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Plant Xyloglucan:Xyloglucosyl Transferases: History
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Subjects: Biochemistry & Molecular Biology
Contributor: Maria Hrmova

Plant xyloglucan:xyloglucosyl transferases, known as xyloglucan endo-transglycosylases (XETs) are the key players that underlie plant cell wall dynamics and mechanics. These fundamental roles are central for the assembly and modifications of cell walls during embryogenesis, vegetative and reproductive growth, and adaptations to living environments under biotic and abiotic (environmental) stresses. Xyloglucan:xyloglucosyl transferases or xyloglucan endo-transglycosylases (XET), classified under EC 2.4.1.207, transfer the glycosyl groups from one glycoside to another. These enzymes were discovered in 1992 independently in bean epicotyls, nasturtium seeds, and pea, tomato and other plant extracts, and since their discovery, significant knowledge has been accumulated on their mode of action.

  • xyloglucosyl transferases
  • evolutionary history
  • glycoside hydrolase family 16
  • mechanism of catalysis
  • molecular modelling and dynamics
  • transglycosylation reactions
  • substrate binding

1. Nomenclatureand classification

The nomenclature of XET enzymes is defined by the International Union of Biochemistry and Molecular Biology (IUBMB) and International Union of Pure and Applied Chemistry Biochemical Nomenclature Committee [1], the Kyoto Encyclopaedia of Genes and Genomes Enzyme Database, and the BRENDA Comprehensive Enzyme System [2]. These nomenclatures either consider the transfer of the ‘glycosyl’ (xyloglucan endo-transglycosylase) [3][4][5] or ‘glucosyl’ (xyloglucan endo-transglucosylase) [6][7] groups.
The complex view on XET enzyme nomenclature and classification is provided by the Carbohydrate-Active enZYmes Database (CAZy) [8] and CAZypedia [9], which classify entries based on protein tertiary structures, substrate specificities and evolutionary history or phylogenomic relationships. XETs are listed in the glycoside hydrolase (GH) family 16, and not in a glycoside transferase (GT) Leloir group of enzymes. According to the Enzyme Commission and International Union of Biochemistry and Molecular Biology, the latter group involves enzymes that use activated sugars (for example UDP-glucose) as the glycosyl donors, and transfer the glycosyl groups to the nucleophilic glycoside (or other) acceptors. The GH16 family, according to tertiary structural features is sub-divided into 23 subfamilies [10], where the subfamily GH16_20 includes XETs but also xyloglucan endohydrolases (XEHs, EC 3.2.1.151) [11]; the latter group catalyses predominantly hydrolytic reactions of XGs. Thus, the GH16_20 group contains the products of XTH (xyloglucan transglycosylase/hydrolase) genes encoding both types of XG-modifying enzymes which notably display close similarities in tertiary structures [12].

2.  Catalytic mechanism

The catalytic function of XETs is defined by breaking a glycosidic bond between 1,4-βdlinked glucosyl residues of XGs (or other polysaccharides) and transferring the xyloglucanyl or another glycoside moiety onto the O-4 atom of a non-reducing end of a glycoside acceptor; this acceptor could be XG, XG-oligosaccharide or another glycoside. This constitutes a so-called ping-pong bi-bi reaction mechanism and not a sequential one [13][14]. Recently, the definition of XET substrate specificity has significantly expanded, and thus the previous information on XETs has become obsolete [15][16][17][18][19][20][21][22][23].
The first steps of the transglycosylation and hydrolytic reactions catalysed by XTHs (that is XETs and XEHs) are binding and cleavage of donor glycoside substrates. The difference between XETs and XEHs occurs in the second step, in which the fragment with the original non-reducing end of the donor substrate is transferred to an acceptor, which in the case of a typical transglycosylase is a saccharide, while in the case of a hydrolase, it is an activated water molecule [8][24]. This second step had the key importance for the nomenclature and classification of XETs as transferases.
The transglycosylation mechanism catalysed by XET enzymes proceeds in two stages with two transition states. The first step involves the deprotonation of a carboxylate that operates as a nucleophile and attacks the anomeric carbon of the donor substrate, forming the glycosyl–enzyme intermediate complex, with the participation of an acid/base carboxylate. In the hybrid aspen Populus tremulus x tremuloides PttXET16A, the nucleophile attacking the anomeric carbon is Glu85, while Glu89 acts as an acid/base (has a dual role), that is it protonates the aglycon saccharide and releases it, and subsequently, it de-protonates the glycoside acceptor. The third important residue in PttXET16A is Asp87 positioned between Glu85 and Glu89 (signature motif ExDxE), which forms a tight bond with the Glu85 nucleophile, and interacts with the substrates through hydrogen bonds during substrate binding. The nucleophile must be de-protonated for the donor substrate attack, while Asp87 and Glu89 remain protonated. The preference for the transfer reaction instead of hydrolytic one by XETs leads to the re-ligation of the nascent donor end of one saccharide to the non-reducing end of the acceptor substrate [12][25]. Curiously, in the crystal structure of PttXET16A (‘true’ XET without hydrolytic activity) [25], only a few water molecules were resolved near the donor and acceptor substrate binding sites, while significantly more water molecules were located around the corresponding sites in XEHs.
Typically, hydrolases also transglycosylate under high(er) substrate concentrations [24][26], when in the later stages of reactions, the glycosyl–enzyme intermediate complex interacts with another sugar molecule (instead of an activated water molecule), thus shifting the reaction equilibrium towards the transfer products. Contrary to these high-substrate-induced transglycosylation reactions, ‘true’ transglycosylases such as XETs encounter a second saccharide regardless of a substrate concentration. While transglycosylation reactions catalysed by hydrolases occur under high(er) concentrations of substrates and generated products, this is not the case of XETs.
In addition to XETs, other plant enzymes with transglycosylase activities were reported, often without providing protein sequence information, with trivial descriptions such as mannan endotransglycosylase/hydrolases recognising (1,4)-β-D-mannan-derived polysaccharides [27], xylan endo-transglycosylases functionalising heteroxylan polysaccharides [28][29][30], mixed-linkage glucan:xyloglucan endotransglucosylases acting on (1,3;1,4)-β-D-glucans [31][32], and hetero-trans-β-glucanases (HTGs) functionalising cellulose [16][33]. However, terming these enzymes as such disguises a fact that some of them could in fact be non-specific XETs.

3. Structural properties, evolutionary relationships, enzyme activity methods

The crystal structures of the XET enzyme PttXET16A (Protein Data Bank—PDB accessions and 1UMZ and 1UN1 at respective 1.80 Å and 2.10 Å resolution; 1UMZ in complex with the XLLG acceptor) [12][25][34], offers insights into its atomic architecture and catalytic mechanism. The crystal structure revealed that PttXET16A has the β-jelly-roll topology and folds into a β-sandwich with convex and concave regions of the two antiparallel β-sheets (Figure 1). The catalytic trio, formed by three Glu85, Glu89 and Asp87 residues, is located around the middle of the convex region of the structure (Figure 1A; magenta cpk sticks). The elongated C-terminus is folded in an α-helix and a β-strand on the concave side of the molecule and stabilised by a disulfide bridge. The PttXET16A structure is also glycosylated at Asn93 with two N-acetylglucosaminyl and one mannosyl residues that are further stabilised by the hydrogen bonds. No other atomic XET structures have been reported; however, structural features of the 3D models of nasturtium, barley (Figure 1B, Figure 2 and Figure 3), and other XETs were defined [18][20][22][23][33][35]. Although, caution should be exercised when extracting and interpreting structural information based on 3D models, where side-chain placements are known to be unreliable even at high similarities between templates and target sequences [36].
Additional information on the PttXET16A enzyme was obtained through molecular dynamics (MD) simulations [12], where the interactions between residues of the PttXET16A and two XLLG nonasaccharides revealed that one XLLG occupied the donor site and created a stable intermediate with the enzyme, while the second XLLG remained bound at the acceptor site. Notably, the reducing-end glucose moiety of XLLG at the donor -1 subsite, altered its low energy 4C1 conformation into 1S3 boat.
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Figure 1. β-Sandwich architectures and β-jelly-roll topologies of Populus XET16A (PDB accession 1UMZ) in complex with the XLLG acceptor substrate (A), and the Tropaeolum XET6.3 3D molecular model in complex with the XXXG donor substrate (B). (A) Left panel: The XLLG acceptor (cpk sticks) bound in the Populus XET16A structure is indicated in dashed lines at 2.6 Å to 3.5 Å separations. The catalytic trio (Glu85, Glu89 and Asp87) is shown in cpk magenta sticks. Right panel: Details of the XLLG binding in Populus XET16A; interacting residues are marked in green cpk sticks and dots. Bottom panel: Some of the residues of Populues XET16A binding XLLG (highlighted in green) are shown in the alignment by PROMALS3D [37] of the Populus and Tropaeolum XET sequences (numbering includes signal peptides). Conservation of residues on the scale 9–6 is shown on the top of the alignment in brown. (B) Left panel: The XXXG donor (cpk sticks) bound in Tropaeolum XET6.3 (cyan; coordinates from [20]) is indicated in dashed lines at separations between 2.3 Å and 3.0 Å. Right panel: Details of XXXG binding in Tropaeolum XET6.3; interacting residues are marked in cyan cpk sticks in dots. Bottom panel: Some of the residues of Tropaeolum XET6.3 binding XLLG (highlighted in cyan) are shown in the alignment by PROMALS3D [37] of Populus XET16A and Tropaeolum XET6.3 sequences (numbering includes signal peptides). Conservation of residues on the scale 9–6 is shown on the top of the alignment in brown. Images were generated in the PyMOL Molecular Graphics System v2.5.2 (Schrődinger LLC, Portland, OR, USA).
Structural differences, which determine whether the XET/XEH enzymes transglycosylate (PDB accessions 1UN1, 1UMZ) or act as the hydrolases (TmNXG1; PDB accession 2UWA) [25][34][38], were reported including the TmNXG1-DELTAYNIIG mutant (PDB accession 2VH9). Slight structural differences, which led to the stronger binding of a donor substrate combined with larger loop flexibility at the acceptor site and the flexibility of residues, underlined the hydrolytic preference [34][38].
As detailed above, structural differences modulate the binding of donor substrates combined with the loop and residues flexibilities and underline the transfer or hydrolytic preferences of XTH enzymes. This is supported by phylogenomic analyses, where XTHs segregate in the three groups, with XEHs belonging to XTH III clade, while XETs clustered within XTH I and XTH II clades [18][20][34][39]. Based on these analyses, it was concluded that GH16 hydrolases including XEHs have evolved from XETs [18][38][40].
As previously summarised [23], XET activity assays use unlabelled donor substrates and radio-chemically or fluorescently labelled acceptors, or radio-labelled donor and unlabelled acceptors. The reaction products in both cases encompass labelled saccharides, while unincorporated donors or acceptors are removed via chromatography [3][15][20], or through a gel [17][41], washed off from a filter paper [42] or ethanol precipitated [43]. When selecting an appropriate approach, it is critical to select the right type of fluorescent tag [44][45] and how to remove efficiently the surplus of substrates [16][23]. Labelled oligosaccharide probes could be also utilised in high-throughput polysaccharide microarrays [46] and through in vivo visualisation [47][48][49][50]. Supplementary XET activity assays include colorimetry [51] that relies on the formation of the blue-green-colored iodine–XG complex, and viscometry that records viscosity changes in substrates.
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Figure 2. Molecular models of the β-sandwich architecture with the β-jelly-roll topology of Hordeum XET3 (top left; yellow), Hordeum XET4 (top right; orange), Hordeum XET5 (bottom left; pink), and Hordeum XET6 (bottom right; lemon) in complex with the XLLG acceptors (cpk sticks). Coordinates of XET3, XET4 and XET6 with XLLG were taken from [35] and those of XET5 from [18]. XLLG was docked in XET5 by HDOCK that performs docking based on a hybrid algorithm of template-based modelling and ab initio free docking [52]; the top docking pose is shown for XET5 from 100 poses ranked by energy docking scores. Details of binding of XLLG by Hordeum XETs are shown in dashed lines. Separations are for XET3: 2.7 Å to 3.6 Å; XET4: 2.6 Å to 3.5 Å; XET5: 2.5 Å to 3.6 Å; XET6: 2.5 Å to 3.2 Å. Images were generated in PyMOL as referenced in Figure 1.

4.  Substrate specificity of transfer reactions with xyloglucan and other than xyloglucan-derived donors and acceptors

Substrate specificity of XETs has recently been evaluated in detail ([23]; cf. Table 1, vide infra); thus, here the researchers only succinctly recapitulate key aspects of this topic.
The systematic description of xyloglucan:xyloglucosyl transferases (EC 2.4.1.207) has from the onset of their discoveries stated that these enzymes recognise XG donor (xyloglucan) and acceptor (xyloglucosyl) substrates—this catalytic activity represents (i) homo-transglycosylation reactions. Yet, since 2007 [15], the novel types of transfer reactions were described in barley XETs and later in other plant sources, termed (ii) hetero-transglycosylation reactions [17][20]. These reactions utilise the neutral donor and neutral or ionic acceptor substrates other than XG-derived [23].
(i) Homo-transglycosylation reactions with XG-derived donors and acceptors
The poplar PttXET16A [25][34] and Pinus radiata PrXTH1 [53] XETs, belonging to the I/II cluster of XTH gene products, were characterised as the homo-transglycosylation-type enzymes with a narrow substrate specificity that utilised XG-derived substrates. Hetero-transglycosylation activities in PttXET16A were undetected with cello-oligosaccharides [12], although the data with PrXTH1 suggested the weak binding of cellooctaose [53]. Phylogenomic analyses of the GH16 family, spanning monocots, eudicots and a basal Angiosperm [22] revealed that PttXET16A clustered with barley (Hordeum vulgare L.) XET5 that catalysed the reactions with XG or hydroxyethyl cellulose and XG- or cellulose-derived oligosaccharides [15].
(ii) Hetero-transglycosylation reactions with neutral donor and neutral or ionic acceptors,other than XG-derived
The key question to answer regarding hetero-transglycosylation reactions is, what is the limit for the activity ratio of the XG/XG-oligosaccharide substrate pair and chemically different substrates? Another key point is that the substrate specificity of only a few XETs in near-homogenous forms has been defined, and thus this property cannot be unequivocally assigned to many XETs.
Although the broad XET substrate specificity in Poaceae was predicted based on molecular modelling [54], the first experimental indication that XETs could catalyse hetero-transglycosylation reactions, i.e., to mediate transfers between saccharides other than XG-derived, was presented by Ait Mohand and Farkaš [17]. This study described glycosyl transfers from XG to cello- and laminari-oligosaccharides, and from carboxymethyl and hydroxyethyl cellulose derivatives to XG oligosaccharides, using the crude protein extracts prepared from germinating nasturtium seeds [17]. This study was followed with the barley XET5 isoform [15], the first XET in a near-homogenous form, with a defined primary structure, that catalysed hetero-transglycosylation reactions in vitro. Except for XGs, this enzyme linked covalently carboxymethyl and hydroxyethyl celluloses, and (1,3;1,4)-β-D-glucans as donors and XG- and cellulose-derived oligosaccharide acceptors. The 44% efficiency of barley XET5 with the hydroxyethyl cellulose donor and the XG-oligosaccharides was comparable to that of XG, but it was significantly lower with (1,3;1,4)-β-D-glucan. Subsequently, the hetero-transglycosylation reactions were defined for a near-homogenous barley XET6 isoform [18].
Other XET enzymes recognising cellulose as the donor substrate, were partially purified XET from parsley roots (Petroselinum crispum Mill. Fus) [43] and near-homogenous XTH3 from Arabidopsis thaliana L. Heynh, which catalysed the hetero-transglycosylation reactions between cellulose and cello-oligosaccharides or cellulose and XG-oligosaccharides [19]. Despite differences in the specificity of hetero-transglycosylation reactions, barley XET5 [15] and Arabidopsis XTH3 [19] clustered in the same phylogenetic group of XTH I, as presumably XG-specific PttXET16A and PrXTH1 [34][53], although they segregated to different sub-clades [23]. The next XETs with known primary structures and utilising besides XG, cellulose or (1,3;1,4)-β-D-glucans as donors, were three acidic EfXTH-A, EfXTH-H and EfXTH-I isoforms from Equisetum fluviatile L. [55], although the homogeneity of these enzymes was not shown. While EfXTH-A displayed the comparable transfer of (1,3;1,4)-β-D-glucan fragments to XG-oligosaccharides, as was the case of barley XET5 (0.2–0.3%) [15], the activity with cellulose was higher with barley XET5. EfXTH-H and EfXTH-I showed the equivalent hetero-transglycosylation activities with (1,3;1,4)-β-D-glucans and cellulose donors [55]. Conversely, HTG and MLG (1,3;1,4-β-D-glucan): xyloglucan endotransglycosylase enzymes from Equisetum preferred cellulose and (1,3;1,4)-β-D-glucans with XG-oligosaccharides as respective donors and acceptors [55]. Such enzymes were so far found only in Equisetum and the charophytic algae [28][56], where they are predicted to re-model hemicelluloses in horsetails shoots [32]. Molecular modelling of HTG suggested the amino acid residues responsible for the evolution of their substrate specificity [33].
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Figure 3. Details of the XXXG donor (cpk sticks) and the [α(1-4)GalAp]5 acceptor (magenta cpk sticks) binding substrates in Hordeum XET3 (A) and Hordeum XET4 (B). In both panels, blue and black dashed lines indicate residue separations between donors or acceptors that are 2.6 Å–3.6 Å (XET3) and 2.7 Å–3.4 Å (XET4). Interacting residues are shown in yellow (XET3) and orange (XET4) cpk sticks and emphasised in dots. Subsites at -4 to -1 for XXXG and +1 to +5 for [α(1-4)GalAp]5 are indicated. Images were generated in PyMOL as referenced in Figure 1. Bottom panel: Some of the residues of Hordeum XETs that bind [α(1-4)GalAp]5 (highlighted in yellow) are shown in the alignment by PROMALS3D [37] of the Hordeum XET sequences (numbering includes signal peptides). Conservation of residues on the scale 9–6 is shown on the top of the alignment in brown.
Recently, Tropaeolum majus L. XET6.3 classified in the XTH II clade [23] in its near-homogenous form displayed hetero-transglycosylation activity with the donor XG or hydroxyethyl cellulose and neutral oligosaccharide acceptors, such as those derived from cellulose, (1,3;1,4)-β-D-glucan, laminarin, pustulan, arabinoxylan, xylan, arabinan, arabinogalactan, mannan, glucomannan and galactomannan [20].
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Figure 4. Docking of the XXXG donor (cpk cyan sticks) and the xylotetraose (Xyl-OS4) acceptor (cpk magenta sticks) substrates in the active sites of Populus XET16A (PDB accession 1UN1) (A) and Tropaeolum XET6.3 [20] (B), and MD simulations of enzyme/substrate complexes after 0, 50, and 1000 ns (simulation times indicated in bottom-left corners). MD simulations were carried out similarly as described in [57]. Separations between the donors and acceptor substrates are between 2.6 Å and 3.6 Å for Populus XET16A and 2.7 Å–3.4 Å for Tropaeolum XET6.3. Residues (numbering includes signal peptides) mediating contacts with substrates are shown in cpk sticks. In Tropaeolum XET6.3, E136 and Q138 stabilise the binding of the Xyl-OS4 acceptor. For clarity, the residues and the subsite binding sites (−4 to −1 for XXXG and +1 to +4 for Xyl-OS) are shown in left panels only. Images were generated in PyMOL as referenced in Figure 1.
Finally, it is important to describe hetero-transglycosylation reactions that accept charged acceptors that were shown in several near-homogenous barley XETs [22] (Figure 3), where all tested neutral oligosaccharides served as the acceptors with barley XET3-XET6 isoforms, although with different efficiency. Barley XET3 and XET4 also transferred efficiently the fragments of XG or hydroxyethyl cellulose to the penta-galacturonic acid ([α(1-4)GalAp]5), a fragment of the linear part of pectin, [22]. These hetero-transglycosylation activities by barley XETs were demonstrated in vitro in CWs of barley roots using fluorescently labelled [α(1-4)GalAp]5) [22]. These findings were supported by the structural modelling of barley XET3 and XET4 with the docked XG heptasaccharide (XXXG) donor and the [α(1-4)GalAp]5 acceptor substrates in the −4 to +5 active site subsites (Figure 3), and suggested that the XET substrate poly-specificity resulted from protein sequences alterations that evoked structural re-arrangements [20][21][22][23][58].
In summary, only specific combinations of residues could underlie certain substrate specificity in XETs, which underpins the functionalisation of substrates other than XG-derived. Despite categorising these residues [20][22], the future understanding of the mechanisms of these reactions depends on the resolution of atomic structures of XETs. Future research will also be focused on in vitro and in silico studies to dissect the molecular mechanisms that drive the transglycosylation reactions catalysed by XETs (Figure 4).

This entry is adapted from the peer-reviewed paper 10.3390/ijms23031656

References

  1. Webb, E.C. Enzyme Nomenclature 1992: Recommendations of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology on the Nomenclature and Classification of Enzymes; Academic Press Inc.: Cambridge, MA, USA; Harcourt Brace Jovanovich Publishers: San Diego, CA, USA, 1992; ISBN 0-12-227164-5.
  2. Placzek, S.; Schomburg, I.; Chang, A.; Jeske, L.; Ulbrich, M.; Tillack, J.; Schomburg, D. BRENDA in 2017: New perspectives and new tools in BRENDA. Nucleic Acids Res. 2017, 45, D380–D388.
  3. Nishitani, K.; Tominaga, R. Endo-xyloglucan transferase, a novel class of glycosyltransferase that catalyzes transfer of a segment of xyloglucan molecule to another xyloglucan molecule. J. Biol. Chem. 1992, 267, 21058–21064.
  4. Farkaš, V.; Sulová, Z.; Stratilová, E.; Hanna, R.; Maclachlan, G. Cleavage of xyloglucan by nasturtium seed xyloglucanase and transglycosylation to xyloglucan subunit oligosaccharides. Arch. Biochem. Biophys. 1992, 298, 365–370.
  5. Fry, S.; Smith, R.; Renwick, K.; Martin, D.; Hodge, S.; Matthews, K. Xyloglucan endotransglycosylase, a new wall-loosening enzyme activity from plants. Biochem. J. 1992, 282, 821–828.
  6. Rose, J.K.C.; Braam, J.; Fry, S.C.; Nishitani, K. The XTH family of enzymes involved in xyloglucan endotransglucosylation and endohydrolysis: Current perspectives and a new unifying nomenclature. Plant Cell Physiol. 2002, 43, 1421–1435.
  7. Atkinson, R.G.; Johnston, S.L.; Yauk, Y.K.; Sharma, N.N.; Schröder, R. Analysis of xyloglucan endotransglucosylase/hydrolase (XTH) gene families in kiwifruit and apple. Postharvest Biol. Technol. 2009, 51, 149–157.
  8. Lombard, V.; Golaconda Ramulu, H.; Drula, E.; Coutinho, P.M.; Henrissat, B. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res. 2014, 42, D490–D495.
  9. CAZypedia Consortium. Ten years of CAZypedia: A living encyclopedia of carbohydrate-active enzymes. Glycobiology 2018, 28, 3–8.
  10. Viborg, A.H.; Terrapon, N.; Lombard, V.; Michel, G.; Gurvan, M.; Czjzek, M.; Henrissat, B.; Brumer, H. A subfamily roadmap for functional glycogenomics of the evolutionarily diverse Glycoside Hydrolase Family 16 (GH16). J. Biol. Chem. 2019, 294, 15973–15986.
  11. Eklöf, J.M.; Brumer, H. The XTH gene family: An update on enzyme structure, function, and phylogeny in xyloglucan remodeling. Plant Physiol. 2010, 153, 456–466.
  12. Mark, P.; Baumann, M.J.; Eklöf, J.M.; Gullfot, F.; Michel, G.; Kallas, A.M.; Teeri, T.T.; Brumer, H.; Czjek, M. Analysis of nasturtium TmNXG1 complexes by crystallography and molecular dynamics provides detailed insight into substrate recognition by family GH16 xyloglucan endo-transglycosylases and endo-hydrolases. Proteins 2009, 75, 820–836.
  13. Baran, R.; Sulová, Z.; Stratilová, E.; Farkaš, V. Ping-Pong character of nasturtium-seed xyloglucan endotransglycosylase (XET) Reaction. Gen. Physiol. Biophys. 2000, 19, 427–440.
  14. Saura-Valls, M.; Faure, R.; Ragas, S.; Piens, K.; Brumer, H.; Teeri, T.T.; Cottaz, S.; Driguez, H.; Planas, A. Kinetic analysis using low-molecular mass xyloglucan oligosaccharides defines the catalytic mechanism of a Populus xyloglucan endotransglycosylase. Biochem. J. 2006, 395, 99–106.
  15. Hrmova, M.; Farkaš, V.; Lahnstein, J.; Fincher, G.B. A barley xyloglucan xyloglucosyl transferase covalently links xyloglucan, cellulosic substrates, and (1,3;1,4)-β-d-glucans. J. Biol. Chem. 2007, 283, 27344.
  16. Franková, L.; Fry, S.C. Biochemistry and physiological roles of enzymes that ‘cut and paste’ plant cell-wall polysaccharides. J. Exp. Bot. 2013, 64, 3519–3550.
  17. Ait Mohand, F.; Farkaš, V. Screening for hetero-transglycosylating activities in extracts from nasturtium (Tropaeolum majus). Carbohydr. Res. 2006, 34, 577–581.
  18. Hrmova, M.; Farkaš, V.; Harvey, A.J.; Lahnstein, J.; Wischmann, B.; Kaewthai, N.; Ezcurra, I.; Teeri, T.T.; Fincher, G.B. Substrate specificity and catalytic mechanism of a xyloglucan xyloglucosyl transferase HvXET6 from barley (Hordeum vulgare L.). FEBS J. 2009, 276, 437–456.
  19. Shinohara, N.; Sunagawa, N.; Tamura, S.; Yokoyama, R.; Ueda, M.; Igarashi, K.; Nishitani, K. The plant cell-wall enzyme AtXTH3 catalyses covalent cross-linking between cellulose and cello-oligosaccharide. Sci. Rep. 2017, 7, 46099.
  20. Stratilová, B.; Firáková, Z.; Klaudiny, J.; Šesták, S.; Kozmon, S.; Strouhalová, D.; Garajová, S.; Ait-Mohand, F.; Horváthová, Á.; Farkaš, V.; et al. Engineering the acceptor substrate specificity in the xyloglucan endotransglycosylase TmXET6.3 from nasturtium seeds (Tropaeolum majus L.). Plant Mol. Biol. 2019, 100, 181–197.
  21. Herburger, K.; Franková, L.; Sanhueza, D.; Roig-Sanchez, S.; Meulewaeter, F.; Hudson, A.; Thomson, A.; Laromaine, A.; Budtova, T.; Fry, S.C. Enzymically attaching oligosaccharide-linked ‘cargoes’ to cellulose and other commercial polysaccharides via stable covalent bonds. Int. J. Biol. Macromol. 2020, 164, 4359–4369.
  22. Stratilová, B.; Šesták, S.; Mravec, J.; Garajová, S.; Pakanová, Z.; Vadinová, K.; Kučerová, D.; Kozmon, S.; Schwerdt, J.G.; Shirley, N.; et al. Another building block in the plant cell wall: Barley xyloglucan xyloglucosyl transferases link covalently xyloglucan and anionic oligosaccharides derived from pectin. Plant J. 2020, 104, 752–767.
  23. Stratilová, B.; Kozmon, S.; Stratilová, E.; Hrmova, M. Xyloglucan xyloglucosyl transferases and the cell wall structure: Subtle but significant. Molecules 2020, 25, 5619.
  24. Hrmova, M.; MacGregor, E.A.; Biely, P.; Stewart, R.J.; Fincher, G.B. Substrate binding and catalytic mechanism of a barley β-d-glucosidase/(1,4)-β-d-glucan exohydrolase. J. Biol. Chem. 1998, 273, 11134–11143.
  25. Johansson, P.; Brumer, H.; Baumann, M.J.; Kallas, A.M.; Henriksson, H.; Denman, S.E.; Teeri, T.T.T.; Jones, T.A. Crystal structures of a poplar xyloglucan endotransglycosylase reveal details of transglycosylation acceptor binding. Plant Cell 2004, 16, 874–886.
  26. Hrmova, M.; Fincher, G.B. Barley β-d-glucan exohydrolases. Substrate specificity and kinetic properties. Carbohydr. Res. 1998, 305, 209–221.
  27. Schröder, R.; Atkinson, R.G.; Redgwell, R.J. Re-interpreting the role of endo-β-mannanases as mannan endotransglycosylase/hydrolases in the plant cell wall. Ann. Bot. 2009, 104, 197–204.
  28. Franková, L.; Fry, S.C. Phylogenetic variation in glycosidases and glycanases acting on plant cell wall polysaccharides, and the detection of transglycosidase and trans-β-xylanase activities. Plant J. 2011, 67, 662–681.
  29. Johnston, S.; Prakash, R.; Chen, N.J.; Kumagai, M.H.; Turano, H.M.; Cooney, J.M.; Atkinson, R.G.; Paull, R.E.; Cheetamun, R.; Bacic, A.; et al. An enzyme activity capable of endotransglycosylation of heteroxylan polysaccharides is present in plant primary cell walls. Planta 2013, 237, 173–187.
  30. Derba-Maceluch, M.; Awano, T.; Takahashi, J.; Lucenius, J.; Ratke, C.; Kontro, I.; Busse-Wicher, M.; Kosik, O.; Tanaka, R.; Winzéll, A.; et al. Suppression of xylan endotransglycosylase PtxtXyn10A affects cellulose microfibril angle in secondary wall in aspen wood. New Phytol. 2015, 205, 666–681.
  31. Fry, S.C.; Mohler, K.E.; Nesselrode, B.H.W.A.; Franková, L. Mixed-linkage β-glucan:xyloglucan endotransglucosylase, a novel wall-remodelling enzyme from Equisetum (horsetails) and charophytic algae. Plant J. 2008, 55, 240–252.
  32. Mohler, K.E.; Simmons, T.J.; Fry, S.C. Mixed-linkage glucan:xyloglucan endotransglucosylase (MXE) re-models hemicelluloses in Equisetum shoots but not in barley shoots or Equisetum callus. New Phytol. 2013, 197, 111–122.
  33. Simmons, T.J.; Mohler, K.E.; Holland, C.; Goubet, F.; Franková, L.; Houston, D.R.; Hudson, A.D.; Meulewaeter, F.; Fry, S.C. Hetero-trans-β-glucanase, an enzyme unique to Equisetum plants, functionalizes cellulose. Plant J. 2015, 83, 753–769.
  34. Baumann, M.J.; Eklöf, J.M.; Michel, G.; Kallas, A.M.; Teeri, T.T.; Czjzek, M.; Brumer, H. Structural evidence for the evolution of xyloglucanase activity from xyloglucan endo-transglycosylases: Biological implications for cell wall metabolism. Plant Cell 2007, 19, 1947–1963.
  35. Vaaje-Kolstad, G.; Farkaš, V.; Fincher, G.B.; Hrmova, M. Barley xyloglucan xyloglucosyl transferases bind xyloglucan-derived oligosaccharides in their acceptor-binding regions in multiple conformational states. Arch. Biochem. Biophys. 2010, 496, 61–68.
  36. Eramian, D.; Eswar, N.; Shen, M.; Sali, A. How well can the accuracy of comparative protein structure models be predicted? Prot. Sci. 2008, 17, 1881–1893.
  37. Pei, J.; Kim, B.-H.; Grishin, N.V. PROMALS3D: A tool for multiple protein sequence and structure alignments. Nucleic Acids Res. 2008, 36, 2295–2300.
  38. McGregor, N.; Yin, V.; Tung, C.C.; Van Petegem, F.; Brumer, H. Crystallographic insight into the evolutionary origins of xyloglucan endotransglycosylases and endohydrolases. Plant J. 2017, 89, 651–670.
  39. Kaewthai, N.; Harvey, A.J.; Hrmova, M.; Brumer, H.; Ezcurra, I.; Teeri, T.T.; Fincher, G.B. Heterologous expression of diverse barley XTH genes in the yeast Pichia pastoris. Plant Biotechnol. 2010, 27, 251–258.
  40. Behar, H.; Graham, S.W.; Brumer, H. Comprehensive cross-genome survey and phylogeny of glycoside hydrolase family 16 members reveals the evolutionary origin of EG 16 and XTH proteins in plant lineages. Plant J. 2018, 95, 1114–1128.
  41. Farkaš, V.; Ait-Mohand, F.; Stratilová, E. Sensitive detection of transglycosylating activity of xyloglucan endotransglycosylase/hydrolase (XTH) after isoelectric focusing in polyacrylamide gels. Plant Physiol. Biochem. 2005, 43, 431–435.
  42. Fry, S.C. Novel ‘dot-blot’ assays for glycosyltransferases and glycosylhydrolases: Optimization for xyloglucan endotransglycosylase (XET) activity. Plant J. 1997, 11, 1141–1150.
  43. Garajová, S.; Flodrová, D.; Ait-Mohand, F.; Farkaš, V.; Stratilová, E. Characterization of two partially purified xyloglucan endotransglycosylases from parsley (Petroselinum crispum) roots. Biologia 2008, 63, 313–319.
  44. Vaaje-Kolstad, G.; Farkaš, V.; Hrmova, M.; Fincher, G.B. Xyloglucan xyloglucosyl transferases from barley (Hordeum vulgare L.) bind oligomeric and polymeric xyloglucan molecules in their acceptor binding sites. Biochim. Biophys. Acta 2010, 1800, 674–684.
  45. Kosík, O.; Garajová, S.; Matulová, M.; Řehulka, P.; Stratilová, E.; Farkaš, V. Effect of the label of oligosaccharide acceptors on the kinetic parameters of nasturtium seed xyloglucan endotransglycosylase (XET). Carbohydr. Res. 2011, 346, 357–361.
  46. Kosík, O.; Auburn, R.P.; Russell, S.; Stratilová, E.; Garajová, S.; Hrmova, M.; Farkaš, V. Polysaccharide microarrays for high-throughput screening of transglycosylase activities in plant extracts. Glycoconj. J. 2010, 27, 79–87.
  47. Vissenberg, K.; Martinez-Vilchez, I.M.; Verbelen, J.P.; Miller, J.G.; Fry, S.C. In vivo colocalization of xyloglucan endotransglycosylase activity and its donor substrate in the elongation zone of Arabidopsis roots. Plant Cell 2000, 12, 1229–1237.
  48. Nishikubo, N.; Awano, T.; Banasiak, A.; Bourquin, V.; Ibatullin, F.; Funada, R.; Brumer, H.; Teeri, T.T.; Hayashi, T.; Sundberg, B.; et al. Xyloglucan endo-transglycosylase (XET) functions in gelatinous layers of tension wood fibers in poplar—a glimpse into the mechanism of the balancing act of trees. Plant Cell Phys. 2007, 48, 843–855.
  49. Ibatullin, F.M.; Banasiak, A.; Baumann, M.J.; Greffe, L.; Takahashi, J.; Mellerowicz, E.J.; Brumer, H.A. Real-time fluorogenic assay for the visualization of glycoside hydrolase activity in planta. Plant Phys. 2009, 151, 1741–1750.
  50. Mravec, J.; Kračun, S.K.; Rydahl, M.G.; Westereng, B.; Miart, F.; Clausen, M.H.; Fangel, J.U.; Daugaard, M.; Van Cutsem, P.; De Fine Licht, H.H.; et al. Tracking developmentally regulated post-synthetic processing of homogalacturonan and chitin using reciprocal oligosaccharide probes. Development 2014, 141, 4841–4850.
  51. Sulová, Z.; Lednická, M.; Farkaš, V. A colorimetric assay for xyloglucan endotransglycosylase from germinating seeds. Anal. Biochem. 1995, 229, 80–85.
  52. Yan, Y.; Zhang, D.; Zhou, P.; Li, B.; Huang, S.-Y. HDOCK: A web server for protein-protein and protein-DNA/RNA docking based on a hybrid strategy. Nucleic Acids Res. 2017, 45, W365–W373.
  53. Morales-Quintana, L.; Carrasco-Orellana, C.; Beltrán, D.; Moya-León, M.A.; Herrera, R. Molecular insights of a xyloglucan endo-transglycosylase/hydrolase of radiata pine (PrXTH1) expressed in response to inclination: Kinetics and computational study. Plant Physiol. Biochem. 2019, 136, 155–161.
  54. Strohmeier, M.; Hrmova, M.; Fischer, M.; Harvey, A.J.; Fincher, J.B.; Pleiss, J. Molecular modeling of family GH16 glycoside hydrolases: Potential roles for xyloglucan transglucosylases/hydrolases in cell wall modification in the Poaceae. Prot. Sci. 2004, 13, 3200–3213.
  55. Holland, C.; Simmons, T.J.; Meulewaeter, F.; Hudson, A.; Fry, S.C. Three highly acidic Equisetum XTHs differ from hetero-trans-β-glucanase in donor substrate specificity and are predominantly xyloglucan homo-transglucosylases. J. Plant Physiol. 2020, 251, 153210.
  56. Herburger, K.; Ryan, L.M.; Popper, Z.A.; Holzinger, A. Localisation and substrate specificities of transglycanases in charophyte algae relate to development and morphology. J. Cell Sci. 2018, 131, jcs203208.
  57. Stratilová, B.; Řehulka, P.; Garajová, S.; Řehulková, H.; Stratilová, E.; Hrmova, M.; Kozmon, S. Structural characterization of the Pet c 1.0201 PR-10 protein isolated from roots of Petroselinum crispum (Mill.) Fuss. Phytochemistry 2020, 175, 112368.
  58. Seven, M.; Derman, Ü.C.; Harvey, A.J. Enzymatic characterization of ancestral/group-IV clade xyloglucan endotransglycosylase/hydrolase enzymes reveals broad substrate specificities. Plant J. 2021, 106, 1060–1073.
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