-state 13C NMR (CP-MAS), Jarvis and Apperley [419] identified cell wall polysaccharides in several seeds and showed their (im)mobility in the wall as well asthe molecular organisationof cellulose. However, the use of CP-MAS NMR leads toa major practical problem, because the application of centrifugal forces can lead to the disintegration of plant tissues. Nevertheless, this technique was used by Fenwick et al. [420] to compare the flexibility of polymers in celery collenchyma cell walls.

6. The Future

In vitro and in situ methods for plant cell wall polymer characterisation are two complementary approaches. The first approach can accurately describe finesse composition, structure and organisation of cell wall polymers in the 3D network. If polymer extraction has been a useful method applied in the past, some new solvents or some combinational of physical and chemical treatments can become preferable for cell wall polymer disorganisation. This new approach strongly reduces polymer modifications giving the physiologists the opportunity to analyse and understand the interaction between each of them. When polymers are in solution, the complex mixture can then be analysed by immune methods or mass spectrometry. For proteins, mass spectrometry and de novo sequencing by MS/MS are the two precedures applied for their identification. For polysaccharides, immune detection is now the fastest method of analysis. High-resolution level of mass spectrometry can represent an alternative to immune detection if polysaccharides are partially hydrolysed with a set of pure enzymes. As tryptic digestion, each polysaccharide of the cell wall can then be partially hydrolysed in oligosaccharides before being identified by MS.

The in situ approach probably gives lower quality of results but is suitable to screen a large plant cell wall population for industrial applications. The in situ methods are non-destructive and can be carried out after plant biopsy. Here, material preparation and resolution levels of some analytical methods are the limiting factors for their application. For example, tomography instrumentation has recently evolved to high-resolution electro tomography that is a 3D TEM with a resolution at the atomic level [421] . This microscopy has not been applied to the cell wall but to syncytial-type cell plate formation permitting the visualisation of vesicle arrangement at a resolution of ~6nm [422] . In the same order, SIMS are probably the future of in situ plant cell wall analysis, but for now the resolution power of the commercial instrumentation does not allow us to access information at the nm level.

Acknowledgements

The work presented here has been made under the chair of excellence Forest Resources and Wood Applications named SylvaLIM (http://www.fondation.unilim.fr/sylvalim) which was funded by the foundation of the University of Limoges, the region Aquitaine-Limousin-Poitou-Charente (ALPC) and the Commision of Europeen Communities.

Cite this paper

Guy Costa,Idelette Plazanet, (2016) Plant Cell Wall, a Challenge for Its Characterisation. Advances in Biological Chemistry,06,70-105. doi: 10.4236/abc.2016.63008

References

  1. 1. Mauseth, J.D. (1988) Plant Anatomy. Benjamin/Cummings Publ. Co., Menlo Park.

  2. 2. Ebringerova, A., Hromadkova, Z. and Heinze, T. (2005) Hemicellulose. Polysaccharides I. Springer, Berlin Heidelberg, 1-67.
    http://dx.doi.org/10.1007/b136816

  3. 3. Scheller, H.V. and Ulvskov, P. (2010) Hemicelluloses. Annual Review of Plant Biology, 61, 263-289.
    http://dx.doi.org/10.1146/annurev-arplant-042809-112315

  4. 4. Hartley, R.D. and Ford, C.W. (1989) Phenolic Constituents of Plant Cell Walls and Wall Biodegradability. Plant Cell Wall Polymers, Biogenesis and Biodegradation, 399, 137-145.
    http://dx.doi.org/10.1021/bk-1989-0399.ch009

  5. 5. de O. Buanafina, M.M. (2009) Feruloylation in Grasses: Current and Future Perspectives. Molecular Plant, 2, 861-872.
    http://dx.doi.org/10.1093/mp/ssp067

  6. 6. Ralph, J. and Helm, R.F. (1993) Lignin/Hydroxycinnamic Acid/Polysaccharide Complexes: Synthetic Models for Regiochemical Characterization. In: Jung, H.G., Buxton, D.R., Hatfield, R.D. and Ralph, J., Eds., Forage Cell Wall Structure and Digestibility, ASA-CSSA-SSSA, Madison, 201-246.

  7. 7. Scalbert, A., Monties, B., Lallemand, J.-Y., Guittet, E. and Rolando, C. (1985) Ether Linkage between Phenolic Acids and Lignin Fractions from Wheat Straw. Phytochemistry, 24, 1359-1362.
    http://dx.doi.org/10.1016/S0031-9422(00)81133-4

  8. 8. Kondo, T., Mizuno, K. and Kato, T. (1990) Cell Wall-Bound p-Coumaric and Ferulic Acids in Italian Ryegrass. Canadian Journal of Plant Science, 70, 495-499.
    http://dx.doi.org/10.4141/cjps90-058

  9. 9. Popper, Z.A. and Fry, S.C. (2004) Primary Cell Wall Composition of Pteridophytes and Spermatophytes. New Phytologist, 164, 165-174.
    http://dx.doi.org/10.1111/j.1469-8137.2004.01146.x

  10. 10. Trethewey, J.A., Campbell, L.M. and Harris, P.J. (2005) (1→ 3),(1→ 4)-β-d-Glucans in the Cell Walls of the Poales (Sensu Lato): An Immunogold Labeling Study Using a Monoclonal Antibody. American Journal of Botany, 92, 1660- 1674.
    http://dx.doi.org/10.3732/ajb.92.10.1660

  11. 11. Fry, S.C., Nesselrode, B.H., Miller, J.G. and Mewburn, B.R. (2008) Mixed-Linkage (1→ 3, 1→ 4)-β-d-Glucan Is a Major Hemicellulose of Equisetum (Horsetail) Cell Walls. New Phytologist, 179, 104-115.
    http://dx.doi.org/10.1111/j.1469-8137.2008.02435.x

  12. 12. Albersheim, P., Darvill, A.G., O’Neill, M.A., Schols, H.A. and Voragen, A.G.J. (1996) An Hypothesis: The Same Six Polysaccharides Are Components of the Primary Cell Walls of All Higher Plants. Pectins and Pectinases, 14, 47-53.
    http://dx.doi.org/10.1016/S0921-0423(96)80245-0

  13. 13. Mohnen, D. (2008) Pectin Structure and Biosynthesis. Current Opinion in Plant Biology, 11, 266-277.
    http://dx.doi.org/10.1016/j.pbi.2008.03.006

  14. 14. Harholt, J., Suttangkakul, A. and Vibe Scheller, H. (2010) Biosynthesis of Pectin. Plant Physiology, 153, 384-395.
    http://dx.doi.org/10.1104/pp.110.156588

  15. 15. Boerjan, W., Ralph, J. and Baucher, M. (2003) Lignin Biosynthesis. Annual Review of Plant Biology, 54, 519-546.
    http://dx.doi.org/10.1146/annurev.arplant.54.031902.134938

  16. 16. Cassab, G.I. (1998) Plant Cell Wall Proteins. Annual Review of Plant Biology, 49, 281-309.
    http://dx.doi.org/10.1146/annurev.arplant.49.1.281

  17. 17. Ellis, M., Egelund, J., Schultz, C.J. and Bacic, A. (2010) Arabinogalactan-Proteins: Key Regulators at the Cell Surface? Plant Physiology, 153, 403-419.
    http://dx.doi.org/10.1104/pp.110.156000

  18. 18. Vanzin, G.F., Madson, M., Carpita, N.C., Raikhel, N.V., Keegstra, K. and Reiter, W.-D. (2002) The Mur2 Mutant of Arabidopsis Thaliana Lacks Fucosylated Xyloglucan Because of a Lesion in Fucosyltransferase AtFUT1. Proceedings of the National Academy of Sciences of the United States of America, 99, 3340-3345.
    http://dx.doi.org/10.1073/pnas.052450699

  19. 19. Marshall, R.D., Neuberger, A., et al. (1970) Aspects of the Structure and Metabolism of Glyco-Proteins. Advances in Carbohydrate Chemistry and Biochemistry, 25, 407-478.
    http://dx.doi.org/10.1016/S0065-2318(08)60433-3

  20. 20. Faillard, H. and Schauer, R. (1972) Glycoproteins: Their Composition, Structure and Function. Elsevier, Amsterdam, 1246-1267.

  21. 21. Kornfeld, R. and Kornfeld, S. (1976) Comparative Aspects of Glycoprotein Structure. Annual Review of Biochemistry, 45, 217-238.
    http://dx.doi.org/10.1146/annurev.bi.45.070176.001245

  22. 22. Clarke, A.E., Anderson, R.L. and Stone, B.A. (1979) Form and Function of Arabinogalactans and Arabinogalactan- Proteins. Phytochemistry, 18, 521-540.
    http://dx.doi.org/10.1016/S0031-9422(00)84255-7

  23. 23. Feiz, L., Irshad, M., Pont-Lezica, R.F., Canut, H. and Jamet, E. (2006) Evaluation of Cell Wall Preparations for Proteomics: A New Procedure for Purifying Cell Walls from Arabidopsis Hypocotyls. Plant Methods, 2, 10.
    http://dx.doi.org/10.1186/1746-4811-2-10

  24. 24. Jamet, E., Canut, H., Boudart, G. and Pont-Lezica, R.F. (2006) Cell Wall Proteins: A New Insight through Proteomics. Trends in Plant Science, 11, 33-39.
    http://dx.doi.org/10.1016/j.tplants.2005.11.006

  25. 25. Irshad, M., Canut, H., Borderies, G., Pont-Lezica, R. and Jamet, E. (2008) A New Picture of Cell Wall Protein Dynamics in Elongating Cells of Arabidopsis Thaliana: Confirmed Actors and Newcomers. BMC Plant Biology, 8, 94.
    http://dx.doi.org/10.1186/1471-2229-8-94

  26. 26. Borderies, G., Jamet, E., Lafitte, C., Rossignol, M., Jauneau, A., Boudart, G., et al. (2003) Proteomics of Loosely Bound Cell Wall Proteins of Arabidopsis Thaliana Cell Suspension Cultures: A Critical Analysis. Electrophoresis, 24, 3421-3432.
    http://dx.doi.org/10.1002/elps.200305608

  27. 27. Charmont, S., Jamet, E., Pont-Lezica, R. and Canut, H. (2005) Proteomic Analysis of Secreted Proteins from Arabidopsis Thaliana Seedlings: Improved Recovery Following Removal of Phenolic Compounds. Phytochemistry, 66, 453-461.
    http://dx.doi.org/10.1016/j.phytochem.2004.12.013

  28. 28. Boudart, G., Jamet, E., Rossignol, M., Lafitte, C., Borderies, G., Jauneau, A., et al. (2005) Cell Wall Proteins in Apoplastic Fluids of Arabidopsis Thaliana Rosettes: Identification by Mass Spectrometry and Bioinformatics. Proteomics, 5, 212-221.
    http://dx.doi.org/10.1002/pmic.200400882

  29. 29. Jiang, L., He, L. and Fountoulakis, M. (2004) Comparison of Protein Precipitation Methods for Sample Preparation Prior to Proteomic Analysis. Journal of Chromatography A, 1023, 317-320.
    http://dx.doi.org/10.1016/j.chroma.2003.10.029

  30. 30. Visser, N.F.C., Lingeman, H. and Irth, H. (2005) Sample Preparation for Peptides and Proteins in Biological Matrices Prior to Liquid Chromatography and Capillary Zone Electrophoresis. Analytical and Bioanalytical Chemistry, 382, 535-558.
    http://dx.doi.org/10.1007/s00216-005-3120-9

  31. 31. Bodzon-Kulakowska, A., Bierczynska-Krzysik, A., Dylag, T., Drabik, A., Suder, P., Noga, M., et al. (2007) Methods for Samples Preparation in Proteomic Research. Journal of Chromatography B, 849, 1-31.
    http://dx.doi.org/10.1016/j.jchromb.2006.10.040

  32. 32. Bunkenborg, J., Pilch, B.J., Podtelejnikov, A.V. and Wisniewski, J.R. (2004) Screening for N-Glycosylated Proteins by Liquid Chromatography Mass Spectrometry. Proteomics, 4, 454-465.
    http://dx.doi.org/10.1002/pmic.200300556

  33. 33. Faye, L., Boulaflous, A., Benchabane, M., Gomord, V. and Michaud, D. (2005) Protein Modifications in the Plant Secretory Pathway: Current Status and Practical Implications in Molecular Pharming. Vaccine, 23, 1770-1778.
    http://dx.doi.org/10.1016/j.vaccine.2004.11.003

  34. 34. Wang, Y., Wu, S. and Hancock, W.S. (2006) Approaches to the Study of N-Linked Glycoproteins in Human Plasma Using Lectin Affinity Chromatography and Nano-HPLC Coupled to Electrospray Linear Ion Trap—Fourier Transform Mass Spectrometry. Glycobiology, 16, 514-523.
    http://dx.doi.org/10.1093/glycob/cwj091

  35. 35. Minic, Z., Jamet, E., Négroni, L., Der Garabedian, P.A., Zivy, M. and Jouanin, L. (2007) A Sub-Proteome of Ara- bidopsis Thaliana Mature Stems Trapped on Concanavalin A Is Enriched in Cell Wall Glycoside Hydrolases. Journal of Experimental Botany, 58, 2503-25012.
    http://dx.doi.org/10.1093/jxb/erm082

  36. 36. Sparbier, K., Koch, S., Kessler, I., Wenzel, T. and Kostrzewa, M. (2005) Selective Isolation of Glycoproteins and Glycopeptides for MALDI-TOF MS Detection Supported by Magnetic Particles. Journal of Biomolecular Techniques: JBT, 16, 407-413.

  37. 37. Zhang, H., Li, X., Martin, D.B. and Aebersold, R. (2003) Identification and Quantification of N-Linked Glycoproteins Using Hydrazide Chemistry, Stable Isotope Labeling and Mass Spectrometry. Nature Biotechnology, 660-666.
    http://dx.doi.org/10.1038/nbt827

  38. 38. Yariv, J., Rapport, M.M. and Graf, L. (1962) The Interaction of Glycosides and Saccharides with Antibody to the Corresponding Phenylazo Glycosides. Biochemical Journal, 85, 383.
    http://dx.doi.org/10.1042/bj0850383

  39. 39. Paulsen, B.S., Craik, D.J., Dunstan, D.E., Stone, B.A. and Bacic, A. (2014) The Yariv Reagent: Behaviour in Different Solvents and Interaction with a Gum Arabic Arabinogalactan Protein. Carbohydrate Polymers, 106, 460-468.
    http://dx.doi.org/10.1016/j.carbpol.2014.01.009

  40. 40. Bond, M.R. and Kohler, J.J. (2007) Chemical Methods for Glycoprotein Discovery. Current Opinion in Chemical Biology, 11, 52-58.
    http://dx.doi.org/10.1016/j.cbpa.2006.11.032

  41. 41. Zhang, Y., Giboulot, A., Zivy, M., Valot, B., Jamet, E. and Albenne, C. (2011) Combining Various Strategies to Increase the Coverage of the Plant Cell Wall Glycoproteome. Phytochemistry, 72, 1109-1023.
    http://dx.doi.org/10.1016/j.phytochem.2010.10.019

  42. 42. Klemm, D., Schmauder, H.-P. and Heinze, T. (2005) Cellulose. Biopolymers Online, Wiley-VCH Verlag GmbH & Co. KGaA.

  43. 43. Xu, A., Wang, J. and Wang, H. (2010) Effects of Anionic Structure and Lithium Salts Addition on the Dissolution of Cellulose in 1-Butyl-3-Methylimidazolium-Based Ionic Liquid Solvent Systems. Green Chemistry, 12, 268-275.
    http://dx.doi.org/10.1039/B916882F

  44. 44. Swatloski, R.P., Spear, S.K., Holbrey, J.D. and Rogers, R.D. (2002) Dissolution of Cellose with Ionic Liquids. Journal of the American Chemical Society, 124, 4974-4975.
    http://dx.doi.org/10.1021/ja025790m

  45. 45. Zhang, H., Wu, J., Zhang, J. and He, J. (2005) 1-Allyl-3-Methylimidazolium Chloride Room Temperature Ionic Liquid: A New and Powerful Nonderivatizing Solvent for Cellulose. Macromolecules, 38, 8272-8277.
    http://dx.doi.org/10.1021/ma0505676

  46. 46. Zavrel, M., Bross, D., Funke, M., Büchs, J. and Spiess, A.C. (2009) High-Throughput Screening for Ionic Liquids Dissolving (Ligno-)Cellulose. Bioresource Technology, 100, 2580-2587.
    http://dx.doi.org/10.1016/j.biortech.2008.11.052

  47. 47. Pinkert, A., Marsh, K.N., Pang, S. and Staiger, M.P. (2009) Ionic Liquids and Their Interaction with Cellulose. Chemical Reviews, 109, 6712-6728.
    http://dx.doi.org/10.1021/cr9001947

  48. 48. Jiang, M., Zhao, M., Zhou, Z., Huang, T., Chen, X. and Wang, Y. (2011) Isolation of Cellulose with Ionic Liquid from Steam Exploded Rice Straw. Industrial Crops and Products, 33, 734-738.
    http://dx.doi.org/10.1016/j.indcrop.2011.01.015

  49. 49. Andanson, J.-M., Bordes, E., Devémy, J., Leroux, F., Pádua, A.A. and Gomes, M.F.C. (2014) Understanding the Role of Co-Solvents in the Dissolution of Cellulose in Ionic Liquids. Green Chemistry, 16, 2528-2538.
    http://dx.doi.org/10.1039/c3gc42244e

  50. 50. Fry, S.C. (1988) The Growing Plant Cell Wall: Chemical and Metabolic Analysis. Reprint Edition, The Blackburn Press, Caldwell, 1-333.

  51. 51. Fry, S.C. (1986) Cross-Linking of Matrix Polymers in the Growing Cell Walls of Angiosperms. Annual Review of Plant Physiology, 37, 165-186.
    http://dx.doi.org/10.1016/S0008-6215(00)82963-8

  52. 52. Aspinall, G.O., Craig, J.W.T. and Whyte, J.L. (1968) Lemon-Peel Pectin: Part I. Fractionation and Partial Hydrolysis of Water-Soluble Pectin. Carbohydrate Research, 7, 442-452.
    http://dx.doi.org/10.1016/S0008-6215(00)82963-8

  53. 53. Ray, B., Loutelier-Bourhis, C., Lange, C., Condamine, E., Driouich, A. and Lerouge, P. (2004) Structural Investigation of Hemicellulosic Polysaccharides from Argania Spinosa: Characterisation of a Novel Xyloglucan Motif. Carbohydrate Research, 339, 201-208.
    http://dx.doi.org/10.1016/j.carres.2003.10.011

  54. 54. Norris, F.W. and Resch, C.E. (1937) The Pectic Substances of Plants. Biochemical Journal, 31, 1945-1951.
    http://dx.doi.org/10.1042/bj0311945

  55. 55. Brett, C.T. and Hillman, J.R. (1985) Biochemistry of Plant Cell Walls. CUP Archive.

  56. 56. Levigne, S., Thomas, M., Ralet, M.-C., Quemener, B. and Thibault, J.-F. (2002) Determination of the Degrees of Methylation and Acetylation of Pectins Using a C18 Column and Internal Standards. Food Hydrocolloids, 16, 547-550.
    http://dx.doi.org/10.1016/S0268-005X(02)00015-2

  57. 57. Garna, H., Mabon, N., Robert, C., Cornet, C., Nott, K., Legros, H., et al. (2007) Effect of Extraction Conditions on the Yield and Purity of Apple Pomace Pectin Precipitated But Not Washed by Alcohol. Journal of Food Science, 72, C001-C009.
    http://dx.doi.org/10.1111/j.1750-3841.2006.00227.x

  58. 58. Yeoh, S., Shi, J. and Langrish, T.A.G. (2008) Comparisons between Different Techniques for Water-Based Extraction of Pectin from Orange Peels. Desalination, 218, 229-237.
    http://dx.doi.org/10.1016/j.desal.2007.02.018

  59. 59. Koubala, B.B., Kansci, G., Mbome, L.I., Crépeau, M.-J., Thibault, J.-F. and Ralet, M.-C. (2008) Effect of Extraction Conditions on Some Physicochemical Characteristics of Pectins from “Améliorée” and “Mango” Mango Peels. Food Hydrocolloids, 22, 1345-1351.
    http://dx.doi.org/10.1016/j.foodhyd.2007.07.005

  60. 60. Bertin, C., Rouau, X. and Thibault, J.-F. (1988) Structure and Properties of Sugar Beet Fibres. Journal of the Science of Food and Agriculture, 44, 15-29.
    http://dx.doi.org/10.1002/jsfa.2740440104

  61. 61. Thakur, B.R., Singh, R.K., Handa, A.K. and Rao, M.A. (1997) Chemistry and Uses of Pectin—A Review. Critical Reviews in Food Science & Nutrition, 37, 47-73.
    http://dx.doi.org/10.1080/10408399709527767

  62. 62. Stoddart, R.W., Barrett, A.J. and Northcote, D.H. (1967) Pectic Polysaccharides of Growing Plant Tissues. Biochemical Journal, 102, 194-204.
    http://dx.doi.org/10.1042/bj1020194

  63. 63. Jarvis, M.C., Hall, M.A., Threlfall, D.R. and Friend, J. (1981) The Polysaccharide Structure of Potato Cell Walls: Che- mical Fractionation. Planta, 152, 93-100.
    http://dx.doi.org/10.1007/BF00391179

  64. 64. Jarvis, M.C. (1982) The Proportion of Calcium-Bound Pectin in Plant Cell Walls. Planta, 154, 344-346.
    http://dx.doi.org/10.1007/BF00393913

  65. 65. Barrett, A.J. and Northcote, D.H. (1965) Apple Fruit Pectic Substances. Biochemical Journal, 94, 617-627.
    http://dx.doi.org/10.1042/bj0940617

  66. 66. Kratchanova, M., Panchev, I., Pavlova, E. and Shtereva, L. (1994) Extraction of Pectin from Fruit Materials Pretreated in an Electromagnetic Field of Super-High Frequency. Carbohydrate Polymers, 25, 141-144.
    http://dx.doi.org/10.1016/0144-8617(94)90197-x

  67. 67. Fishman, M.L., Chau, H.K., Hoagland, P. and Ayyad, K. (1999) Characterization of Pectin, Flash-Extracted from Orange Albedo by Microwave Heating, under Pressure. Carbohydrate Research, 323, 126-138.
    http://dx.doi.org/10.1016/S0008-6215(99)00244-X

  68. 68. Fishman, M.L., Chau, H.K., Hoagland, P.D. and Hotchkiss, A.T. (2006) Microwave-Assisted Extraction of Lime Pectin. Food Hydrocolloids, 20, 1170-1177.
    http://dx.doi.org/10.1016/j.foodhyd.2006.01.002

  69. 69. Zhiwei, L., Nan, W. and Mengyu, Z. (2002) The Application of Microwave Assisted Extraction Technique in Food Chemistry. Journal of Wuhan Polytechnic University, 2, 18-21.

  70. 70. Sahari, M.A., Akbarian, A. and Hamedi, M. (2003) Effect of Variety and Acid Washing Method on Extraction Yield and Quality of Sunflower Head Pectin. Food Chemistry, 83, 43-47.
    http://dx.doi.org/10.1016/S0308-8146(03)00034-7

  71. 71. Mesbahi, G., Jamalian, J. and Farahnaky, A. (2005) A Comparative Study on Functional Properties of Beet and Citrus Pectins in Food Systems. Food Hydrocolloids, 19, 731-738.
    http://dx.doi.org/10.1016/j.foodhyd.2004.08.002

  72. 72. Singthong, J., Ningsanond, S., Cui, S.W. and Goff, H.D. (2005) Extraction and Physicochemical Characterization of Krueo Ma Noy Pectin. Food Hydrocolloids, 19, 793-801.
    http://dx.doi.org/10.1016/j.foodhyd.2004.09.007

  73. 73. Liu, Z.D., Wei, G.H., Guo, Y.C. and Kennedy, J.F. (2006) Image Study of Pectin Extraction from Orange Skin Assisted by Microwave. Carbohydrate Polymers, 64, 548-552.
    http://dx.doi.org/10.1016/j.carbpol.2005.11.006

  74. 74. Wang, S., Chen, F., Wu, J., Wang, Z., Liao, X. and Hu, X. (2007) Optimization of Pectin Extraction Assisted by Microwave from Apple Pomace Using Response Surface Methodology. Journal of Food Engineering, 78, 693-700.
    http://dx.doi.org/10.1016/j.jfoodeng.2005.11.008

  75. 75. Wu, J., Peng, K., Zhang, Y., Hu, X., Liao, S., Chen, F., et al. (2009) Comparison of Quality of Apple Pectin between Conventional Solution Extraction and Microwave-Assisted Extraction. Transactions of the Chinese Society of Agricultural Engineering, 25, 350-355.

  76. 76. Prabasari, I., Pettolino, F., Liao, M.-L. and Bacic, A. (2011) Pectic Polysaccharides from Mature Orange (Citrus sinensis) Fruit Albedo Cell Walls: Sequential Extraction and Chemical Characterization. Carbohydrate Polymers, 84, 484-494.

    http://dx.doi.org/10.1016/j.carbpol.2010.12.012

  77. 77. Guo, X., Han, D., Xi, H., Rao, L., Liao, X., Hu, X., et al. (2012) Extraction of Pectin from Navel Orange Peel Assisted by Ultra-High Pressure, Microwave or Traditional Heating: A Comparison. Carbohydrate Polymers, 88, 441-448.
    http://dx.doi.org/10.1016/j.carbpol.2011.12.026

  78. 78. Kratchanova, M., Pavlova, E. and Panchev, I. (2004) The Effect of Microwave Heating of Fresh Orange Peels on the Fruit Tissue and Quality of Extracted Pectin. Carbohydrate Polymers, 56, 181-185.
    http://dx.doi.org/10.1016/j.carbpol.2004.01.009

  79. 79. Godin, B., Agneessens, R., Gofflot, S., Lamaudière, S., Sinnaeve, G., Gerin, P.A., et al. (2011) Revue bibliographique sur les méthodes d’analyse des polysaccharides structuraux des biomasses lignocellulosiques. Biotechnologie, Agro- nomie, Société et Environnement, 15, 165-182.

  80. 80. Favela-Torres, E., Volke-Sepúlveda, T. and Viniegra-González, G. (2006) Production of Hydrolytic Depolymerising Pectinases. Food Technology and Biotechnology, 44, 221.

  81. 81. Sakai, T., Sakamoto, T., Hallaert, J. and Vandamme, E.J. (1993) Pectin, Pectinase, and Protopectinase: Production, Properties, and Applications. Advances in Applied Microbiology, 39, 213-294.
    http://dx.doi.org/10.1016/S0065-2164(08)70597-5

  82. 82. Alkorta, I., Garbisu, C., Llama, M.J. and Serra, J.L. (1998) Industrial Applications of Pectic Enzymes: A Review. Process Biochemistry, 33, 21-28.
    http://dx.doi.org/10.1016/S0032-9592(97)00046-0

  83. 83. Jayani, R.S., Saxena, S. and Gupta, R. (2005) Microbial Pectinolytic Enzymes: A Review. Process Biochemistry, 40, 2931-2944.
    http://dx.doi.org/10.1016/j.procbio.2005.03.026

  84. 84. Pauly, M., Qin, Q., Greene, H., Albersheim, P., Darvill, A. and York, W.S. (2001) Changes in the Structure of Xyloglucan during Cell Elongation. Planta, 212, 842-850.
    http://dx.doi.org/10.1007/s004250000448

  85. 85. Byg, I., Diaz, J., Ogendal, L.H., Harholt, J., Jorgensen, B., Rolin, C., et al. (2012) Large-Scale Extraction of Rhamnogalacturonan I from Industrial Potato Waste. Food Chemistry, 131, 1207-1216.
    http://dx.doi.org/10.1016/j.foodchem.2011.09.106

  86. 86. Wise, L.E. and Ratliff, E.K. (1947) Quantitative Isolation of Hemicelluloses and Summative Analysis of Wood. Analytical Chemistry, 19, 459-462.
    http://dx.doi.org/10.1021/ac60007a010

  87. 87. Norris, F.W. and Preece, I.A. (1930) Studies on Hemicelluloses: The Hemicelluloses of Wheat Bran. Biochemical Journal, 24, 59.
    http://dx.doi.org/10.1042/bj0240059

  88. 88. Lawther, J.M., Sun, R. and Banks, W.B. (1996) Effects of Extraction Conditions and Alkali Type on Yield and Composition of Wheat Straw Hemicellulose. Journal of Applied Polymer Science, 60, 1827-1837.
    http://dx.doi.org/10.1002/(SICI)1097-4628(19960613)60:11<1827::AID-APP6>3.0.CO;2-N

  89. 89. Rutenberg, M.W. and William, H. (1957) Process for Extraction of Hemicellulose. Google Patents.

  90. 90. Doner, L.W. and Hicks, K.B. (1997) Isolation of Hemicellulose from Corn Fiber by Alkaline Hydrogen Peroxide Extraction. Cereal Chemistry Journal, 74, 176-181.
    http://dx.doi.org/10.1094/CCHEM.1997.74.2.176

  91. 91. Sjostrom, E. and Alén, R. (1998) Analytical Methods in Wood Chemistry, Pulping, and Papermaking. Springer, New York, 37-77.

  92. 92. Chanliaud, E., Saulnier, L. and Thibault, J.-F. (1995) Alkaline Extraction and Characterisation of Heteroxylans from Maize Bran. Journal of Cereal Science, 21, 195-203.
    http://dx.doi.org/10.1016/0733-5210(95)90035-7

  93. 93. Sun, R.C. and Tomkinson, J. (2002) Characterization of Hemicelluloses Obtained by Classical and Ultrasonically Assisted Extractions from Wheat Straw. Carbohydrate Polymers, 50, 263-271.
    http://dx.doi.org/10.1016/S0144-8617(02)00037-1

  94. 94. Hagglund, E., Lindberg, B. and McPherson, J. (1956) Dimethylsulphoxide, a Solvent for Hemicelluloses. Acta Chemica Scandinavica, 10, 1160-1164.
    http://dx.doi.org/10.3891/acta.chem.scand.10-1160

  95. 95. Buranov, A.U. and Mazza, G. (2010) Extraction and Characterization of Hemicelluloses from Flax Shives by Different Methods. Carbohydrate Polymers, 79, 17-25.
    http://dx.doi.org/10.1016/j.carbpol.2009.06.014

  96. 96. Hromadkova, Z., Kováciková, J. and Ebringerová, A. (1999) Study of the Classical and Ultrasound-Assisted Extraction of the Corn Cob Xylan. Industrial Crops and Products, 9, 101-109.
    http://dx.doi.org/10.1016/S0926-6690(98)00020-X

  97. 97. Janker-Obermeier, I., Sieber, V., Faulstich, M. and Schieder, D. (2012) Solubilization of Hemicellulose and Lignin from Wheat Straw through Microwave-Assisted Alkali Treatment. Industrial Crops and Products, 39, 198-203.
    http://dx.doi.org/10.1016/j.indcrop.2012.02.022

  98. 98. Krawczyk, H., Persson, T., Andersson, A. and Jonsson, A.-S. (2008) Isolation of Hemicelluloses from Barley Husks. Food and Bioproducts Processing, 86, 31-36.
    http://dx.doi.org/10.1016/j.fbp.2007.10.018

  99. 99. Filho, E.X.F. (1998) Hemicellulase and Biotechnology. S. G. Pandalai.

  100. 100. Reilly, P.J. (1981) Xylanases: Structure and Function. Trends in the Biology of Fermentations for Fuels and Chemicals. Springer, 111-129.
    http://dx.doi.org/10.1007/978-1-4684-3980-9_8

  101. 101. Collins, T., Gerday, C. and Feller, G. (2005) Xylanases, Xylanase Families and Extremophilic Xylanases. FEMS Microbiology Reviews, 29, 3-23.
    http://dx.doi.org/10.1016/j.femsre.2004.06.005

  102. 102. Wyman, C.E., Decker, S.R., Himmel, M.E., Brady, J.W., Skopec, C.E. and Viikari, L. (2005) Hydrolysis of Cellulose and Hemicellulose. Polysaccharides: Structural Diversity and Functional Versatility, 1, 1023-1062.

  103. 103. Dhawan, S. and Kaur, J. (2007) Microbial Mannanases: An Overview of Production and Applications. Critical Reviews in Biotechnology, 27, 197-216.
    http://dx.doi.org/10.1080/07388550701775919

  104. 104. Palm, M. and Zacchi, G. (2003) Extraction of Hemicellulosic Oligosaccharides from Spruce Using Microwave Oven or Steam Treatment. Biomacromolecules, 4, 617-623.
    http://dx.doi.org/10.1021/bm020112d

  105. 105. Pepper, J.M., Baylis, P.E.T. and Adler, E. (1959) The Isolation and Properties of Lignins Obtained by the Acidolysis of Spruce and Aspen Woods in Dioxane-Water Medium. Canadian Journal of Chemistry, 37, 1241-1248.
    http://dx.doi.org/10.1139/v59-183

  106. 106. Jaaskelainen, A.S., Sun, Y., Argyropoulos, D.S., Tamminen, T. and Hortling, B. (2003) The Effect of Isolation Method on the Chemical Structure of Residual Lignin. Wood Science and Technology, 37, 91-102.
    http://dx.doi.org/10.1007/s00226-003-0163-y

  107. 107. Mortha, G., Nikandrov, A., Robert, D., Lachenal, D. and Zaroubine, M.Y. (2001) Characteristics of Lignins Extracted from Oak Wood and Kraft Pulps by Acetic Acid/ZnCl2 Acidolysis: Comparison with Other Methods. Proceedings of 11th International Symposium on Wood and Pulping Chemistry, Nice, 11-14 June 2001, 245-250.

  108. 108. Evtuguin, D.V., Neto, C.P., Silva, A.M., Domingues, P.M., Amado, F.M., Robert, D., et al. (2001) Comprehensive Study on the Chemical Structure of Dioxane Lignin from Plantation Eucalyptus globulus Wood. Journal of Agricultural and Food Chemistry, 49, 4252-42561.
    http://dx.doi.org/10.1021/jf010315d

  109. 109. Guerra, A., Filpponen, I., Lucia, L.A., Saquing, C., Baumberger, S. and Argyropoulos, D.S. (2006) Toward a Better Understanding of the Lignin Isolation Process from Wood. Journal of Agricultural and Food Chemistry, 54, 5939- 5947.
    http://dx.doi.org/10.1021/jf060722v

  110. 110. Guerra, A., Filpponen, I., Lucia, L.A. and Argyropoulos, D.S. (2006) Comparative Evaluation of Three Lignin Isolation Protocols for Various Wood Species. Journal of Agricultural and Food Chemistry, 54, 9696-9705.
    http://dx.doi.org/10.1021/jf062433c

  111. 111. Pew, J.C. and Weyna, P. (1962) Fine Grinding, Enzyme Digestion, and the Lignin-Cellulose Bond in Wood. Tappi, 45, 247-256.

  112. 112. Balakshin, M.Y., Capanema, E.A. and Chang, H.-M. (2008) Recent Advances in the Isolation and Analysis of Lignins and Lignin-Carbohydrate Complexes. In: Fellow TQHBS, Characterization of Lignocellulosic Materials, Blackwell Publishing Ltd., 148-170.
    http://dx.doi.org/10.1002/9781444305425.ch9

  113. 113. Wu, S. and Argyropoulos, D.S. (2003) An Improved Method for Isolating Lignin in High Yield and Purity. Journal of Pulp and Paper Science, 29, 235-240.

  114. 114. Sun, X.-F., Jing, Z., Fowler, P., Wu, Y. and Rajaratnam, M. (2011) Structural Characterization and Isolation of Lignin and Hemicelluloses from Barley Straw. Industrial Crops and Products, 33, 588-598.
    http://dx.doi.org/10.1016/j.indcrop.2010.12.005

  115. 115. Kilpelainen, I., Xie, H., King, A., Granstrom, M., Heikkinen, S. and Argyropoulos, D.S. (2007) Dissolution of Wood in Ionic Liquids. Journal of Agricultural and Food Chemistry, 55, 9142-9148.
    http://dx.doi.org/10.1021/jf071692e

  116. 116. Yang, D., Zhong, L.-X., Yuan, T.-Q., Peng, X.-W. and Sun, R.-C. (2013) Studies on the Structural Characterization of Lignin, Hemicelluloses and Cellulose Fractionated by Ionic Liquid Followed by Alkaline Extraction from Bamboo. Industrial Crops and Products, 43, 141-149.
    http://dx.doi.org/10.1016/j.indcrop.2012.07.024

  117. 117. Prado, R., Erdocia, X. and Labidi, J. (2013) Lignin Extraction and Purification with Ionic Liquids. Journal of Chemical Technology and Biotechnology, 88, 1248-1257.
    http://dx.doi.org/10.1002/jctb.3965

  118. 118. Muhammad, N., Man, Z. and Bustam Khalil, M.A. (2012) Ionic liquid—A Future Solvent for the Enhanced Uses of Wood Biomass. European Journal of Wood and Wood Products, 70, 125-133.
    http://dx.doi.org/10.1007/s00107-011-0526-2

  119. 119. Chanzy, H., Peguy, A., Chaunis, S. and Monzie, P. (1980) Oriented Cellulose Films and Fibers from a Mesophase System. Journal of Polymer Science: Polymer Physics Edition, 18, 1137-1144.
    http://dx.doi.org/10.1002/pol.1980.180180517

  120. 120. McCormick, C.L., Callais, P.A. and Hutchinson Jr., B.H. (1985) Solution Studies of Cellulose in Lithium Chloride and N, N-Dimethylacetamide. Macromolecules, 18, 2394-2401.
    http://dx.doi.org/10.1021/ma00154a010

  121. 121. Cai, J. and Zhang, L. (2005) Rapid Dissolution of Cellulose in LiOH/Urea and NaOH/Urea Aqueous Solutions. Macro- molecular Bioscience, 5, 539-548.
    http://dx.doi.org/10.1002/mabi.200400222

  122. 122. Nishio, Y., Roy, S.K. and Manley, R.S.J. (1987) Blends of Cellulose with Polyacrylonitrile Prepared from N, N-Dimethylacetamide-Lithium Chloride Solutions. Polymer, 28, 1385-1390.
    http://dx.doi.org/10.1016/0032-3861(87)90456-3

  123. 123. Isogai, A. and Atalla, R.H. (1998) Dissolution of Cellulose in Aqueous NaOH Solutions. Cellulose, 5, 309-319.
    http://dx.doi.org/10.1080/07366579008050914

  124. 124. Dawsey, T.R. and McCormick, C.L. (1990) The Lithium Chloride/Dimethylacetamide Solvent for Cellulose: A Literature Review. Journal of Macromolecular Science—Reviews in Macromolecular Chemistry and Physics, 30, 405-440.
    http://dx.doi.org/10.1080/07366579008050914

  125. 125. Seddon, K.R. (1997) Ionic Liquids for Clean Technology. Journal of Chemical Technology and Biotechnology, 68, 351-356.
    http://dx.doi.org/10.1002/(SICI)1097-4660(199704)68:4<351::AID-JCTB613>3.0.CO;2-4

  126. 126. Sheldon, R.A., Lau, R.M., Sorgedrager, M.J., van Rantwijk, F. and Seddon, K.R. (2002) Biocatalysis in Ionic Liquids. Green Chemistry, 4, 147-151.
    http://dx.doi.org/10.1039/b110008b

  127. 127. Lee, S.H. and Lee, S.B. (2005) The Hildebrand Solubility Parameters, Cohesive Energy Densities and Internal Ener- gies of 1-Alkyl-3-Methylimidazolium-Based Room Temperature Ionic Liquids. Chemical Communications, 2005, 3469-3471.
    http://dx.doi.org/10.1039/b503740a

  128. 128. Xie, H., Zhang, S. and Li, S. (2006) Chitin and Chitosan Dissolved in Ionic Liquids as Reversible Sorbents of CO2. Green Chemistry, 8, 630-633.
    http://dx.doi.org/10.1039/b517297g

  129. 129. Vesa, M. and Reijo, A. (WO 017001 A1) Dissolution Method for Lignocellulosic Materials.

  130. 130. Fort, D.A., Remsing, R.C., Swatloski, R.P., Moyna, P., Moyna, G. and Rogers, R.D. (2007) Can Ionic Liquids Dissolve wood? Processing and Analysis of Lignocellulosic Materials with 1-n-Butyl-3-Methylimidazolium Chloride. Green Chemistry, 9, 63.
    http://dx.doi.org/10.1039/B607614A

  131. 131. Lan, W., Liu, C.-F. and Sun, R.-C. (2011) Fractionation of Bagasse into Cellulose, Hemicelluloses, and Lignin with Ionic Liquid Treatment Followed by Alkaline Extraction. Journal of Agricultural and Food Chemistry, 59, 8691-8701.
    http://dx.doi.org/10.1021/jf201508g

  132. 132. Sun, N., Rahman, M., Qin, Y., Maxim, M.L., Rodríguez, H. and Rogers, R.D. (2009) Complete Dissolution and Partial Delignification of Wood in the Ionic Liquid 1-Ethyl-3-Methylimidazolium Acetate. Green Chemistry, 11, 646.
    http://dx.doi.org/10.1039/b822702k

  133. 133. Miyafuji, H., Miyata, K., Saka, S., Ueda, F. and Mori, M. (2009) Reaction Behavior of Wood in an Ionic Liquid, 1-Ethyl-3-Methylimidazolium Chloride. Journal of Wood Science, 55, 215-219.
    http://dx.doi.org/10.1007/s10086-009-1020-x

  134. 134. Singh, S., Simmons, B.A. and Vogel, K.P. (2009) Visualization of Biomass Solubilization and Cellulose Regeneration during Ionic Liquid Pretreatment of Switchgrass. Biotechnology and Bioengineering, 104, 68-75.
    http://dx.doi.org/10.1002/bit.22386

  135. 135. Abe, M., Yamanaka, S., Yamada, H., Yamada, T. and Ohno, H. (2015) Almost Complete Dissolution of Woody Biomass with Tetra-n-Butylphosphonium Hydroxide Aqueous Solution at 60℃. Green Chemistry, 17, 4432-4438.
    http://dx.doi.org/10.1039/C5GC00646E

  136. 136. Mosier, N., Wyman, C., Dale, B., Elander, R., Lee, Y.Y., Holtzapple, M., et al. (2005) Features of Promising Technologies for Pretreatment of Lignocellulosic Biomass. Bioresource Technology, 96, 673-686.
    http://dx.doi.org/10.1016/j.biortech.2004.06.025

  137. 137. Yan, Y., Li, X., Wan, M., Chen, J., Li, S., Cao, M., et al. (2015) Effect of Extraction Methods on Property and Bioactivity of Water-Soluble Polysaccharides from Amomum villosum. Carbohydrate Polymers, 117, 632-635.
    http://dx.doi.org/10.1016/j.carbpol.2014.09.070

  138. 138. Wang, J., Zhang, J., Zhao, B., Wang, X., Wu, Y. and Yao, J. (2010) A Comparison Study on Microwave-Assisted Extraction of Potentilla anserina L. Polysaccharides with Conventional Method: Molecule Weight and Antioxidant Activities Evaluation. Carbohydrate Polymers, 80, 84-93.
    http://dx.doi.org/10.1016/j.carbpol.2009.10.073

  139. 139. Mishra, A., Mishra, S., Bhargav, S., Bhargava, C.S. and Thakur, M. (2014) Microwave Assisted Extraction, Antioxidant Potential and Chromatographic Studies of Some Rasayana Drugs. Chinese Journal of Integrative Medicine, 21, 1-7.

  140. 140. Zeng, H., Zhang, Y., Lin, S., Jian, Y., Miao, S. and Zheng, B. (2015) Ultrasonic-Microwave Synergistic Extraction (UMSE) and Molecular Weight Distribution of Polysaccharides from Fortunella margarita (Lour.) Swingle. Separation and Purification Technology, 144, 97-106.
    http://dx.doi.org/10.1016/j.seppur.2015.02.015

  141. 141. Fan, T., Hu, J., Fu, L. and Zhang, L. (2015) Optimization of Enzymolysis-Ultrasonic Assisted Extraction of Polysaccharides from Momordica charabtia L. by Response Surface Methodology. Carbohydrate Polymers, 115, 701-706.
    http://dx.doi.org/10.1016/j.carbpol.2014.09.009

  142. 142. Puri, M., Sharma, D. and Barrow, C.J. (2012) Enzyme-Assisted Extraction of Bioactives from Plants. Trends in Biotechnology, 30, 37-44.
    http://dx.doi.org/10.1016/j.tibtech.2011.06.014

  143. 143. Gornall, A.G., Bardawill, C.J. and David, M.M. (1949) Determination of Serum Proteins by Means of the Biuret Reaction.

  144. 144. Lowry, O.H., Rosebrough, N.J., Farr, A.L., Randall, R.J., et al. (1951) Protein Measurement with the Folin Phenol Reagent. Journal of Biological Chemistry, 193, 265-275.

  145. 145. Smith, P.K., Krohn, R.I., Hermanson, G.T., Mallia, A.K., Gartner, F.H., Provenzano, M., et al. (1985) Measurement of Protein Using Bicinchoninic Acid. Analytical Biochemistry, 150, 76-85.
    http://dx.doi.org/10.1016/0003-2697(85)90442-7

  146. 146. Bradford, M.M. (1976) A Rapid and Sensitive Method for the Quantitation of Microgram Quantities of Protein Utilizing the Principle of Protein-Dye Binding. Analytical Biochemistry, 72, 248-254.
    http://dx.doi.org/10.1016/0003-2697(76)90527-3

  147. 147. Stoscheck, C.M. (1987) Protein Assay Sensitive at Nanogram Levels. Analytical Biochemistry, 160, 301-305.
    http://dx.doi.org/10.1016/0003-2697(87)90051-0

  148. 148. Watanabe, N., Kamei, S., Ohkubo, A., Yamanaka, M., Ohsawa, S., Makino, K., et al. (1986) Urinary Protein as Measured with a Pyrogallol Red-Molybdate Complex, Manually and in a Hitachi 726 Automated Analyzer. Clinical Chemistry, 32, 1551-1554.

  149. 149. Fujita, Y., Mori, I. and Kitano, S. (1984) Determination of Proteins by Using the Color Reaction with Pyrocatechol Violet-Molybdenum (VI) Complex. Chemical & Pharmaceutical Bulletin, 32, 4161-4164.
    http://dx.doi.org/10.1248/cpb.32.4161

  150. 150. Antharavally, B.S., Mallia, K.A., Rangaraj, P., Haney, P. and Bell, P.A. (2009) Quantitation of Proteins Using a Dye-Metal-Based Colorimetric Protein Assay. Analytical Biochemistry, 385, 342-425.
    http://dx.doi.org/10.1016/j.ab.2008.11.024

  151. 151. Dishe, Z. and Popper, H. (1926) Uber Eine Neue Kolorimetrischen Mikrobestim Mungs Methode der Kohlehydrate in Organen und Korpersaften. Biologische Zeitung, 175, 371-411.

  152. 152. Tillmans, J. and Philippi, K. (1929) The Carbohydrate Content of the Important Proteins of Foodstuffs and a Colorimetric Procedure for the Determination of Nitrogen-Free Sugar in Protein. Biochemische Zeitschrift, 215, 36-60.

  153. 153. Dische, Z. (1947) A New Specific Color Reaction of Hexuronic Acids. Journal of Biological Chemistry, 167, 189-198.

  154. 154. Bitter, T. and Muir, H.M. (1962) A Modified Uronic Acid Carbazole Reaction. Analytical Biochemistry, 4, 330-334.
    http://dx.doi.org/10.1016/0003-2697(62)90095-7

  155. 155. Monsigny, M., Petit, C. and Roche, A.-C. (1988) Colorimetric Determination of Neutral Sugars by a Resorcinol Sulfuric Acid Micromethod. Analytical Biochemistry, 175, 525-530.
    http://dx.doi.org/10.1016/0003-2697(88)90578-7

  156. 156. Blumenkrantz, N. and Asboe-Hansen, G. (1973) New Method for Quantitative Determination of Uronic Acids. Analytical Biochemistry, 54, 484-489.
    http://dx.doi.org/10.1016/0003-2697(73)90377-1

  157. 157. Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, P.A. and Smith, F. (1956) Colorimetric Method for Determination of Sugars and Related Substances. Analytical Chemistry, 28, 350-356.
    http://dx.doi.org/10.1021/ac60111a017

  158. 158. Masuko, T., Minami, A., Iwasaki, N., Majima, T., Nishimura, S.-I. and Lee, Y.C. (2005) Carbohydrate Analysis by a Phenol-Sulfuric Acid Method in Microplate Format. Analytical Biochemistry, 339, 69-72.
    http://dx.doi.org/10.1016/j.ab.2004.12.001

  159. 159. Montreuil, J. and Spik, G. (1963) Microdosage des glucides: Méthodes colorimétriques de dosage des glucides totaux. Faculté des Sciences de Lille.

  160. 160. Wicker, L. and Leiting, V.A. (1995) Microscale Galacturonic Acid Assay. Analytical Biochemistry, 229, 148-50.
    http://dx.doi.org/10.1006/abio.1995.1395

  161. 161. Johnson, D.B., Moore, W.E. and Zank, L.C. (1961) The Spectrophotometric Determination of Lignin in Small Wood Samples. Tappi, 44, 793-798.

  162. 162. Morrison, I. (1972) Improvements in the Acetyl Bromide Technique to Determine Lignin and Digestibility and Its Application to Legumes. Journal of the Science of Food and Agriculture, 23, 1463-1469.
    http://dx.doi.org/10.1002/jsfa.2740231211

  163. 163. Morrison, I.M. (1972) A Semi-Micro Method for the Determination of Lignin and Its Use in Predicting the Digestibility of Forage Crops. Journal of the Science of Food and Agriculture, 23, 455-463.
    http://dx.doi.org/10.1002/jsfa.2740230405

  164. 164. Fengel, D. and Wegener, G. (1983) Wood: Chemistry, Ultrastructure, Reactions. Walter de Gruyter.
    http://dx.doi.org/10.1515/9783110839654

  165. 165. Moreira-Vilar, F.C., de Cássia Siqueira-Soares, R., Finger-Teixeira, A., de Oliveira, D.M., Ferro, A.P., da Rocha, G.J., et al. (2014) The Acetyl Bromide Method Is Faster, Simpler and Presents Best Recovery of Lignin in Different Herbaceous Tissues than Klason and Thioglycolic Acid Methods. PLoS ONE, 9, e110000.
    http://dx.doi.org/10.1371/journal.pone.0110000

  166. 166. Kline, L.M., Hayes, D.G., Womac, A.R. and Labbe, N. (2010) Simplified Determination of Lignin Content in Hard and Soft Woods via UV-Spectrophotometric Analysis of Biomass Dissolved in Ionic Liquids. BioResources, 5, 1366-1383.

  167. 167. Nkansah, K. and Dawson-Andoh, B. (2010) Rapid Characterization of Biomass Using Fluorescence Spectroscopy Coupled with Multivariate Data Analysis. II. Northern Red Oak (Quercus rubra). Journal of Renewable and Sustainable Energy, 2, Article ID: 043101.
    http://dx.doi.org/10.1063/1.3404181

  168. 168. Nkansah, K. and Dawson-Andoh, B. (2010) Rapid Characterization of Biomass Using Fluorescence Spectroscopy Coupled with Multivariate Data Analysis. I. Yellow Poplar (Liriodendron tulipifera L.). Journal of Renewable and Sustainable Energy, 2, Article ID: 023103.
    http://dx.doi.org/10.1063/1.3290749

  169. 169. Schulz, H. and Baranska, M. (2007) Identification and Quantification of Valuable Plant Substances by IR and Raman Spectroscopy. Vibrational Spectroscopy, 43, 13-25.
    http://dx.doi.org/10.1016/j.vibspec.2006.06.001

  170. 170. Sene, C.F., McCann, M.C., Wilson, R.H. and Grinter, R. (1994) Fourier-Transform Raman and Fourier-Transform Infrared Spectroscopy (An Investigation of Five Higher Plant Cell Walls and Their Components). Plant Physiology, 106, 1623-1631.

  171. 171. Larkin, P. (2011) Infrared and Raman Spectroscopy; Principles and Spectral Interpretation. Elsevier, Amsterdam, 1-7.

  172. 172. Atalla, R.H. and Dimick, B.E. (1975) Raman-Spectral Evidence for Differences between the Conformations of Cellulose I and Cellulose II. Carbohydrate Research, 39, C1-C3.
    http://dx.doi.org/10.1016/S0008-6215(00)82656-7

  173. 173. Fengel, D. and Ludwig, M. (1991) Moglichkeiten und Grenzen der FTIR-Spektroskopie bei der Charakterisierung von Cellulose. I: Vergleich von Verschiedenen Cellulosefasern und Bakterien-Cellulose. Das Papier, 45, 45-51.

  174. 174. Langkilde, F.W. and Svantesson, A. (1995) Identification of Celluloses with Fourier-Transform (FT) Mid-Infrared, FT-Raman and Near-Infrared Spectrometry. Journal of Pharmaceutical and Biomedical Analysis, 13, 409-414.
    http://dx.doi.org/10.1016/0731-7085(95)01298-Y

  175. 175. Agarwal, U.P. (2014) 1064 nm FT-Raman Spectroscopy for Investigations of Plant Cell Walls and Other Biomass Materials. Frontiers in Plant Science, 5.
    http://dx.doi.org/10.3389/fpls.2014.00490

  176. 176. Hulleman, S.H., van Hazendonk, J.M. and van Dam, J.E. (1994) Determination of Crystallinity in Native Cellulose from Higher Plants with Diffuse Reflectance Fourier Transform Infrared Spectroscopy. Carbohydrate Research, 261, 163-172.
    http://dx.doi.org/10.1016/0008-6215(94)80015-4

  177. 177. Rowe, R.C., McKillop, A.G. and Bray, D. (1994) The Effect of Batch and Source Variation on the Crystallinity of Microcrystalline Cellulose. International Journal of Pharmaceutics, 101, 169-172.
    http://dx.doi.org/10.1021/ma970768c

  178. 178. Kataoka, Y. and Kondo, T. (1998) FT-IR Microscopic Analysis of Changing Cellulose Crystalline Structure during Wood Cell Wall Formation. Macromolecules, 31, 760-764.
    http://dx.doi.org/10.1021/ma970768c

  179. 179. Cael, J.J., Gardner, K.H., Koenig, J.L. and Blackwell, J. (1975) Infrared and Raman Spectroscopy of Carbohydrates. Paper V. Normal Coordinate Analysis of Cellulose I. The Journal of Chemical Physics, 62, 1145-1153.
    http://dx.doi.org/10.1063/1.430558

  180. 180. Marchessault, R.H., Sundararajan, P.R., et al. (1983) Cellulose. The Polysaccharides, 2, 11-95.
    http://dx.doi.org/10.1016/b978-0-12-065602-8.50007-8

  181. 181. Mathlouthi, M. and Koenig, J.L. (1986) Vibrational Spectra of Carbohydrates. Advances in Carbohydrate Chemistry and Biochemistry, 44, 7-89.
    http://dx.doi.org/10.1016/S0065-2318(08)60077-3

  182. 182. Stewart, D. and Morrison, I.M. (1992) Ft-ir Spectroscopy as a Tool for the Study of Biological and Chemical Treatments of Barley Straw. Journal of the Science of Food and Agriculture, 60, 431-436.
    http://dx.doi.org/10.1002/jsfa.2740600405

  183. 183. Sun, R.C. and Hughes, S. (1999) Fractional Isolation and Physico-Chemical Characterization of Alkali-Soluble Polysaccharides from Sugar Beet Pulp. Carbohydrate Polymers, 38, 273-281.
    http://dx.doi.org/10.1016/S0144-8617(98)00102-7

  184. 184. Sun, R. and Hughes, S. (1998) Fractional Extraction and Physico-Chemical Characterization of Hemicelluloses and Cellulose from Sugar Beet Pulp. Carbohydrate Polymers, 36, 293-299.
    http://dx.doi.org/10.1016/S0144-8617(97)00255-5

  185. 185. Sun, R., Fang, J.M., Rowlands, P. and Bolton, J. (1998) Physicochemical and Thermal Characterization of Wheat Straw Hemicelluloses and Cellulose. Journal of Agricultural and Food Chemistry, 46, 2804-2809.
    http://dx.doi.org/10.1021/jf971078a

  186. 186. Sun, R., Lawther, J.M. and Banks, W.B. (1996) Fractional and Structural Characterization of Wheat Straw Hemicelluloses. Carbohydrate Polymers, 29, 325-331.
    http://dx.doi.org/10.1016/S0144-8617(96)00018-5

  187. 187. Filippov, M.P. (1992) Practical Infrared Spectroscopy of Pectic Substances. Food Hydrocolloids, 6, 115-142.
    http://dx.doi.org/10.1016/S0268-005X(09)80060-X

  188. 188. Engelsen, S.B. and Norgaard, L. (1996) Comparative Vibrational Spectroscopy for Determination of Quality Parameters in Amidated Pectins as Evaluated by Chemometrics. Carbohydrate Polymers, 30, 9-24.
    http://dx.doi.org/10.1016/S0144-8617(96)00068-9

  189. 189. Coimbra, M.A., Barros, A., Barros, M., Rutledge, D.N. and Delgadillo, I. (1998) Multivariate Analysis of Uronic Acid and Neutral Sugars in Whole Pectic Samples by FT-IR Spectroscopy. Carbohydrate Polymers, 37, 241-248.
    http://dx.doi.org/10.1016/S0144-8617(98)00066-6

  190. 190. Chatjigakis, A.K., Pappas, C., Proxenia, N., Kalantzi, O., Rodis, P. and Polissiou, M. (1998) FT-IR Spectroscopic Determination of the Degree of Esterification of Cell Wall Pectins from Stored Peaches and Correlation to Textural Changes. Carbohydrate Polymers, 37, 395-408.
    http://dx.doi.org/10.1016/S0144-8617(98)00057-5

  191. 191. Barros, A.S., Mafra, I., Ferreira, D., Cardoso, S., Reis, A., Da Silva, J.L., et al. (2002) Determination of the Degree of Methylesterification of Pectic Polysaccharides by FT-IR Using an Outer Product PLS1 Regression. Carbohydrate Polymers, 50, 85-94.
    http://dx.doi.org/10.1016/S0144-8617(02)00017-6

  192. 192. Manrique, G.D. and Lajolo, F.M. (2002) FT-IR Spectroscopy as a Tool for Measuring Degree of Methyl Esterification in Pectins Isolated from Ripening Papaya Fruit. Postharvest Biology and Technology, 25, 99-107.
    http://dx.doi.org/10.1016/S0925-5214(01)00160-0

  193. 193. Synytsya, A., Copiková, J., Matějka, P. and Machovic, V. (2003) Fourier Transform Raman and Infrared Spectroscopy of Pectins. Carbohydrate Polymers, 54, 97-106.
    http://dx.doi.org/10.1016/S0144-8617(03)00158-9

  194. 194. Kacurakova, M., Capek, P., Sasinkova, V., Wellner, N. and Ebringerova, A. (2000) FT-IR Study of Plant Cell Wall Model Compounds: Pectic Polysaccharides and Hemicelluloses. Carbohydrate Polymers, 43, 195-203.
    http://dx.doi.org/10.1016/S0144-8617(00)00151-X

  195. 195. Kacuráková, M. and Wilson, R.H. (2001) Developments in Mid-Infrared FT-IR Spectroscopy of Selected Carbo- hydrates. Carbohydrate Polymers, 44, 291-303.
    http://dx.doi.org/10.1016/S0144-8617(00)00245-9

  196. 196. Lupoi, J.S., Singh, S., Simmons, B.A. and Henry, R.J. (2014) Assessment of Lignocellulosic Biomass Using Analytical Spectroscopy: An Evolution to High-Throughput Techniques. BioEnergy Research, 7, 1-23.
    http://dx.doi.org/10.1007/s12155-013-9352-1

  197. 197. Bjarnestad, S. and Dahlman, O. (2002) Chemical Compositions of Hardwood and Softwood Pulps Employing Photoacoustic Fourier Transform Infrared Spectroscopy in Combination with Partial Least-Squares Analysis. Analytical Chemistry, 74, 5851-5858.
    http://dx.doi.org/10.1021/ac025926z

  198. 198. Liu, L., Ye, X.P., Womac, A.R. and Sokhansanj, S. (2010) Variability of Biomass Chemical Composition and Rapid Analysis Using FT-NIR Techniques. Carbohydrate Polymers, 81, 820-829.
    http://dx.doi.org/10.1016/j.carbpol.2010.03.058

  199. 199. Jaaskelainen, A.-S., Saariaho, A.-M. and Vuorinen, T. (2005) Quantification of Lignin and Hexenuronic Acid in Bleached Hardwood Kraft Pulps: A New Calibration Method for UVRR Spectroscopy and Evaluation of the Con- ventional Methods. Journal of Wood Chemistry and Technology, 25, 51-65.
    http://dx.doi.org/10.1081/WCT-200058239

  200. 200. Agarwal, U.P. (2011) Lignin Quantitation by FT-Raman Spectroscopy. Proceedings 16th International Symposium on Wood, Fiber and Pulping Chemistry, Tianjin, 8-10 June 2011, 170-173.

  201. 201. Sun, L., Varanasi, P., Yang, F., Loqué, D., Simmons, B.A. and Singh, S. (2012) Rapid Determination of Syringyl: Guaiacyl Ratios Using FT-Raman Spectroscopy. Biotechnology and Bioengineering, 109, 647-656.
    http://dx.doi.org/10.1002/bit.24348

  202. 202. Ona, T., Sonoda, T., Ito, K., Shibatal, M., Katayama, T., Kato, T., et al. (1998) Non-Destructive Determination of Lig- nin Syringyl/Guaiacyl Monomeric Composition in Native Wood by Fourier Transform Raman Spectroscopy. Journal of Wood Chemistry and Technology, 18, 43-51.
    http://dx.doi.org/10.1080/02773819809350124

  203. 203. Saariaho, A.-M., Argyropoulos, D.S., Jaaskelainen, A.-S. and Vuorinen, T. (2005) Development of the Partial Least Squares Models for the Interpretation of the UV Resonance Raman Spectra of Lignin Model Compounds. Vibrational Spectroscopy, 37, 111-121.
    http://dx.doi.org/10.1016/j.vibspec.2004.08.001

  204. 204. Saariaho, A.-M., Jaaskelainen, A.-S., Nuopponen, M. and Vuorinen, T. (2003) Ultra Violet Resonance Raman Spec- troscopy in Lignin Analysis: Determination of Characteristic Vibrations of p-Hydroxyphenyl, Guaiacyl, and Syringyl Lignin Structures. Applied Spectroscopy, 57, 58-66.
    http://dx.doi.org/10.1366/000370203321165214

  205. 205. Jose, C., Gutiérrez, A., Rodriguez, I.M., Ibarra, D. and Martinez, A.T. (2007) Composition of Non-Woody Plant Lignins and Cinnamic Acids by Py-GC/MS, Py/TMAH and FT-IR. Journal of Analytical and Applied Pyrolysis, 79, 39-46.
    http://dx.doi.org/10.1016/j.jaap.2006.09.003

  206. 206. Casas, A., Oliet, M., Alonso, M.V. and Rodriguez, F. (2012) Dissolution of Pinus radiata and Eucalyptus globulus Woods in Ionic Liquids under Microwave Radiation: lignin Regeneration and Characterization. Separation and Purification Technology, 97, 115-122.
    http://dx.doi.org/10.1016/j.seppur.2011.12.032

  207. 207. Kihara, M., Takayama, M., Wariishi, H. and Tanaka, H. (2002) Determination of the Carbonyl Groups in Native Lignin Utilizing Fourier Transform Raman Spectroscopy. Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy, 58, 2213-2221.
    http://dx.doi.org/10.1016/S1386-1425(01)00693-X

  208. 208. Boudart, G., Jamet, E., Rossignol, M., Lafitte, C., Borderies, G., Jauneau, A., et al. (2005) Cell Wall Proteins in Apoplastic Fluids of Arabidopsis thaliana Rosettes: Identification by Mass Spectrometry and Bioinformatics. Proteomics, 5, 212-221.
    http://dx.doi.org/10.1002/pmic.200400882

  209. 209. Bayer, E.M., Bottrill, A.R., Walshaw, J., Vigouroux, M., Naldrett, M.J., Thomas, C.L., et al. (2006) Arabidopsis Cell Wall Proteome Defined Using Multidimensional Protein Identification Technology. Proteomics, 6, 301-311.
    http://dx.doi.org/10.1002/pmic.200500046

  210. 210. Minic, Z., Jamet, E., Négroni, L., Der Garabedian, P.A., Zivy, M. and Jouanin, L. (2007) A Sub-Proteome of Arabi- dopsis thaliana Mature Stems Trapped on Concanavalin A Is Enriched in Cell Wall Glycoside Hydrolases. Journal of Experimental Botany, 58, 2503-2512.
    http://dx.doi.org/10.1093/jxb/erm082

  211. 211. Casasoli, M., Spadoni, S., Lilley, K.S., Cervone, F., De Lorenzo, G. and Mattei, B. (2008) Identification by 2-D DIGE of Apoplastic Proteins Regulated by Oligogalacturonides in Arabidopsis thaliana. Proteomics, 8, 1042-1054.
    http://dx.doi.org/10.1002/pmic.200700523

  212. 212. Irshad, M., Canut, H., Borderies, G., Pont-Lezica, R. and Jamet, E. (2008) A New Picture of Cell Wall Protein Dynamics in Elongating Cells of Arabidopsis thaliana: Confirmed Actors and Newcomers. BMC Plant Biology, 8, 94.
    http://dx.doi.org/10.1186/1471-2229-8-94

  213. 213. Gevaert, K. and Vandekerckhove, J. (2000) Protein Identification Methods in Proteomics. Electrophoresis, 21, 1145- 1154.
    http://dx.doi.org/10.1002/(SICI)1522-2683(20000401)21:6<1145::AID-ELPS1145>3.0.CO;2-Z

  214. 214. Aebersold, R. and Mann, M. (2003) Mass Spectrometry-Based Proteomics. Nature, 422, 198-207.
    http://dx.doi.org/10.1038/nature01511

  215. 215. Petersen, J., Rogowska-Wrzesinska, A. and Jensen, O.N. (2013) Functional Proteomics of Barley and Barley Chloro- plasts-Strategies, Methods and Perspectives. Frontiers in Plant Science, 4.
    http://dx.doi.org/10.3389/fpls.2013.00052

  216. 216. Seidler, J., Zinn, N., Boehm, M.E. and Lehmann, W.D. (2010) De novo Sequencing of Peptides by MS/MS. Proteo- mics, 10, 634-649.
    http://dx.doi.org/10.1002/pmic.200900459

  217. 217. Bond, M.R. and Kohler, J.J. (2007) Chemical Methods for Glycoprotein Discovery. Current Opinion in Chemical Biology, 11, 52-58.
    http://dx.doi.org/10.1016/j.cbpa.2006.11.032

  218. 218. Schmidt, A., Kellermann, J. and Lottspeich, F. (2005) A Novel Strategy for Quantitative Proteomics Using Isotope-Coded Protein Labels. Proteomics, 5, 4-15.
    http://dx.doi.org/10.1002/pmic.200400873

  219. 219. Ross, P.L., Huang, Y.N., Marchese, J.N., Williamson, B., Parker, K., Hattan, S., et al. (2004) Multiplexed Protein Quantitation in Saccharomyces cerevisiae Using Amine-Reactive Isobaric Tagging Reagents. Molecular & Cellular Proteomics, 3, 1154-1169.
    http://dx.doi.org/10.1074/mcp.M400129-MCP200

  220. 220. Wiese, S., Reidegeld, K.A., Meyer, H.E. and Warscheid, B. (2007) Protein Labeling by iTRAQ: A New Tool for Quantitative Mass Spectrometry in Proteome Research. Proteomics, 7, 340-350.
    http://dx.doi.org/10.1002/pmic.200600422

  221. 221. Ong, S.-E., Blagoev, B., Kratchmarova, I., Kristensen, D.B., Steen, H., Pandey, A., et al. (2002) Stable Isotope Label- ing by Amino Acids in Cell Culture, SILAC, as a Simple and Accurate Approach to Expression Proteomics. Molecular & Cellular Proteomics, 1, 376-386.
    http://dx.doi.org/10.1074/mcp.M200025-MCP200

  222. 222. Bindschedler, L.V., Palmblad, M. and Cramer, R. (2008) Hydroponic Isotope Labelling of Entire Plants (HILEP) for Quantitative Plant Proteomics; an Oxidative Stress Case Study. Phytochemistry, 69, 1962-1972.
    http://dx.doi.org/10.1016/j.phytochem.2008.04.007

  223. 223. Gouw, J.W., Tops, B.B., Mortensen, P., Heck, A.J. and Krijgsveld, J. (2008) Optimizing Identification and Quantitation of 15N-Labeled Proteins in Comparative Proteomics. Analytical Chemistry, 80, 7796-803.
    http://dx.doi.org/10.1021/ac801249v

  224. 224. Reinhold, B.B., Chan, S.Y., Reuber, T.L., Marra, A., Walker, G.C. and Reinhold, V.N. (1994) Detailed Structural Characterization of Succinoglycan, the Major Exopolysaccharide of Rhizobium Meliloti Rm1021. Journal of Bacteriology, 176, 1997-2002.

  225. 225. Reinhold, B.B., Hauer, C.R., Plummer, T.H. and Reinhold, V.N. (1995) Detailed Structural Analysis of a Novel, Specific O-Linked Glycan from the Prokaryote Flavobacterium meningosepticum. Journal of Biological Chemistry, 270, 13197-13203.
    http://dx.doi.org/10.1074/jbc.270.22.13197

  226. 226. Reinhold, V.N., Reinhold, B.B. and Chan, S. (1996) Carbohydrate Sequence Analysis by Electrospray Ionization-Mass Spectrometry. Methods in Enzymology, 271, 377-402.
    http://dx.doi.org/10.1016/S0076-6879(96)71018-2

  227. 227. Todd, J.F. and March, R.E. (1999) A Retrospective Review of the Development and Application of the Quadrupole Ion Trap Prior to the Appearance of Commercial Instruments. International Journal of Mass Spectrometry, 190, 9-35.
    http://dx.doi.org/10.1016/S1387-3806(99)00065-2

  228. 228. Park, Y. and Lebrilla, C.B. (2005) Application of Fourier Transform Ion Cyclotron Resonance Mass Spectrometry to Oligosaccharides. Mass Spectrometry Reviews, 24, 232-64.
    http://dx.doi.org/10.1002/mas.20010

  229. 229. Lerouxel, O., Choo, T.S., Séveno, M., Usadel, B., Faye, L., Lerouge, P., et al. (2002) Rapid Structural Phenotyping of Plant Cell Wall Mutants by Enzymatic Oligosaccharide Fingerprinting. Plant Physiology, 130, 1754-1763.
    http://dx.doi.org/10.1104/pp.011965

  230. 230. Obel, N., Erben, V. and Pauly, M. (2006) Functional Wall Glycomics through Oligosaccharide Mass Profiling. The Science and Lore of the Plant Cell Wall Brown Walker Press, Boca Raton, 258-266.

  231. 231. Obel, N., Erben, V., Schwarz, T., Kuhnel, S., Fodor, A. and Pauly, M. (2009) Microanalysis of Plant Cell Wall Polysaccharides. Molecular Plant, 2, 922-932.
    http://dx.doi.org/10.1093/mp/ssp046

  232. 232. Reale, S., Di Tullio, A., Spreti, N. and De Angelis, F. (2004) Mass Spectrometry in the Biosynthetic and Structural Investigation of Lignins. Mass Spectrometry Reviews, 23, 87-126.
    http://dx.doi.org/10.1080/02773818308085170

  233. 233. Obst, J.R. (1983) Analytical Pyrolysis of Hardwood and Softwood Lignins and Its Use in Lignin-Type Determination of Hardwood Vessel Elements. Journal of Wood Chemistry and Technology, 3, 377-397.
    http://dx.doi.org/10.1080/02773818308085170

  234. 234. Meier, D. and Faix, O. (1992) Pyrolysis-Gas Chromatography-Mass Spectrometry. Springer, Berlin Heidelberg, 177- 199.
    http://dx.doi.org/10.1007/978-3-642-74065-7_13

  235. 235. Galletti, G.C., Bocchini, P., Smacchia, A.M. and Reeves III, J.B. (1996) Monitoring Phenolic Composition of Maturing Maize Stover by High Performance Liquid Chromatography and Pyrolysis/Gas Chromatography/Mass Spectrometry. Journal of the Science of Food and Agriculture, 71, 1-9.
    http://dx.doi.org/10.1002/(SICI)1097-0010(199605)71:1<1::AID-JSFA535>3.0.CO;2-A

  236. 236. Freudenberg, K. and Lautsch, W. (1939) Zur Konstitution des Fichtenlignins. Naturwissenschaften, 27, 227-228.
    http://dx.doi.org/10.1007/BF02716492

  237. 237. Freudenberg, K., Lautsch, W. and Engler, K. (1940) Die bildung von vanillin aus fichtenlignin. Berichte Der Deutschen Chemischen Gesellschaft (A and B Series), 73, 167-171.
    http://dx.doi.org/10.1002/cber.19400730302

  238. 238. Chen, C.-L. (1992) Nitrobenzene and Cupric Oxide Oxidations. Springer, Berlin Heidelberg, 301-321.
    http://dx.doi.org/10.1007/978-3-642-74065-7_21

  239. 239. Hedges, J.I. and Mann, D.C. (1979) The Characterization of Plant Tissues by Their Lignin Oxidation Products. Geochimica et Cosmochimica Acta, 43, 1803-1807.
    http://dx.doi.org/10.1016/0016-7037(79)90028-0

  240. 240. Freudenberg, K. and Müller, H.F. (1938) Quecksilber und Jod enthaltende Derivate des Fichtenlignins. Berichte Der Deutschen Chemischen Gesellschaft (A and B Series), 71, 2500-2504.
    http://dx.doi.org/10.1002/cber.19380711215

  241. 241. Hyatt, J.A. (1989) Hydroxypropyl Lignins and Model Compounds: Synthesis and Characterization by Electron-Impact Mass Spectrometry. ACS Symposium Series, Oxford University Press, 425-435.
    http://dx.doi.org/10.1021/bk-1989-0397.ch033

  242. 242. Lapierre, C., Monties, B., Rolando, C. and de Chirale, L. (1985) Thioacidolysis of Lignin: Comparison with Acidolysis. Journal of Wood Chemistry and Technology, 5, 277-292.
    http://dx.doi.org/10.1080/02773818508085193

  243. 243. Lapierre, C., Rolando, C. and Monties, B. (1983) Characterization of Poplar Lignins Acidolysis Products: Capillary Gas-Liquid and Liquid-Liquid Chromatography of Monomeric Compounds. Holzforschung-International Journal of the Biology, Chemistry, Physics and Technology of Wood, 37, 189-198.
    http://dx.doi.org/10.1515/hfsg.1983.37.4.189

  244. 244. Sarkanen, K.V., Islam, A. and Anderson, C.D. (1992) Ozonation. Springer, Berlin Heidelberg, 387-406.
    http://dx.doi.org/10.1007/978-3-642-74065-7_26

  245. 245. Lu, F. and Ralph, J. (1997) Derivatization Followed by Reductive Cleavage (DFRC Method), a New Method for Lignin Analysis: Protocol for Analysis of DFRC Monomers. Journal of Agricultural and Food Chemistry, 45, 2590- 2592.
    http://dx.doi.org/10.1021/jf970258h

  246. 246. Clifford, D.J., Carson, D.M., McKinney, D.E., Bortiatynski, J.M. and Hatcher, P.G. (1995) A New Rapid Technique for the Characterization of Lignin in Vascular Plants: Thermochemolysis with Tetramethylammonium Hydroxide (TMAH). Organic Geochemistry, 23, 169-175.
    http://dx.doi.org/10.1016/0146-6380(94)00109-E

  247. 247. De Angelis, F., Fregonese, P. and Verì, F. (1996) Structural Investigation of Synthetic Lignins by Matrix-Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry. Rapid Communications in Mass Spectrometry, 10, 1304-1308.
    http://dx.doi.org/10.1002/(SICI)1097-0231(19960731)10:10<1304::AID-RCM591>3.0.CO;2-0

  248. 248. Evtuguin, D.V., Domingues, P., Amado, F.L., Neto, C.P. and Correia, A.J. (1999) Electrospray Ionization Mass Spectrometry as a Tool for Lignins Molecular Weight and Structural Characterisation. Holzforschung, 53, 525-528.
    http://dx.doi.org/10.1515/HF.1999.086

  249. 249. Palmblad, M., Gellerstedt, G., et al. (2003) Investigation of Lignin Oligomers Using Electrospray Ionisation Mass Spectrometry. Holzforschung, 57, 37-43.

  250. 250. Metzger, J.O., Bicke, C., Faix, O., Tuszynski, W., Angermann, R., Karas, M., et al. (1992) Matrix-Assisted Laser Desorption Mass Spectrometry of Lignins. Angewandte Chemie International Edition in English, 31, 762-764.
    http://dx.doi.org/10.1002/anie.199207621

  251. 251. Bocchini, P., Galletti, G.C., Seraglia, R., Traldi, P., Camarero, S. and Martinez, A.T. (1996) Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry of Natural and Synthetic Lignin. Rapid Communications in Mass Spectrometry, 10, 1144-1147.

  252. 252. Kim, H., Ralph, J. and Akiyama, T. (2008) Solution-State 2D NMR of Ball-Milled Plant Cell Wall Gels in DMSO-d 6. BioEnergy Research, 1, 56-66.

  253. 253. Davis, E.A., Derouet, C., Herve Du Penhoat, C. and Morvan, C. (1990) Isolation and an N.M.R. Study of Pectins from Flax (Linum usitatissimum L.). Carbohydrate Research, 197, 205-215.
    http://dx.doi.org/10.1016/0008-6215(90)84143-I

  254. 254. Newman, R.H., Ha, M.-A. and Melton, L.D. (1994) Solid-State 13C NMR Investigation of Molecular Ordering in the Cellulose of Apple Cell Walls. Journal of Agricultural and Food Chemistry, 42, 1402-1406.
    http://dx.doi.org/10.1021/jf00043a002

  255. 255. Foster, T.J. and Ablett, S. (1996) Mobility-Resolved I3C-NMR. Biopolymers, 39, 1-66.

  256. 256. Duus, J.O., Gotfredsen, C.H. and Bock, K. (2000) Carbohydrate Structural Determination by NMR Spectroscopy: Modern Methods and Limitations. Chemical Reviews, 100, 4589-4614.
    http://dx.doi.org/10.1021/cr990302n

  257. 257. Jarvis, M.C. and McCann, M.C. (2000) Macromolecular Biophysics of the Plant Cell Wall: Concepts and Methodology. Plant Physiology and Biochemistry, 38, 1-13.
    http://dx.doi.org/10.1016/S0981-9428(00)00172-8

  258. 258. Gurjanov, O.P., Ibragimova, N.N., Gnezdilov, O.I. and Gorshkova, T.A. (2008) Polysaccharides, Tightly Bound to Cellulose in Cell Wall of Flax Bast Fibre: Isolation and Identification. Carbohydrate Polymers, 72, 719-729.
    http://dx.doi.org/10.1016/j.carbpol.2007.10.017

  259. 259. Dick-Pérez, M., Zhang, Y., Hayes, J., Salazar, A., Zabotina, O.A. and Hong, M. (2011) Structure and Interactions of Plant Cell-Wall Polysaccharides by Two- and Three-Dimensional Magic-Angle-Spinning Solid-State NMR. Biochemistry, 50, 989-1000.
    http://dx.doi.org/10.1021/bi101795q

  260. 260. Hedenstrom, M., Wiklund-Lindstrom, S., Oman, T., Lu, F., Gerber, L., Schatz, P., et al. (2009) Identification of Lignin and Polysaccharide Modifications in Populus Wood by Chemometric Analysis of 2D NMR Spectra from Dissolved Cell Walls. Molecular Plant, 2, 933-942.
    http://dx.doi.org/10.1093/mp/ssp047

  261. 261. Hall, M., Bansal, P., Lee, J.H., Realff, M.J. and Bommarius, A.S. (2010) Cellulose Crystallinity—A Key Predictor of the Enzymatic Hydrolysis rate. FEBS Journal, 277, 1571-1582.
    http://dx.doi.org/10.1111/j.1742-4658.2010.07585.x

  262. 262. Newman, R.H. (2005) Homogeneity in Cellulose Crystallinity between Samples of Pinus radiata Wood. Holzfors- chung, 58, 91-96.

  263. 263. Park, S., Baker, J.O., Himmel, M.E., Parilla, P.A. and Johnson, D.K. (2010) Research Cellulose Crystallinity Index: Measurement Techniques and Their Impact on Interpreting Cellulase Performance. Biotechnol Biofuels, 3.

  264. 264. Sathitsuksanoh, N., Zhu, Z., Wi, S. and Zhang, Y.-H.P. (2011) Cellulose Solvent-Based Biomass Pretreatment Breaks Highly Ordered Hydrogen Bonds in Cellulose Fibers of Switchgrass. Biotechnology and Bioengineering, 108, 521-529.
    http://dx.doi.org/10.1002/bit.22964

  265. 265. Lu, F. and Ralph, J. (2003) Non-Degradative Dissolution and Acetylation of Ball-Milled Plant Cell Walls: High-Resolution Solution-State NMR. The Plant Journal, 35, 535-544.
    http://dx.doi.org/10.1046/j.1365-313X.2003.01817.x

  266. 266. Ralph, J. and Lu, F. (2004) Cryoprobe 3D NMR of Acetylated Ball-Milled Pine Cell Walls. Organic & Biomolecular Chemistry, 2, 2714-2715.
    http://dx.doi.org/10.1039/b412633e

  267. 267. Moulthrop, J.S., Swatloski, R.P., Moyna, G. and Rogers, R.D. (2005) High-Resolution 13C NMR Studies of Cellulose and Cellulose Oligomers in Ionic Liquid Solutions. Chemical Communications, 1557-1559.
    http://dx.doi.org/10.1039/b417745b

  268. 268. Pu, Y., Jiang, N. and Ragauskas, A.J. (2007) Ionic Liquid as a Green Solvent for Lignin. Journal of Wood Chemistry and Technology, 27, 23-33.

  269. 269. Yelle, D.J., Ralph, J. and Frihart, C.R. (2008) Characterization of Nonderivatized Plant Cell Walls Using High-Resolution Solution-State NMR Spectroscopy. Magnetic Resonance in Chemistry, 46, 508-517.
    http://dx.doi.org/10.1002/mrc.2201

  270. 270. Jiang, N., Pu, Y., Samuel, R. and Ragauskas, A.J. (2009) Perdeuterated Pyridinium Molten salt (Ionic Liquid) for Direct Dissolution and NMR Analysis of Plant Cell Walls. Green Chemistry, 11, 1762-1766.
    http://dx.doi.org/10.1039/b913609f

  271. 271. Kim, H. and Ralph, J. (2010) Solution-State 2D NMR of Ball-Milled Plant Cell Wall Gels in DMSO-d6/Pyridine-d5. Organic & Biomolecular Chemistry, 8, 576-591.
    http://dx.doi.org/10.1039/B916070A

  272. 272. Mansfield, S.D., Kim, H., Lu, F. and Ralph, J. (2012) Whole Plant Cell Wall Characterization Using Solution-State 2D NMR. Nature Protocols, 7, 1579-1589.
    http://dx.doi.org/10.1038/nprot.2012.064

  273. 273. Cheng, K., Sorek, H., Zimmermann, H., Wemmer, D.E. and Pauly, M. (2013) Solution-State 2D NMR Spectroscopy of Plant Cell Walls Enabled by a Dimethylsulfoxide-d 6/1-Ethyl-3-Methylimidazolium Acetate Solvent. Analytical Che- mistry, 85, 3213-3221.
    http://dx.doi.org/10.1021/ac303529v

  274. 274. Lupoi, J.S., Singh, S., Simmons, B.A. and Henry, R.J. (2014) Assessment of Lignocellulosic Biomass Using Analytical Spectroscopy: An Evolution to High-Throughput Techniques. BioEnergy Research, 7, 1-23.
    http://dx.doi.org/10.1007/s12155-013-9352-1

  275. 275. Wen, J.-L., Sun, S.-L., Xue, B.-L. and Sun, R.-C. (2013) Recent Advances in Characterization of Lignin Polymer by Solution-State Nuclear Magnetic Resonance (NMR) Methodology. Materials, 6, 359-391.
    http://dx.doi.org/10.3390/ma6010359

  276. 276. Pinto, P.C., Evtuguin, D.V. and Pascoal Neto, C. (2005) Chemical Composition and Structural Features of the Macro- molecular Components of Plantation Acacia Mangium Wood. Journal of Agricultural and Food Chemistry, 53, 7856- 7862.
    http://dx.doi.org/10.1021/jf058081b

  277. 277. Yan, J., Hu, Z., Pu, Y., Brummer, E.C. and Ragauskas, A.J. (2010) Chemical Compositions of Four Switchgrass Populations. Biomass and Bioenergy, 34, 48-53.
    http://dx.doi.org/10.1016/j.biombioe.2009.09.010

  278. 278. Capanema, E.A., Balakshin, M.Y. and Kadla, J.F. (2004) A Comprehensive Approach for Quantitative Lignin Characterization by NMR Spectroscopy. Journal of Agricultural and Food Chemistry, 52, 1850-1860.
    http://dx.doi.org/10.1021/jf035282b

  279. 279. Bunzel, M. and Ralph, J. (2006) NMR Characterization of Lignins Isolated from Fruit and Vegetable Insoluble Dietary Fiber. Journal of Agricultural and Food Chemistry, 54, 8352-8361.
    http://dx.doi.org/10.1021/jf061525z

  280. 280. Jamet, E., Albenne, C., Boudart, G., Irshad, M., Canut, H. and Pont-Lezica, R. (2008) Recent Advances in Plant Cell Wall Proteomics. Proteomics, 8, 893-908.
    http://dx.doi.org/10.1002/pmic.200700938

  281. 281. Galvani, M., Hamdan, M., Herbert, B. and Righetti, P.G. (2001) Alkylation Kinetics of Proteins in Preparation for Two-Dimensional Maps: A Matrix Assisted Laser Desorption/Ionization-Mass Spectrometry Investigation. Electrophoresis, 22, 2058-2065.
    http://dx.doi.org/10.1002/1522-2683(200106)22:10<2058::AID-ELPS2058>3.0.CO;2-Z

  282. 282. Mineki, R., Taka, H., Fujimura, T., Kikkawa, M., Shindo, N. and Murayama, K. (2002) In Situ Alkylation with Acrylamide for Identification of Cysteinyl Residues in Proteins during One-and Two-Dimensional Sodium Dodecyl Sulphate-Polyacrylamide Gel Electrophoresis. Proteomics, 2, 1672-1681.
    http://dx.doi.org/10.1002/pmic.200300589

  283. 283. Luche, S., Diemer, H., Tastet, C., Chevallet, M., Van Dorsselaer, A., Leize-Wagner, E., et al. (2004) About Thiol Derivatization and Resolution of Basic Proteins in Two-Dimensional Electrophoresis. Proteomics, 4, 551.
    http://dx.doi.org/10.1002/pmic.200300589

  284. 284. Olsson, I., Larsson, K., Palmgren, R. and Bjellqvist, B. (2002) Organic Disulfides as a Means to Generate Streak-Free Two-Dimensional Maps with Narrow Range Basic Immobilized pH Gradient Strips as First Dimension. Proteomics, 2, 1630-1632.
    http://dx.doi.org/10.1002/1615-9861(200211)2:11<1630::AID-PROT1630>3.0.CO;2-N

  285. 285. Jackson, P. (1990) The Use of Polyacrylamide-Gel Electrophoresis for the High-Resolution Separation of Reducing Saccharides Labelled with the Fluorophore 8-Aminonaphthalene-1, 3, 6-Trisulphonic Acid. Detection of Picomolar Quantities by an Imaging System Based on a Cooled Charge-Coupled Device. Biochemical Journal, 270, 705-713.
    http://dx.doi.org/10.1042/bj2700705

  286. 286. Stack, R.J. and Sullivan, M.T. (1992) Electrophoretic Resolution and Fluorescence Detection of N-Linked Glyco- protein Oligosaccharides after Reductive Amination with 8-Aminonaphthalene-1, 3, 6-Trisulphonic Acid. Glycobio- logy, 2, 85-92.
    http://dx.doi.org/10.1093/glycob/2.1.85

  287. 287. Bardor, M., Cabanes-Macheteau, M., Faye, L. and Lerouge, P. (2000) Monitoring the N-Glycosylation of Plant Glyco- proteins by Fluorophore-Assisted Carbohydrate Electrophoresis. Electrophoresis, 21, 2550-2556.
    http://dx.doi.org/10.1002/1522-2683(20000701)21:12<2550::AID-ELPS2550>3.0.CO;2-G

  288. 288. Mort, A.J. and Chen, E.M.W. (1996) Separation of 8-Aminonaphthalene-1, 3, 6-Trisulfonate (ANTS)-Labeled Oligomers Containing Galacturonic Acid by Capillary Electrophoresis: Application to Determining the Substrate Specificity of Endopolygalacturonases. Electrophoresis, 17, 379-383.
    http://dx.doi.org/10.1002/elps.1150170215

  289. 289. Goubet, F., Jackson, P., Deery, M.J. and Dupree, P. (2002) Polysaccharide Analysis Using Carbohydrate Gel Electro- phoresis: A Method to Study Plant Cell Wall Polysaccharides and Polysaccharide Hydrolases. Analytical Biochemistry, 300, 53-68.
    http://dx.doi.org/10.1006/abio.2001.5444

  290. 290. Starr, C.M., Masada, R.I., Hague, C., Skop, E. and Klock, J.C. (1996) Fluorophore-Assisted Carbohydrate Electropho- resis in the Separation, Analysis, and Sequencing of Carbohydrates. Journal of Chromatography A, 720, 295-321.
    http://dx.doi.org/10.1016/0021-9673(95)00749-0

  291. 291. O’Shea, M.G., Samuel, M.S., Konik, C.M. and Morell, M.K. (1998) Fluorophore-Assisted Carbohydrate Electrophoresis (FACE) of Oligosaccharides: Efficiency of Labelling and High-Resolution Separation. Carbohydrate Research, 307, 1-12.
    http://dx.doi.org/10.1016/S0008-6215(97)10085-4

  292. 292. Volpi, N., Maccari, F. and Linhardt, R.J. (2008) Capillary Electrophoresis of Complex Natural Polysaccharides. Electrophoresis, 29, 3095-3106.
    http://dx.doi.org/10.1002/elps.200800109

  293. 293. Campa, C., Coslovi, A., Flamigni, A. and Rossi, M. (2006) Overview on Advances in Capillary Electrophoresis-Mass Spectrometry of Carbohydrates: A Tabulated Review. Electrophoresis, 27, 2027-2050.
    http://dx.doi.org/10.1002/elps.200500960

  294. 294. Amon, S., Zamfir, A.D. and Rizzi, A. (2008) Glycosylation Analysis of Glycoproteins and Proteoglycans Using Capillary Electrophoresis-Mass Spectrometry Strategies. Electrophoresis, 29, 2485-2507.
    http://dx.doi.org/10.1002/elps.200800105

  295. 295. Mechref, Y. and Novotny, M.V. (2009) Glycomic Analysis by Capillary Electrophoresis-Mass Spectrometry. Mass Spectrometry Reviews, 28, 207-222.
    http://dx.doi.org/10.1002/mas.20196

  296. 296. Zaia, J. (2013) Capillary Electrophoresis-Mass Spectrometry of Carbohydrates. Capillary Electrophoresis of Biomole- cules, 984, 13-25.
    http://dx.doi.org/10.1007/978-1-62703-296-4_2

  297. 297. Masselter, S., Zemann, A. and Bobleter, O. (1995) Analysis of Lignin Degradation Products by Capillary Electrophoresis. Chromatographia, 40, 51-57.
    http://dx.doi.org/10.1007/BF02274608

  298. 298. Bonn, G.K., Pfeifer, P.A., Hormeyer, H. and Bobleter, O. (1984) Analysis of Acidic and Phenolic Biomass Degradation Products by Isotachophoresis. Fresenius’ Zeitschrift Für Analytische Chemie, 318, 30-32.
    http://dx.doi.org/10.1007/BF00532834

  299. 299. Lee, K.J., Jung, J.-H., Lee, J.M., So, Y., Kwon, O., Callewaert, N., et al. (2009) High-Throughput Quantitative Analysis of Plant N-Glycan Using a DNA Sequencer. Biochemical and Biophysical Research Communications, 380, 223-229.
    http://dx.doi.org/10.1016/j.bbrc.2009.01.070

  300. 300. SOUPI, M., Bourven, I., Simon, S., Lhernould, S., Omokolo, D., Guibaud, G., et al. (2014) SEC Coupled with UV and Fluorescence Detection, an Efficient Method for β-Glucosyl-Yariv Arabinogalactan Protein (AGP) Monitoring. International Journal of Research In Agriculture and Food Science, 2, 5-15.

  301. 301. Edge, A. (2003) Deglycosylation of Glycoproteins with Trifluoromethanesulphonic Acid: Elucidation of Molecular Structure and Function. Biochemical Journal, 376, 339-350.
    http://dx.doi.org/10.1042/bj20030673

  302. 302. Doco, T., O’neill, M.A. and Pellerin, P. (2001) Determination of the Neutral and Acidic Glycosyl-Residue Composi- tions of Plant Polysaccharides by GC-EI-MS Analysis of the Trimethylsilyl Methyl Glycoside Derivatives. Carbohy- drate Polymers, 46, 249-259.
    http://dx.doi.org/10.1016/S0144-8617(00)00328-3

  303. 303. Chaplin, M.F. (1982) A Rapid and Sensitive Method for the Analysis of Carbohydrate Components in Glycoproteins Using Gas-Liquid Chromatography. Analytical Biochemistry, 123, 336-341.
    http://dx.doi.org/10.1016/0003-2697(82)90455-9

  304. 304. Willfor, S., Pranovich, A., Tamminen, T., Puls, J., Laine, C., Suurnakki, A., et al. (2009) Carbohydrate Analysis of Plant Materials with Uronic Acid-Containing Polysaccharides—A Comparison between Different Hydrolysis and Subsequent Chromatographic Analytical Techniques. Industrial Crops and Products, 29, 571-580.
    http://dx.doi.org/10.1016/j.indcrop.2008.11.003

  305. 305. Blakeney, A.B., Harris, P.J., Henry, R.J. and Stone, B.A. (1983) A Simple and Rapid Preparation of Alditol Acetates for Monosaccharide Analysis. Carbohydrate Research, 113, 291-299.
    http://dx.doi.org/10.1016/0008-6215(83)88244-5

  306. 306. Chambers, R.E. and Clamp, J.R. (1971) An Assessment of Methanolysis and Other Factors Used in the Analysis of Carbohydrate-Containing Materials. Biochemical Journal, 125, 1009-1018.
    http://dx.doi.org/10.1042/bj1251009

  307. 307. Bleton, J., Mejanelle, P., Sansoulet, J., Goursaud, S. and Tchapla, A. (1996) Characterization of Neutral Sugars and Uronic Acids after Methanolysis and Trimethylsilylation for Recognition of Plant Gums. Journal of Chromatography A, 720, 27-49.
    http://dx.doi.org/10.1016/0021-9673(95)00308-8

  308. 308. Biermann, C.J. (1989) Introduction to Analysis of Carbohydrates by Gas-Liquid Chromatography (GLC). W: Analysis of Carbohydrates by GLC and MS, Red: Biermann CJ, McGinnis GD CRC Press Inc, Florida, 1-18.

  309. 309. Englmaier, P. (1989) Carbohydrate Trifiuoroacetates. In: Biermann, C.J. and McGinnis, G.D., Eds., Analysis of Carbohydrates by GLC and MS, CRC Press, Inc., Boca Raton, FL, 127-141.

  310. 310. Black, G.E. and Fox, A. (1996) Recent Progress in the Analysis of Sugar Monomers from Complex Matrices Using Chromatography in Conjunction with Mass Spectrometry or Stand-Alone Tandem Mass Spectrometry. Journal of Chromatography A, 720, 51-60.
    http://dx.doi.org/10.1016/0021-9673(95)00335-5

  311. 311. McGinnis, G.D. and Biermann, C.J. (1989) Analysis of Monosaccharides as Per-O-Acetylated Aldononitrile (PAAN) Derivatives by Gas-Liquid Chromatography (GLC). Analysis of Carbohydrates by GLC and MS, CRC Press, Inc., Boca Raton, Florida, 119-125.

  312. 312. Neeser, J.-R. and Schweizer, T.F. (1989) Analysis of Carbohydrates as 0-Alkyloxime Derivatives by Gas-Liquid Chromatography (GLC). In: Biermann, C.J. and McGinnis, G.D., Eds., Analysis of Carbohydrates by GLC and MS, CRC Press, Inc., Boca Raton, 143-155.

  313. 313. Chen, W., Smeekens, J.M. and Wu, R. (2014) Comprehensive Analysis of Protein N-Glycosylation Sites by Combining Chemical Deglycosylation with LC-MS. Journal of Proteome Research, 13, 1466-1473.
    http://dx.doi.org/10.1021/pr401000c

  314. 314. Ruiz-May, E., Thannhauser, T.W., Zhang, S. and Rose, J.K. (2012) Analytical Technologies for Identification and Characterization of the Plant N-Glycoproteome. Frontiers in Plant Science, 3.
    http://dx.doi.org/10.3389/fpls.2012.00150

  315. 315. Pauly, M., Albersheim, P., Darvill, A. and York, W.S. (1999) Molecular Domains of the Cellulose/Xyloglucan Network in the Cell Walls of Higher Plants. The Plant Journal, 20, 629-639.
    http://dx.doi.org/10.1046/j.1365-313X.1999.00630.x

  316. 316. Zhang, Z., Xiao, Z. and Linhardt, R.J. (2009) Thin Layer Chromatography for the Separation and Analysis of Acidic Carbohydrates. Journal of Liquid Chromatography & Related Technologies, 32, 1711-1732.
    http://dx.doi.org/10.1080/10826070902956402

  317. 317. Hartley, R.D. (1971) Improved Methods for the Estimation by Gas-Liquid Chromatography of Lignin Degradation Products from Plants. Journal of Chromatography A, 54, 335-344.
    http://dx.doi.org/10.1016/S0021-9673(01)80289-2

  318. 318. Hedges, J.I. and Ertel, J.R. (1982) Characterization of Lignin by Gas Capillary Chromatography of Cupric Oxide Oxidation Products. Analytical Chemistry, 54, 174-178.
    http://dx.doi.org/10.1021/ac00239a007

  319. 319. Heitner, C., Dimmel, D. and Schmidt, J. (2010) Lignin and Lignans: Advances in Chemistry [Internet]. CRC Press.
    http://dx.doi.org/10.1201/EBK1574444865

  320. 320. Gidh, A.V., Decker, S.R., Vinzant, T.B., Himmel, M.E. and Williford, C. (2006) Determination of Lignin by Size Exclusion Chromatography Using Multi Angle Laser Light Scattering. Journal of Chromatography A, 1114, 102-110.
    http://dx.doi.org/10.1016/j.chroma.2006.02.044

  321. 321. Willats, W.G. and Knox, J.P. (2003) Molecules in Context: Probes for Cell Wall Analysis. In: Rose, J.K.C., Ed., The Plant Cell Wall, Blackwell Publishing/CRC, Oxford, 92-110.

  322. 322. Moller, I., Marcus, S.E., Haeger, A., Verhertbruggen, Y., Verhoef, R., Schols, H., et al. (2008) High-Throughput Screening of Monoclonal Antibodies against Plant Cell Wall Glycans by Hierarchical Clustering of Their Carbohydrate Microarray Binding Profiles. Glycoconjugate Journal, 25, 37-48.
    http://dx.doi.org/10.1007/s10719-007-9059-7

  323. 323. Freshour, G., Clay, R.P., Fuller, M.S., Albersheim, P., Darvill, A.G. and Hahn, M.G. (1996) Developmental and Tissue-Specific Structural Alterations of the Cell-Wall Polysaccharides of Arabidopsis thaliana Roots. Plant Physiology, 110, 1413-1429.

  324. 324. Jones, L., Seymour, G.B. and Knox, J.P. (1997) Localization of Pectic Galactan in Tomato Cell Walls Using a Monoclonal Antibody Specific to (1[->]4)-[Beta]-D-Galactan. Plant Physiology, 113, 1405-1412.

  325. 325. Willats, W.G., Marcus, S.E. and Knox, J.P. (1998) Generation of a Monoclonal Antibody Specific to (1→ 5)-α-l- Arabinan. Carbohydrate Research, 308, 149-152.
    http://dx.doi.org/10.1016/S0008-6215(98)00070-6

  326. 326. McCartney, L., Ormerod, A.P., Gidley, M.J. and Knox, J.P. (2000) Temporal and Spatial Regulation of Pectic (1-->4)- Beta-D-Galactan in Cell Walls of Developing Pea Cotyledons: Implications for Mechanical Properties. The Plant Journal: For Cell and Molecular Biology, 22, 105-113.
    http://dx.doi.org/10.1046/j.1365-313x.2000.00719.x

  327. 327. Clausen, M.H., Ralet, M.-C., Willats, W.G.T., McCartney, L., Marcus, S.E., Thibault, J.-F., et al. (2004) A Mono-clonal Antibody to Feruloylated-(1-->4)-Beta-D-Galactan. Planta, 219, 1036-1041.
    http://dx.doi.org/10.1007/s00425-004-1309-3

  328. 328. Altaner, C., Hapca, A.I., Knox, J.P. and Jarvis, M.C. (2007) Detection of β-1-4-Galactan in Compression Wood of Sitka spruce [Picea sitchensis (Bong.) Carrière] by Immunofluorescence. Holzforschung, 61, 311-316.
    http://dx.doi.org/10.1515/HF.2007.049

  329. 329. Ralet, M.-C., Tranquet, O., Poulain, D., Moise, A. and Guillon, F. (2010) Monoclonal Antibodies to Rhamnogalacturonan I Backbone. Planta, 231, 1373-1383.
    http://dx.doi.org/10.1007/s00425-010-1116-y

  330. 330. Clausen, M.H., Willats, W.G.T. and Knox, J.P. (2003) Synthetic Methyl Hexagalacturonate Hapten Inhibitors of Anti-Homogalacturonan Monoclonal Antibodies LM7, JIM5 and JIM7. Carbohydrate Research, 338, 1797-1800.
    http://dx.doi.org/10.1016/S0008-6215(03)00272-6

  331. 331. Willats, W.G., Orfila, C., Limberg, G., Buchholt, H.C., van Alebeek, G.-J.W., Voragen, A.G., et al. (2001) Modulation of the Degree and Pattern of Methyl-Esterification of Pectic Homogalacturonan in Plant Cell Walls Implications for Pectin Methyl Esterase Action, Matrix Properties, and Cell Adhesion. Journal of Biological Chemistry, 276, 19404- 19413.
    http://dx.doi.org/10.1074/jbc.M011242200

  332. 332. Freshour, G., Bonin, C.P., Reiter, W.-D., Albersheim, P., Darvill, A.G. and Hahn, M.G. (2003) Distribution of Fucose-Containing Xyloglucans in Cell Walls of the Mur1 Mutant of Arabidopsis. Plant Physiology, 131, 1602-1612.
    http://dx.doi.org/10.1104/pp.102.016444

  333. 333. Freshour, G., Clay, R.P., Fuller, M.S., Albersheim, P., Darvill, A.G. and Hahn, M.G. (1996) Developmental and Tissue-Specific Structural Alterations of the Cell-Wall Polysaccharides of Arabidopsis thaliana Roots. Plant Physiology, 110, 1413-1429.

  334. 334. Marcus, S.E., Verhertbruggen, Y., Hervé, C., Ordaz-Ortiz, J.J., Farkas, V., Pedersen, H.L., et al. (2008) Pectic Homo- galacturonan Masks Abundant Sets of Xyloglucan Epitopes in Plant Cell Walls. BMC Plant Biology, 8, 60.
    http://dx.doi.org/10.1186/1471-2229-8-60

  335. 335. Willats, W.G., McCartney, L., Steele-King, C.G., Marcus, S.E., Mort, A., Huisman, M., et al. (2004) A Xylogalacturonan Epitope Is Specifically Associated with Plant Cell Detachment. Planta, 218, 673-681.
    http://dx.doi.org/10.1007/s00425-003-1147-8

  336. 336. McCartney, L. (2005) Monoclonal Antibodies to Plant Cell Wall Xylans and Arabinoxylans. Journal of Histochemistry and Cytochemistry, 53, 543-546.
    http://dx.doi.org/10.1007/BF01280168

  337. 337. Dolan, L., Linstead, P. and Roberts, K. (1995) An AGP Epitope Distinguishes a Central Metaxylem Initial from Other Vascular Initials in the Arabidopsis Root. Protoplasma, 189, 149-155.
    http://dx.doi.org/10.1007/BF01280168

  338. 338. Pennell, R.I., Janniche, L., Kjellbom, P., Scofield, G.N., Peart, J.M. and Roberts, K. (1991) Developmental Regulation of a Plasma Membrane Arabinogalactan Protein Epitope in Oilseed Rape Flowers. The Plant Cell Online, 3, 1317- 13126.
    http://dx.doi.org/10.1105/tpc.3.12.1317

  339. 339. Puhlmann, J., Bucheli, E., Swain, M.J., Dunning, N., Albersheim, P., Darvill, A.G., et al. (1994) Generation of Monoclonal Antibodies against Plant Cell-Wall Polysaccharides (I. Characterization of a Monoclonal Antibody to a Terminal [alpha]-(1->2)-Linked Fucosyl-Containing Epitope. Plant Physiology, 104, 699-710.
    http://dx.doi.org/10.1104/pp.104.2.699

  340. 340. Smallwood, M., Yates, E.A., Willats, W.G.T., Martin, H. and Knox, J.P. (1996) Immunochemical Comparison of Membrane-Associated and Secreted Arabinogalactan-Proteins in Rice and Carrot. Planta, 198, 452-459.
    http://dx.doi.org/10.1007/BF00620063

  341. 341. Smallwood, M., Martin, H. and Knox, J.P. (1995) An Epitope of Rice Threonine- and Hydroxyproline-Rich Glycoprotein Is Common to Cell Wall and Hydrophobic Plasma-Membrane Glycoproteins. Planta, 196, 510-522.
    http://dx.doi.org/10.1007/BF00203651

  342. 342. Pattathil, S., Avci, U., Baldwin, D., Swennes, A.G., McGill, J.A., Popper, Z., et al. (2010) A Comprehensive Toolkit of Plant Cell Wall Glycan-Directed Monoclonal Antibodies. Plant Physiology, 153, 514-525.
    http://dx.doi.org/10.1104/pp.109.151985

  343. 343. Pennell, R.I. and Roberts, K. (1995) Chapter 9 Monoclonal Antibodies to Cell-Specific Cell Surface Carbohydrates in Plant Cell Biology and Development. Methods in Cell Biology, 49, 123-141.
    http://dx.doi.org/10.1016/s0091-679x(08)61450-8

  344. 344. Boraston, A.B., Bolam, D., Gilbert, H. and Davies, G. (2004) Carbohydrate-Binding Modules: Fine-Tuning Polysaccharide Recognition. Biochemical Journal, 382, 769-781.
    http://dx.doi.org/10.1042/BJ20040892

  345. 345. Hilden, L. and Johansson, G. (2004) Recent Developments on Cellulases and Carbohydrate-Binding Modules with Cellulose Affinity. Biotechnology Letters, 26, 1683-1693.
    http://dx.doi.org/10.1007/s10529-004-4579-8

  346. 346. Shoseyov, O., Shani, Z. and Levy, I. (2006) Carbohydrate Binding Modules: Biochemical Properties and Novel Applications. Microbiology and Molecular Biology Reviews, 70, 283-295.
    http://dx.doi.org/10.1074/jbc.M605903200

  347. 347. Blake, A.W., McCartney, L., Flint, J.E., Bolam, D.N., Boraston, A.B., Gilbert, H.J., et al. (2006) Understanding the Biological Rationale for the Diversity of Cellulose-Directed Carbohydrate-Binding Modules in Prokaryotic Enzymes. Journal of Biological Chemistry, 281, 29321-29329.
    http://dx.doi.org/10.1074/jbc.M605903200

  348. 348. Ding, S., Xu, Q., Ali, M.K., Baker, J.O., Bayer, E.A., Barak, Y., et al. (2006) Versatile Derivatives of Carbohydrate- Binding Modules for Imaging of Complex Carbohydrates Approaching the Molecular Level of Resolution. Biotechniques, 41, 435.
    http://dx.doi.org/10.2144/000112244

  349. 349. Hildén, L., Daniel, G. and Johansson, G. (2003) Use of a Fluorescence Labelled, Carbohydrate-Binding Module from Phanerochaete chrysosporium Cel7D for Studying Wood Cell Wall Ultrastructure. Biotechnology Letters, 25, 553-558.
    http://dx.doi.org/10.1023/A:1022846304826

  350. 350. McCartney, L., Blake, A.W., Flint, J., Bolam, D.N., Boraston, A.B., Gilbert, H.J., et al. (2006) Differential Recog nition of Plant Cell Walls by Microbial Xylan-Specific Carbohydrate-Binding Modules. Proceedings of the National Academy of Sciences of the United States of America, 103, 4765-4770.
    http://dx.doi.org/10.1073/pnas.0508887103

  351. 351. Filonova, L., Kallas, A.M., Greffe, L., Johansson, G., Teeri, T.T. and Daniel, G. (2007) Analysis of the Surfaces of Wood Tissues and Pulp Fibers Using Carbohydrate-Binding Modules Specific for Crystalline Cellulose and Mannan. Biomacromolecules, 8, 91-97.
    http://dx.doi.org/10.1021/bm060632z

  352. 352. McCartney, L., Gilbert, H.J., Bolam, D.N., Boraston, A.B. and Knox, J.P. (2004) Glycoside Hydrolase Carbohydrate- Binding Modules as Molecular Probes for the Analysis of Plant Cell Wall Polymers. Analytical Biochemistry, 326, 49- 54.
    http://dx.doi.org/10.1016/j.ab.2003.11.011

  353. 353. McCartney, L., Gilbert, H.J., Bolam, D.N., Boraston, A.B. and Knox, J.P. (2004) Glycoside Hydrolase Carbohydrate- Binding Modules as Molecular Probes for the Analysis of Plant Cell Wall Polymers. Analytical Biochemistry, 326, 49-54.
    http://dx.doi.org/10.1016/j.ab.2003.11.011

  354. 354. Knox, J.P. (2008) Revealing the Structural and Functional Diversity of Plant Cell Walls. Current Opinion in Plant Biology, 11, 308-313.
    http://dx.doi.org/10.1016/S0008-6215(98)00070-6

  355. 355. Willats, W.G., Marcus, S.E. and Knox, J.P. (1998) Generation of a Monoclonal Antibody Specific to (1→ 5)-α-l-Arabinan. Carbohydrate Research, 308, 149-152.
    http://dx.doi.org/10.1016/S0008-6215(98)00070-6

  356. 356. Tang, P.W., Gool, H.C., Hardy, M., Lee, Y.C. and Felzi, T. (1985) Novel Approach to the Study of the Antigenicities and Receptor Functions of Carbohydrate Chains of Glycoproteins. Biochemical and Biophysical Research Communications, 132, 474-480.
    http://dx.doi.org/10.1016/0006-291X(85)91158-1

  357. 357. Obel, N., Erben, V., Schwarz, T., Kühnel, S., Fodor, A. and Pauly, M. (2009) Microanalysis of Plant Cell Wall Polysaccharides. Molecular Plant, 2, 922-932.
    http://dx.doi.org/10.1093/mp/ssp046

  358. 358. Moller, I., Sorensen, I., Bernal, A.J., Blaukopf, C., Lee, K., Obro, J., et al. (2007) High-Throughput Mapping of Cell-Wall Polymers within and between Plants Using Novel Microarrays: Glycan Microarrays for Plant Cell-Wall Analysis. The Plant Journal, 50, 1118-1128.
    http://dx.doi.org/10.1111/j.1365-313X.2007.03114.x

  359. 359. Plazanet, I., Zerrouki, R., Lhernould, S., Breton, C. and Costa, G. (2015) Direct Immunological Detection of Wood Cell Wall Polysaccharides after Microwave-Assisted Ionic Liquid Disruption. Glycobiology Journals, 4, 2.

  360. 360. Czaja, A.T. (1934) Untersuchungen über metachromatische Firbungen von Pflanzengeweben. Planta, 21, 531-601.
    http://dx.doi.org/10.1007/BF01909490

  361. 361. Tolivia, D. and Tolivia, J. (1987) Fasga: A New Polychromatic Method for Simultaneous and Differential Staining of Plant Tissues. Journal of Microscopy, 148, 113-117.
    http://dx.doi.org/10.1111/j.1365-2818.1987.tb02859.x

  362. 362. Wiesner, J. (1878) Note fiber das Verhalten des Phloroglucins und einiger verwandter K6rper zur verholzten Zellmembran. Sitzungsber Math Naturwiss CI Akad Wiss Wien, 77, 60-66.

  363. 363. Maule, C. (1901) Das verhalten verholzter membranen gegen kaliumpermanganat, eine holzreaktion neuer art. A. Zimmer’s Verlag (Ernst Mohrmann).

  364. 364. Bond, J., Donaldson, L., Hill, S. and Hitchcock, K. (2008) Safranine Fluorescent Staining of Wood Cell Walls. Bio- technic & Histochemistry, 83, 161-171.
    http://dx.doi.org/10.1080/10520290802373354

  365. 365. Donaldson, L.A. (2002) Abnormal Lignin Distribution in Wood from Severely Drought Stressed Pinus radiata trees. IAWA JL (NS), 23, 161-178.
    http://dx.doi.org/10.1163/22941932-90000295

  366. 366. Donaldson, L.A. and Bond, J. (2005) Fluorescence Microscopy of Wood. New Zealand Forest Research Institute [CD ROM], Rotorua.

  367. 367. Brundrett, M.C., Enstone, D.E. and Peterson, C.A. (1988) A Berberine-Aniline Blue Fluorescent Staining Procedure for Suberin, Lignin, and Callose in Plant Tissue. Protoplasma, 146, 133-142.
    http://dx.doi.org/10.1007/BF01405922

  368. 368. Castellan, A., Trichet, V., Pommier, J.-C., Siohan, A. and Armagnacq, S. (1995) Photo and Thermal Stability of To- tally Chlorine Free Softwood Pulps Studied by UV/V Is Diffuse Reflectance and Fluorescence Spectroscopy. Journal of Pulp and Paper Science, 21, J291–6.

  369. 369. Olmstead, J.A. and Gray, D.G. (1997) Fluorescence Spectroscopy of Cellulose, Lignin and Mechanical Pulps: A Review. Journal of Pulp and Paper Science, 23, J571-J581.

  370. 370. Anderson, C.T., Carroll, A., Akhmetova, L. and Somerville, C. (2010) Real-Time Imaging of Cellulose Reorientation during Cell Wall Expansion in Arabidopsis Roots. Plant Physiology, 152, 787-796.
    http://dx.doi.org/10.1104/pp.109.150128

  371. 371. Anderson, C.T. and Carroll, A. (2014) Identification and Use of Fluorescent Dyes for Plant Cell Wall Imaging Using High-Throughput Screening. Plant Chemical Genomics, 1056, 103-109.
    http://dx.doi.org/10.1007/978-1-62703-592-7_10

  372. 372. Hoch, H.C., Galvani, C.D., Szarowski, D.H. and Turner, J.N. (2005) Two New Fluorescent Dyes Applicable for Visualization of Fungal Cell Walls. Mycologia, 97, 580-588.
    http://dx.doi.org/10.3852/mycologia.97.3.580

  373. 373. Knox, J.P. (1992) Molecular Probes for the Plant Cell Surface. Protoplasma, 167, 1-9.
    http://dx.doi.org/10.1007/BF01353575

  374. 374. Knox, J.P. (2008) Revealing the Structural and Functional Diversity of Plant Cell Walls. Current Opinion in Plant Biology, 11, 308-313.
    http://dx.doi.org/10.1016/j.pbi.2008.03.001

  375. 375. Willats, W.G., Steele-King, C.G., McCartney, L., Orfila, C., Marcus, S.E. and Knox, J.P. (2000) Making and Using Antibody Probes to Study Plant Cell Walls. Plant Physiology and Biochemistry, 38, 27-36.
    http://dx.doi.org/10.1016/S0981-9428(00)00170-4

  376. 376. Boraston, A.B., Bolam, D., Gilbert, H. and Davies, G. (2004) Carbohydrate-Binding Modules: Fine-Tuning Polysaccharide Recognition. Biochemical Journal, 382, 769-781.
    http://dx.doi.org/10.1042/BJ20040892

  377. 377. Donaldson, L.A. and Knox, J.P. (2012) Localization of Cell Wall Polysaccharides in Normal and Compression Wood of Radiata Pine: Relationships with Lignification and Microfibril Orientation. Plant Physiology, 158, 642-653.
    http://dx.doi.org/10.1104/pp.111.184036

  378. 378. Hepler, P.K. and Gunning, B.E. (1998) Confocal Fluorescence Microscopy of Plant Cells. Protoplasma, 201, 121-157.
    http://dx.doi.org/10.1007/BF01287411

  379. 379. Schwille, P., Haupts, U., Maiti, S. and Webb, W.W. (1999) Molecular Dynamics in Living Cells Observed by Fluorescence Correlation Spectroscopy with One- and Two-Photon Excitation. Biophysical Journal, 77, 2251-2265.
    http://dx.doi.org/10.1016/S0006-3495(99)77065-7

  380. 380. Blancaflor, E.B. and Gilroy, S. (2000) Plant Cell Biology in the New Millennium: New Tools and New Insights. American Journal of Botany, 87, 1547-1560.
    http://dx.doi.org/10.2307/2656730

  381. 381. Fricker, M.D. and Meyer, A.J. (2001) Confocal Imaging of Metabolism in Vivo: Pitfalls and Possibilities. Journal of Experimental Botany, 52, 631-640.
    http://dx.doi.org/10.1007/s00709-003-0026-2

  382. 382. Feijó, J.A. and Moreno, N. (2004) Imaging Plant Cells by Two-Photon Excitation. Protoplasma, 223, 1-32.
    http://dx.doi.org/10.1007/s00709-003-0026-2

  383. 383. Tirlapur, U.K. and Konig, K. (1999) Near-Infrared Femtosecond Laser Pulses as a Novel Non-Invasive Means for Dye-Permeation and 3D Imaging of Localised Dye-Coupling in the Arabidopsis Root Meristem. The Plant Journal, 20, 363-370.
    http://dx.doi.org/10.1046/j.1365-313X.1999.t01-1-00603.x

  384. 384. Squirrell, J.M., Wokosin, D.L., White, J.G. and Bavister, B.D. (1999) Long-Term Two-Photon Fluorescence Imaging of Mammalian Embryos without Compromising Viability. Nature Biotechnology, 17, 763-767.
    http://dx.doi.org/10.1038/11698

  385. 385. Tirlapur, U.K. and Konig, K. (2001) Femtosecond Near-Infrared Lasers as a Novel Tool for Non-Invasive Real-Time High-Resolution Time-Lapse Imaging of Chloroplast Division in Living Bundle Sheath Cells of Arabidopsis. Planta, 214, 1-10.
    http://dx.doi.org/10.1007/s004250100597

  386. 386. Tirlapur, U.K. and Konig, K. (2002) Femtosecond Near-Infrared Laser Pulses as a Versatile Non-Invasive Tool for Intra-Tissue Nanoprocessing in Plants without Compromising Viability. The Plant Journal, 31, 365-374.
    http://dx.doi.org/10.1046/j.1365-313X.2002.01346.x

  387. 387. Moreno, N., Bougourd, S., Haseloff, J. and Feijó, J.A. (2006) Imaging Plant Cells. Handbook of Biological Confocal Microscopy, 166, 769-787.
    http://dx.doi.org/10.1104/pp.114.245597

  388. 388. Schuetz, M., Benske, A., Smith, R.A., Watanabe, Y., Tobimatsu, Y., Ralph, J., et al. (2014) Laccases Direct Ligni- fication in the Discrete Secondary Cell Wall Domains of Protoxylem. Plant Physiology, 166, 798-807.
    http://dx.doi.org/10.1104/pp.114.245597

  389. 389. Sosinsky, G.E., Crum, J., Jones, Y.Z., Lanman, J., Smarr, B., Terada, M., et al. (2008) The Combination of Chemical Fixation Procedures with High Pressure Freezing and Freeze Substitution Preserves Highly Labile Tissue Ultra-structure for Electron Tomography Applications. Journal of Structural Biology, 161, 359-371.
    http://dx.doi.org/10.1016/j.jsb.2007.09.002

  390. 390. Griffith, P.R. and De Haseth, J.A. (1986) Fourier Transform Infrared Spectroscopy. Chem Anal Ser Monogr Anal Chem Appl, 83.

  391. 391. McCann, M.C., Hammouri, M., Wilson, R., Belton, P. and Roberts, K. (1992) Fourier Transform Infrared Micro-spectroscopy Is a New Way to Look at Plant Cell Walls. Plant Physiology, 100, 1940-1947.
    http://dx.doi.org/10.1104/pp.100.4.1940

  392. 392. Himmelsbach, D.S. and Akin, D.E. (1998) Near-Infrared Fourier-Transform Raman Spectroscopy of Flax (Linum usitatissimum L.) Stems. Journal of Agricultural and Food Chemistry, 46, 991-998.
    http://dx.doi.org/10.1021/jf970656k

  393. 393. Himmelsbach, D.S., Khalili, S. and Akin, D.E. (2002) The Use of FT-IR Microspectroscopic Mapping to Study the Effects of Enzymatic Retting of Flax (Linum usitatissimum L) Stems. Journal of the Science of Food and Agriculture, 82, 685-696.
    http://dx.doi.org/10.1007/s00226-004-0274-0

  394. 394. Labbe, N., Rials, T.G., Kelley, S.S., Cheng, Z.-M., Kim, J.-Y. and Li, Y. (2005) FT-IR Imaging and Pyrolysis-Molecular Beam Mass Spectrometry: New Tools to Investigate Wood Tissues. Wood Science and Technology, 39, 61-76.
    http://dx.doi.org/10.1007/s00226-004-0274-0

  395. 395. Gierlinger, N., Sapei, L. and Paris, O. (2008) Insights into the Chemical Composition of Equisetum hyemale by High Resolution Raman Imaging. Planta, 227, 969-980.
    http://dx.doi.org/10.1007/s00425-007-0671-3

  396. 396. Yin, C.-Y. and Goh, B.-M. (2011) Thermal Degradation of Rice Husks in Air and Nitrogen: Thermogravimetric and Kinetic Analyses. Energy Sources, Part A: Recovery, Utilization, and Environmental Effects, 34, 246-252.
    http://dx.doi.org/10.1080/15567030903586048

  397. 397. Gou, J.-Y., Park, S., Yu, X.-H., Miller, L.M. and Liu, C.-J. (2008) Compositional Characterization and Imaging of “Wall-Bound” Acylesters of Populus trichocarpa Reveal Differential Accumulation of Acyl Molecules in Normal and Reactive Woods. Planta, 229, 15-24.
    http://dx.doi.org/10.1007/s00425-008-0799-9

  398. 398. Chang, S., Salmén, L., Olsson, A.-M. and Clair, B. (2014) Deposition and Organization of Cell Wall Polymers during Tension Wood Cell Wall Maturation Studied by FTIR Microspectroscopy. Planta, 239, 243-254.

  399. 399. Ji, Z., Ding, D., Ling, Z., Zhang, X., Zhou, X. and Xu, F. (2014) In Situ Microscopic Investigation of Plant Cell Walls Deconstruction in Biorefinery. In: Mendez-Vilas, A.., Ed., Microscopy: Advances in Scientific Research and Education, Formatex Research Center, Spain, 426-433.

  400. 400. Gierlinger, N., Luss, S., Konig, C., Konnerth, J., Eder, M. and Fratzl, P. (2010) Cellulose Microfibril Orientation of Picea abies and Its Variability at the Micron-Level Determined by Raman Imaging. Journal of Experimental Botany, 61, 587-595.
    http://dx.doi.org/10.1093/jxb/erp325

  401. 401. Schmidt, M., Schwartzberg, A.M., Carroll, A., Chaibang, A., Adams, P.D. and Schuck, P.J. (2010) Raman imaging of cell wall polymers in Arabidopsis thaliana. Biochemical and Biophysical Research Communications, 395, 521–3. http://dx.doi.org/10.1016/j.bbrc.2010.04.055

  402. 402. Agarwal, U.P. (2006) Raman Imaging to Investigate Ultrastructure and Composition of Plant Cell Walls: Distribution of lignin and Cellulose in Black Spruce Wood (Picea mariana). Planta, 224, 1141-1153.
    http://dx.doi.org/10.1007/s00425-006-0295-z

  403. 403. Schmidt, M., Schwartzberg, A.M., Perera, P.N., Weber-Bargioni, A., Carroll, A., Sarkar, P., et al. (2009) Label-Free in Situ Imaging of Lignification in the Cell Wall of Low Lignin Transgenic Populus trichocarpa. Planta, 230, 589-597.
    http://dx.doi.org/10.1007/s00425-009-0963-x

  404. 404. Zhang, Z., Ma, J., Ji, Z. and Xu, F. (2012) Comparison of Anatomy and Composition Distribution between Normal and Compression Wood of Pinus bungeana zucc. Revealed by Microscopic Imaging Techniques. Microscopy and Micro-analysis, 18, 1459-1466.
    http://dx.doi.org/10.1017/S1431927612013451

  405. 405. Chu, L.-Q., Masyuko, R., Sweedler, J.V. and Bohn, P.W. (2010) Base-Induced Delignification of Miscanthus x Giganteus Studied by Three-Dimensional Confocal Raman Imaging. Bioresource Technology, 101, 4919-4925.
    http://dx.doi.org/10.1016/j.biortech.2009.10.096

  406. 406. Zhang, X., Ma, J., Ji, Z., Yang, G.-H., Zhou, X. and Xu, F. (2014) Using Confocal Raman Microscopy to Real-Time Monitor Poplar Cell Wall Swelling and Dissolution during Ionic Liquid Pretreatment. Microscopy Research and Technique, 77, 609-618.
    http://dx.doi.org/10.1002/jemt.22379

  407. 407. Evans, C.L. and Xie, X.S. (2008) Coherent Anti-Stokes Raman Scattering Microscopy: Chemical Imaging for Biology and Medicine. Annual Review of Analytical Chemistry, 1, 883-909.
    http://dx.doi.org/10.1146/annurev.anchem.1.031207.112754

  408. 408. Chandra, S., Smith, D.R. and Morrison, G.H. (2000) Peer Reviewed: A Subcellular Imaging by Dynamic SIMS Ion Microscopy. Analytical Chemistry, 72, 104-114.

  409. 409. Guerquin-Kern, J.-L., Wu, T.-D., Quintana, C. and Croisy, A. (2005) Progress in Analytical Imaging of the Cell by Dynamic Secondary Ion Mass Spectrometry (SIMS Microscopy). Biochimica et Biophysica Acta (BBA)-General Subjects, 1724, 228-238.
    http://dx.doi.org/10.1016/j.bbagen.2005.05.013

  410. 410. Smart, K.E., Smith, J.A.C., Kilburn, M.R., Martin, B.G., Hawes, C. and Grovenor, C.R. (2010) High-Resolution Elemental Localization in Vacuolate Plant Cells by Nanoscale Secondary ion Mass Spectrometry. The Plant Journal, 63, 870-879.
    http://dx.doi.org/10.1111/j.1365-313X.2010.04279.x

  411. 411. Moore, K.L., Schroder, M., Wu, Z., Martin, B.G., Hawes, C.R., McGrath, S.P., et al. (2011) High-Resolution Secondary ion Mass Spectrometry Reveals the Contrasting Subcellular Distribution of Arsenic and Silicon in Rice Roots. Plant Physiology, 156, 913-924.
    http://dx.doi.org/10.1104/pp.111.173088

  412. 412. Boughton, B.A., Thinagaran, D., Sarabia, D., Bacic, A. and Roessner, U. (2015) Mass Spectrometry Imaging for Plant Biology: A Review. Phytochemistry Reviews, 15, 1-44.

  413. 413. Tokareva, E.N., Pranovich, A.V. and Holmbom, B.R. (2011) Characteristic Fragment Ions from Lignin and Polysaccharides in ToF-SIMS. Wood Science and Technology, 45, 767-785.
    http://dx.doi.org/10.1007/s00226-010-0392-9

  414. 414. Saito, K., Mitsutani, T., Imai, T., Matsushita, Y., Yamamoto, A. and Fukushima, K. (2008) Chemical Differences between Sapwood and Heartwood of Chamaecyparis obtusa Detected by ToF-SIMS. Applied Surface Science, 255, 1088-1091.
    http://dx.doi.org/10.1016/j.apsusc.2008.05.145

  415. 415. Araújo, P., Ferreira, M.S., de Oliveira, D.N., Pereira, L., Sawaya, A.C.H.F., Catharino, R.R., et al. (2014) Mass Spectrometry Imaging: An Expeditious and Powerful Technique for Fast in Situ Lignin Assessment in Eucalyptus. Analytical Chemistry, 86, 3415-3419.
    http://dx.doi.org/10.1021/ac500220r

  416. 416. Imai, T., Tanabe, K., Kato, T. and Fukushima, K. (2005) Localization of Ferruginol, a Diterpene Phenol, in Cryptomeria japonica Heartwood by Time-of-Flight Secondary Ion Mass Spectrometry. Planta, 221, 549-556.
    http://dx.doi.org/10.1007/s00425-004-1476-2

  417. 417. Kuroda, K., Fujiwara, T., Hashida, K., Imai, T., Kushi, M., Saito, K., et al. (2014) The Accumulation Pattern of Ferruginol in the Heartwood-Forming Cryptomeria japonica Xylem as Determined by Time-of-Flight Secondary Ion Mass Spectrometry and Quantity Analysis. Annals of Botany, 113, 1029-1036.
    http://dx.doi.org/10.1093/aob/mcu028

  418. 418. Lunsford, K.A., Peter, G.F. and Yost, R.A. (2011) Direct Matrix-Assisted Laser Desorption/Ionization Mass Spectrometric Imaging of Cellulose and Hemicellulose in Populus Tissue. Analytical Chemistry, 83, 6722-6730.
    http://dx.doi.org/10.1021/ac2013527

  419. 419. Jarvis, M.C. and Apperley, D.C. (1990) Direct Observation of Cell Wall Structure in Living Plant Tissues by Solid-State 13C NMR Spectroscopy. Plant Physiology, 92, 61-65.
    http://dx.doi.org/10.1104/pp.92.1.61

  420. 420. Fenwick, K.M., Jarvis, M.C. and Apperley, D.C. (1997) Estimation of Polymer Rigidity in Cell Walls of Growing and Nongrowing Celery Collenchyma by Solid-State Nuclear Magnetic Resonance in Vivo. Plant Physiology, 115, 587- 592.

  421. 421. Bals, S., Van Aert, S. and Van Tendeloo, G. (2013) High Resolution Electron Tomography. Current Opinion in Solid State and Materials Science, 17, 107-114.
    http://dx.doi.org/10.1016/j.cossms.2013.03.001

  422. 422. Otegui, M.S., Mastronarde, D.N., Kang, B.-H., Bednarek, S.Y. and Staehelin, L.A. (2001) Three-Dimensional Analysis of Syncytial-Type Cell Plates during Endosperm Cellularization Visualized by High Resolution Electron Tomography. The Plant Cell, 13, 2033-2051.
    http://dx.doi.org/10.1105/tpc.13.9.2033

  423. 423. Doblin, M.S., Pettolino, F. and Bacic, A. (2010) Evans Review: Plant Cell Walls: The Skeleton of the Plant World. Functional Plant Biology, 37, 357-381.
    http://dx.doi.org/10.1071/FP09279

  424. 424. Pabst, M., Fischl, R.M., Brecker, L., Morelle, W., Fauland, A., Kofeler, H., et al. (2013) Rhamnogalacturonan II Structure Shows Variation in the Side Chains Monosaccharide Composition and Methylation Status within and across Different Plant Species. The Plant Journal, 76, 61-72.
    http://dx.doi.org/10.1111/tpj.12271

NOTES

*Corresponding author.