Chemical Papers 66 (9) 814–820 (2012)DOI: 10.2478/s11696-012-0167-x

ORIGINAL PAPER

Incorporation of β-(1,6)-linked glucooligosaccharides(pustulooligosaccharides) into plant cell wall structures‡

aZuzana Zemková, a,bSoňa Garajová, cDana Flodrová, dPavel Řehulka,aIvan Zelko, aRenáta Vadkertiová, aVladimír Farkaš, a,bEva Stratilová*

aInstitute of Chemistry, Centre for Glycomics, Slovak Academy of Sciences, Dúbravská cesta 9, SK-845 38 Bratislava, Slovakia

bInstitute of Chemistry, Centre of Excellence for White-Green Biotechnology, Slovak Academy of Sciences,

Trieda Andreja Hlinku 2, SK-949 76 Nitra, Slovakia

cInstitute of Analytical Chemistry, Academy of Sciences of the Czech Republic, Veveří 97, CZ-60200 Brno, Czech Republic

dInstitute of Molecular Pathology, Faculty of Military Health Sciences, University of Defence,

Třebešská 1575, CZ-50001 Hradec Králové, Czech Republic

Received 30 October 2011; Revised 16 February 2012; Accepted 26 February 2012

Protein extract of germinating nasturtium (Tropaeolum majus) seeds containing xyloglucan endo-transglycosylase (xyloglucan xyloglucosyl transferase, EC 2.4.1.207, abbreviated XET) exhibited theheterotransglycosylating activity with donor/acceptor substrate pair xyloglucan/sulphorhodaminelabelled pustulooligosaccharides (XG/PUOS–SR) in a dot blot assay. The heterotransglycosylatingactivity was confirmed by the substrate–product changes during transglycosylation by HPLC size-exclusion chromatography. Another donor substrate capable of being coupled with PUOS–SR wascellulose, probably owing to its structural similarity to xyloglucan. Surprisingly, microscopic com-parison of the incorporation of the labelled xyloglucan nonasaccharide XGO9–SR (specific substratefor XET) and PUOS–SR into the cell wall structures clearly showed differences in their binding tospecific cell structures: the primary cell wall and the plasma membrane. These findings indicatethe existence in nasturtium of XETs with different localisation, substrate specificity and, probably,function.c© 2012 Institute of Chemistry, Slovak Academy of Sciences

Keywords: xyloglucan endotransglycosylase, XET, heterotransglycosylation, cell wall, plasmamembrane

Introduction

Plant and fungal cells share many similarities in-cluding a sturdy cell wall with a complex structurelargely composed of polysaccharides and proteins. Theproperties of the cell walls are determined by the com-position and structure of their components. The com-plexity of plant and fungal cell walls is enhanced by thefact that the individual polysaccharides are mutuallyinterlinked by covalent bonds (Popper & Fry, 2008;

Cabib et al., 2008). The synthesis and the modifica-tions of cell wall polysaccharides needed for the req-uisite properties of the cell walls in each developmen-tal stage are accomplished by the enzymes from thegroup of glycosyltransferases (enzymes transferring aglycosyl group from one compound, the donor, to an-other compound, the acceptor) and glycoside hydro-lases (enzymes hydrolysing the glycosidic bond be-tween two or more carbohydrates or between a car-bohydrate and a non-carbohydrate moiety).

*Corresponding author, e-mail: chemevi@savba.sk‡Presented at the 5th Meeting on Chemistry & Life 2011, Brno, 14–16 September 2011.

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According to CAZy (Carbohydrate Active En-zymes database; Cantarel et al., 2009) based on thestructurally related catalytic and carbohydrate bind-ing domains, the transglycosylases belong to the groupof glycoside hydrolases whereas, according to Enzymenomenclature based on the chemical reactions theycatalyse, they belong to glycosyltransferases. This isbecause transglycosylases are special enzymes trans-ferring the fragment of the donor molecule with anewly created reducing end to another saccharide(transglycosylation) instead of to water (hydrolysis)(Sinnott, 1990). From the perspective of CAZy, glyco-syltransferases catalyse only the transfer of sugar moi-eties from activated donor molecules to specific accep-tor molecules, forming glycosidic bonds, but not trans-glycosylation. Transglycosylation can be observed inalmost all the retaining glycoside hydrolases, althoughonly at high concentrations of the final products ofhydrolysis. The “true” transglycosylases catalyse thetransfer of a donor fragment to an acceptor saccha-ride throughout the reaction. These enzymes enablethe cell walls to behave like dynamic structures, i.e.they enable the expansion and remodelling of the cellwall during cell growth without its weakening (Nishi-tani, 1997).To date, the best-known enzyme from this group

is plant xyloglucan endotransglycosylase, XET (Fryet al., 1992; Nishitani & Tominaga 1992; Farkas etal., 1992). It was presumed that some XETs fromseeds and fruits were able to catalyse both transglyco-sylation and hydrolysis, i.e. act as transglycosylases/hydrolases, XTHs (Rose et al., 2002; Eklof & Brumer,2010). One of these enzymes was XTH from nastur-tium seeds. Later, both typical xyloglucan endohydro-lase (Bauman et al., 2007; Mark et al., 2009) and mul-tiple forms of typical transglycosylases were identifiedin this source (Stratilová et al., 2010). At the sametime, a new and unexpected feature of some XETswas discovered – the ability to catalyse heterotrans-glycosylation, i.e. transfer and ligation of fragmentsbetween structurally different covalent bond-formingsaccharides (Ait Mohand & Farkaš, 2006; Hrmova etal., 2007, 2009; Stratilová et al., 2010). This findingled to a new concept in understanding the formationof covalent cross-linking of polysaccharide moleculesin the cell wall.The discovery of heterotransglycosylation stimu-

lated other investigators into an effort to identify newtypes of plant transglycosylases. Fry et al. (2008)demonstrated the presence of an enzyme catalysingthe formation of covalent bonds between mixed-linkage β-(1,3)-/β-(1,6)-glucan and labelled xyloglu-cooligosaccharides. Schroder et al. (2009) found thatβ-mannanase from tomato is a transglycosylase ableto transfer portions of galactomannan molecules tooligosaccharides derived from galactomannan.As there are many similarities between plant and

fungal cell walls (Klis et al., 2006; Lesage & Bussey,

2006), the question of the presence of transglycosy-lases catalysing homo- and/or heterotransglycosyla-tion in the cell walls of fungi has gained interest.Amongst others, the proteins Crh1 and Crh2 (Cabibet al., 2007) were found to be serious candidatesfor catalysing transglycosylation by the formation ofcovalent linkages between chitin and pustulan (Cabibet al., 2008).This work is focused on the ability of crude ex-

tracts from germinating nasturtium seeds to catalysethe incorporation of pustulooligosaccharides, β-(1,6)-linked glucooligosaccharides, typical of the yeast, butnot plant cell wall.

Experimental

Polysaccharides used in this study were of the fol-lowing origin: tamarind seed xyloglucan (XG, Mr >106 kDa) was kindly donated by Dr. Mayumi Shi-rakawa (Dainippon Pharmaceutical Co., Ltd., Osaka,Japan), beech wood β-D-glucuronoxylan (XYL) wasa gift from Dr. Peter Biely (Institute of Chemistryof SAS, Bratislava, Slovakia), hydroxyethylcellulose(HEC, medium viscosity) was purchased from Fluka(Switzerland), pustulan (β-(1,6)-D-glucan, PUS) wassupplied from Calbiochem (UK), barley (1,3;1,4)-β-D-glucan (MLG) and wheat arabinoxylan (AX) fromMegazyme (Ireland), laminarin (β-(1,3)-D-glucan) andlocust bean gum galactomannan from Sigma (USA).Xyloglucan nonasaccharide XLLG (for abbreviated

nomenclature of xyloglucan oligosaccharides see Fryet al., 1993) was prepared by digestion of XG withTrichoderma cellulase (Cellulysin, Calbiochem, USA)(Sulová et al., 1995). The nonasaccharide was sepa-rated from the mixture of xyloglucan-derived oligosac-charides on a Bio-Gel P-10 (Bio-Rad, CA, USA)column (1.0 cm × 100 cm). Pustulooligosaccharides(PUOS) were prepared by acid digestion (2 M TFA,100◦C, 20–90 min) of pustulan. Progress of the re-action was monitored by thin-layer chromatography(TLC on Silicagel 60 plates, Merck, Slovakia, de-veloped twice in butan-1-ol : ethanol : water (ϕr= 5 : 2 : 3), detection by spraying with sulphuricacid in ethanol (ϕr = 0.1) followed by heating at100◦C for 5 min). After hydrolysis, the oligosaccha-rides were purified by being passed through a column(2.4 cm× 105 cm) of Bio-Gel P-6 (Bio-Rad, CA, USA)eluted with water, the fractions corresponding to in-dividual oligomers were pooled and freeze-dried. Themolecular masses of the oligosaccharides were con-firmed by matrix-assisted laser desorption/ionisationtime-of-flight mass spectrometry (MALDI-TOF-MS)analyses performed on a Kratos Kompact MALDI 3(Shimazu, Japan) using 2,5-dihydroxybenzoic acid asa matrix.Oligosaccharides were coupled with sulphorho-

damine (SR) as previously described (Kosík & Farkaš,2008). The purity of individual SR-labelled oligosac-

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Fig. 1. TLC analysis of mixture of fluorescently labelled pus-tulooligosaccharides. Purified PUO3–SR, PUO5–SR,PUO6/7–SR, and SR were used as standards.

charides derived from XG and from PUS was checkedby TLC on silica gel plates using the butan-1-ol : ethanol : water (ϕr = 5 : 2 : 3) solvent system. Thechromatograms were viewed under the UV lamp. Themolar concentrations of the labelled oligosaccharideswere calculated on the basis of the molar absorptioncoefficient for LRSC, ε566 = 85000 M−1 cm−1 (Certifi-cate of Analysis for Lissamine rhodamine B sulphonylchloride, lot No. CM16-027. Anaspec, San José, CA,USA).The PUOS–SR used in this work represented a

mixture of oligosaccharides (DP 3–6) prepared bymixing the equimolar amount of labelled individuals.Its composition was checked by TLC on Silicagel 60plates (Merck, Slovakia), developed twice in butan-1-ol : ethanol : water (ϕr = 5 : 2 : 3). Purified PUO3–SR,PUO5–SR, PUO6/7–SR, and SR were used as stan-dards. The spots were visualised under a UV lamp(Fig. 1).The substrates were further characterised by

MALDI-TOF mass spectrometry. The measurementswere carried out using a 4800 Proteomics Analyser(Applied Biosystems, Framingham, USA). Typically,0.5 µL of derivatised saccharide (50 ng µL−1 in water)was mixed with 0.5 µL of α-cyano-4-hydroxycinnamicacid (5 mg mL−1 in acetonitrile : water (ϕr = 3 : 2)) onthe stainless steel sample spot and dried. The MS datawere acquired in the negative reflectron mode, cali-brated externally by measuring 6 standard peptides,and processed using Data Explorer software v.4.8 (Ap-plied Biosystems, Foster City, CA, USA). The compo-sition of the corresponding oligosaccharide ions, theirtheoretical m/z, experimental m/z and relative abun-dance are included in Table 1. The MS data character-ising XGO9–SR were published by Kosík et al. (2011).

Nasturtium (Tropaeolum majus, var. Goldshine or-ange) seeds were germinated in wet perlite at 20–22◦Cunder a natural day–night regime for 12 days. Theseparated cotyledons covered with seed coats were ho-mogenised with two volumes of an extracting buffercontaining 0.1 M imidazole, pH 6.0, and 1 M NaCl(4◦C, 12 h). After filtration through Miracloth (Cal-biochem, USA) and centrifugation (23000g, 20 min,4◦C), the extract obtained was allowed to precipitatewith ammonium sulphate to 90 % saturation (24 h,4◦C), centrifuged again, the sediment dissolved in asmall volume of 1 M NaCl solution, dialysed againstdistilled water, and freeze-dried. The preparation wasdenoted as a crude extract and stored at –20◦C untilfurther use.The activity assays were performed in 0.1 M am-

monium acetate, pH 5.7, at laboratory temperatureand at various concentrations of the acceptor sub-strate. The fluorimetric method (Fry, 1997) utilisedXG or HEC (0.3 %) as the donor (5 µL), XGO9–SR orPUOS–SR (1 µL, 2.72–27.2 µm) as the glycosyl accep-tor and a concentrated solution of nasturtium proteinextract (4 µL). At defined time intervals, the reactionswere stopped by the addition of an equal volume offormic acid (ϕr = 0.4) and 4 µL aliquots of the mixturewere applied in triplicates onto the Whatman 3MMpaper template corresponding in size and shape to 96-well microtitration ELISA plate, and dried with hotair. The unbound labelled oligosaccharides were re-moved by washing with ethanol (ϕr = 0.6) containingformic acid (ϕr = 0.05) (12 h), followed by repeatedwashing with ethanol (ϕr = 0.6). After drying, theintensity of fluorescence on the paper was quantifiedusing an ELISA micro-plate reader equipped with flu-orescent detector (Synergy HT 1, BioTek, Winooski,VT, USA). The filters used had parameters of λexc= 530 nm and λem = 575 nm. The built-in fluores-cence was converted into absolute amounts using thecalibration curve for sulphorhodamine.The scaled-up reaction mixtures incubated for var-

ious time intervals and the control (without enzyme)were analysed by HPLC using isocratic size-exclusionchromatography on a Hewlett Packard 1100 (PaloAlto, CA, USA) system with a fluorescent detectorcontrolled by ChemStation software (Agilent, SantaClara, CA, USA). The detector was programmed forexcitation at 530 nm and emission at 575 nm. Analyseswere performed on a TSKgel G3000 SWXL column,7.8 mm × 300 mm (TosoHaas, Tokyo, Japan) at 24◦C,eluted with 100 mM ammonium acetate, pH 5.7 con-taining aqueous acetonitrile (ϕr = 0.2) at a flow-rateof 0.5 mL min−1.The germinating nasturtium seeds were hand-

sectioned for in situ microscopic analysis. The pro-cedures for the in situ transglycosylation activity de-tection including the negative control were based onthe works of Vissenberg et al. (2000), Nishikubo etal. (2007), and Ibatullin et al. (2009) with modifica-

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Table 1. Characterisation of PUOS–SR mixture by MALDI-TOF mass spectrometry analysis

Compound Composition Calculated m/z Found m/z Relative abundance/%

PUO1–SR [C27H30N2O7S2 – H]− 720.227 720.218 1.59PUO2–SR [C39H53N3O16S2 – H]− 882.279 882.305 2.68PUO3–SR [C45H63N3O21S2 – H]− 1044.332 1044.363 29.9PUO4–SR [C51H73N3O26S2 – H]− 1206.385 1206.423 35.1PUO5–SR [C57H83N3O31S2 – H]− 1368.438 1368.478 25.4PUO6–SR [C63H93N3O36S2 – H]− 1530.491 1530.537 4.01PUO7–SR [C69H103N3O41S2 – H]− 1692.544 1692.591 1.32

Fig. 2. Dependence of enzyme activity on concentration of acceptor substrate: XGO9–SR (A) and PUOS–SR (B). 0.3 % XG usedas donor substrate in the dot blot assay.

tions. Tissue sections were incubated in solutions con-taining 27.2 µM XGO9–SR or PUOS–SR in 0.1 Msuccinate buffer, pH 5.5, for 1 h for XGOS–SR and6 h for PUOS–SR at laboratory temperature. The re-action was stopped by washing the tissue slices withethanol : formic acid : water (ϕr = 66 : 5 : 2) for about12 h followed by repeated washing with ethanol (ϕr= 0.6). The incorporation of oligosaccharides was ob-served by a fluorescent microscope Nikon Eclipse 80i(CFI 15 × 40 CFI Plan Fluor, Nikon, Tokyo, Japan)using an excitation wavelength of 550 nm and emissionwavelength of 570 nm and documented with DS-FilNikon camera (5 Mpx resolution).

Results and discussion

TLC of the PUOS–SR mixture showed fluores-cently labelled oligosaccharides with the degree ofpolymerisation (DP) between 3 and 6 with tracesof PUO7–SR and residual SR (Fig. 1). PUO3–SR,PUO4–SR, and PUO5–SR accounted for the major-ity share.MS analysis confirmed these expected structures

but PUO2–SR and traces of labelled monomer werealso found (Table 1). The reason for using the mixturerather than a defined oligosaccharide was the unknowninfluence of acceptor DP on the potential transglyco-sylation reaction.The protein extract of germinating nasturtium

seeds containing multiple forms of XET exhibitedtransglycosylating activity with the following sub-

strate pairs: XG–xyloglucan oligosaccharides (a typ-ical reaction catalysed by XET), XG–laminarioligo-saccharides, XG–(1,3;1,4)-β-glucooligosaccharides,XG–xylooligosaccharides, and XG–pustulooligosac-charides (gentiooligosaccharides) (Ait Mohand & Far-kaš, 2006; Stratilová et al., 2010) in the fluorescencedot blot assay with the SR-labelled acceptors (Kosíket al., 2011).The positive reaction of pustulooligosaccharides

(PUOS–SR) with XG was surprising because pustu-lan has not been known as a natural component ofdicotyledonous plant cell walls. This linear β-(1,6)-linked polysaccharide is a typical component of theyeast cell wall (Cabib et al., 2008). Nevertheless, theactivity of the enzymes with XGOS–SR was approxi-mately 100 times higher than with PUOS–SR (Fig. 2).The heterotransglycosylation activity of the ex-

tract from nasturtium seeds was confirmed by size-exclusion chromatography on HPLC using fluores-cence and RI detectors to monitor the substrate–product changes during transglycosylation (Fig. 3).It can also be seen that the reaction rates withXGO9–SR or PUOS–SR and XG differed by two or-ders of magnitude (Figs. 2, 3A, and 3B). Anotherdonor substrate, cellulose (in the form of soluble hy-droxyethylcellulose, HEC), was also able to incor-porate the PUOS–SR (Fig. 3D). The incorporationof XGOS–SR to HEC (Fig. 3C) was reported notonly with nasturtium but also with barley seed XET(Hrmova et al., 2007, 2009). The structural analogyof XG and HEC was considered as the main reason

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Fig. 3. Elution profile of reaction mixtures composed of nasturtium seed extract and substrate pairs: XG/XGO9–SR (A) (solidline, 0 h; dashed line (-) 0.5 h; dashed line (–) 16 h; dotted line 3 d), XG/PUOS–SR (B) (solid line, 0 h; dashed line (–) 2h; dashed line (-) 6 h; doted line 3 d), HEC/XGO9–SR (C) (solid line, 0 h; dashed line (-) 0.25 h; dashed line (–) 1.25 h),and HEC/PUOS–SR (D) (solid line, 0 h; dashed line (-) 2.5 h) on TSKgel G3000. SWXL column connected to HPLC. Theconcentrations of XGO9–SR and PUOS–SR were 27.2 µM, of XG, and HEC 0.3 %.

A B

C D

Fig. 4. Tissue slices from germinating nasturtium seeds incubated with XGO9–SR, 1 h (A, B) and PUOS–SR, 6 h (C, D) observedin a fluorescent microscope. Magnification 15 × 40 (A, C) and detail (B, D).

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for this donor non-specificity. Similarly, the HEC wascleaved with xyloglucan endohydrolase (Baumann etal., 2007).The comparison of heterotransglycosylation reac-

tion rates with the HEC–XGOS–SR and HEC–PUOS–SR pairs showed higher incorporation of XGOS thanof PUOS, again indicating the action of the ma-jor unspecific XET form of germinating nasturtiumseeds (Stratilová et al., 2010). The incorporation ofPUOS–SR in other polysaccharides, such as mixed-linked glucan, pustulan, laminarin, galactomannan,glucuronoxylan, or arabinoxylan, tested by HPLC,was not observed.Surprisingly, a microscopic comparison of incor-

poration of the labelled xyloglucan nonasaccharideXGO9–SR and PUOS–SR in cell structures clearlyshowed different binding sites. While XGO9–SR wasincorporated predominantly in the primary cell wall,the PUOS–SR was concentrated in the plasma mem-brane (Fig. 4). The binding of other labelled oligosac-charides with a positive reaction with XG (Stratilováet al., 2010) showed a similar pattern to XGO9–SR(results not shown). At present, we have no explana-tion for the different binding of XGO9–SR and PUOS–SR to cell structures. One reason could be the differentlocalisation of particular XET isoenzymes of differentsubstrate specificity.

Conclusions

The results described underline the importance ofthe simultaneous application of different methods forverifying the existence of newly-found heterotransgly-cosylating activities in plant extracts, as well as theimportance of assaying both in vivo and in vitro activ-ities. A primary action of defined unspecific XET incell walls leading to the incorporation of PUOS–SR instructural XG or HEC was indicated in one case. Be-cause of the systematic rebuilding of the cell wall, thePUOS–SR were quickly replaced with XG or XGOS–SR in this case. The primary binding of PUOS–SR tothe plasma membrane structures could be due to theirinteraction with nascent chains of cellulose, which issynthesised just in this structure (Carroll & Specht,2011). This assumption is supported by the inabil-ity of the nasturtium protein extract to catalyse theincorporation of PUOS–SR in other polysaccharidesexcept for XG and HEC. These findings indicate theexistence in nasturtium of XETs with different local-isation, function and substrate specificity.

Acknowledgements. The authors are grateful to Dr. Liškováand Dr. Erdelská, and Prof. Lux for identification of the nas-turtium tissue used. This contribution is the result of theproject implementation: Centre of Excellence for White-GreenBiotechnology, ITMS 26220120054, supported by the Research& Development Operational Programme funded by the ERDF(40 %). The financial support of VEGA grants Nos 2/0005/10,2/0020/12, 2/0046/10, support from the Institutional Research

Plan AV0Z40310501 of the Institute of Analytical Chemistry,v.v.i., Academy of Sciences of the Czech Republic and of the In-stitutional Research Plan No. MO0FVZ0000501 from the Fac-ulty of Military Health Sciences, University of Defence, CzechRepublic is also gratefully acknowledged.

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