XET Activity is Found Near Sites of Growth and Cell Elongation in ...

XET Activity is Found Near Sites of Growth and Cell Elongationin Bryophytes and Some Green Algae: New Insights into the Evolution

of Primary Cell Wall Elongation

VICKY S. T. VAN SANDT1, HERMAN STIEPERAERE3, YVES GUISEZ2,

JEAN-PIERRE VERBELEN1 and KRIS VISSENBERG1,*1Plant Physiology and Morphology, and 2Plant Physiology, Department of Biology, University of Antwerp,

Groenenborgerlaan 171, B-2020 Antwerp, Belgium and 3National Botanic Garden of Belgium,Domein van Bouchout, B-1860 Meise, Belgium

Received: 10 July 2006 Returned for revision: 24 August 2006 Accepted: 11 September 2006 Published electronically: 10 November 2006

† Background and Aims In angiosperms xyloglucan endotransglucosylase (XET)/hydrolase (XTH) is involved inreorganization of the cell wall during growth and development. The location of oligo-xyloglucan transglucosyla-tion activity and the presence of XTH expressed sequence tags (ESTs) in the earliest diverging extant plants, i.e.in bryophytes and algae, down to the Phaeophyta was examined. The results provide information on the presenceof an XET growth mechanism in bryophytes and algae and contribute to the understanding of the evolution of cellwall elongation in general.† Methods Representatives of the different plant lineages were pressed onto an XET test paper and assayed. XETor XET-related activity was visualized as the incorporation of fluorescent signal. The Physcomitrella genome data-base was screened for the presence of XTHs. In addition, using the 30 RACE technique searches were made forthe presence of possible XTH ESTs in the Charophyta.† Key Results XET activity was found in the three major divisions of bryophytes at sites corresponding to growingregions. In the Physcomitrella genome two putative XTH-encoding cDNA sequences were identified that containall domains crucial for XET activity. Furthermore, XET activity was located at the sites of growth in Chara(Charophyta) and Ulva (Chlorophyta) and a putative XTH ancestral enzyme in Chara was identified. No XETactivity was identified in the Rhodophyta or Phaeophyta.† Conclusions XET activity was shown to be present in all major groups of green plants. These data suggest thatan XET-related growth mechanism originated before the evolutionary divergence of the Chlorobionta and opennew insights in the evolution of the mechanisms of primary cell wall expansion.

Key words: XTH, XET activity, primary cell wall, xyloglucan, cell elongation, growth, Physcomitrella patens,bryophytes, Charophyta, Chlorophyta, Rhodophyta, Phaeophyta.

INTRODUCTION

A basic characteristic of vascular plants, mosses and manyalgae is the substantial post-mitotic increase in volume,known as expansion or elongation, preceding or coincidingwith cell differentiation. The mechanisms steering thisprocess are to date largely unknown (Hu et al., 2006). Aconstant feature, however, is the increase in volume of thevacuole and the increase in surface area of the primarycell wall. The wall is an indispensable and characteristicfeature of plant cells. Among many other functions(Albersheim, 1976; Carpita and Gibeaut, 1993) it serves tomaintain the shape of plant cells and thereby greatly con-tributes to the structural integrity and morphology of theentire plant. In angiosperms the primary cell wall (PCW)is composed of cellulose microfibrils that are embedded ina highly hydrated matrix of hemicelluloses, pectins andglycoproteins (Darvill et al., 1980; McNeil et al., 1984;Fry, 1986; McCann and Roberts, 1991; Carpita andGibeaut, 1993; Pauly et al., 1999). Xyloglucan (XyG) isthe major hemicellulose in non-gramineous plants and wasshown to be present in all vascular plants (Popper and Fry,2004). It can form non-covalent hydrogen bonds withcellulose (Hayashi et al., 1987) and is therefore believed

to tether adjacent cellulose microfibrils, thereby forming aload-bearing cellulose/XyG network in plant cell walls(Fry, 1989; McCann et al., 1990). During turgor-drivencell wall expansion this network needs to be modified toallow slippage of the cellulose microfibrils, but withoutlosing the integrity of the cellulose/XyG network as thismay lead to detrimental cell lysis (Marga et al., 2005).Two enzyme families are considered to play a majorpart in this highly controlled mechanism of cell wall loos-ening: expansins and xyloglucan endotransglucosylase/hydrolases (XTHs). Expansins are thought to disturb thehydrogen bonds between XyG and the cellulose micro-fibrils and hence create the possibility for slippage of thefibrils (McQueen-Mason and Cosgrove, 1994; Cosgrove,2000; Sampedro and Cosgrove, 2005). XTHs, by contrast,break (¼hydrolase or XEH action) or break and rejoin thetethering XyG with another available XyG (¼endotrans-glucosylase or XET action). This action also allows thereorganization of the XyG within the (growing) wall(Thompson and Fry, 2001), as the incorporation of newlysecreted XyG (Fry et al., 1992; Nishitani and Tominaga,1992; Rose et al., 2002). In angiosperms, XTHs wereindeed shown to play major roles during plant growthand differentiation (Campbell and Braam, 1999; Albertet al., 2004; Matsui et al., 2005; Vissenberg et al., 2005).* For correspondence. E-mail [email protected]

# The Author 2006. Published by Oxford University Press on behalf of the Annals of Botany Company. All rights reserved.

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Annals of Botany 99: 39–51, 2007

doi:10.1093/aob/mcl232, available online at www.aob.oxfordjournals.org

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As XTHs are so far the only characterized and well-described enzyme group displaying transglucosylationactivity in the plant cell wall, the study of their presenceand function in lower plants will shed new light on howthe PCW and the PCW elongation mechanism originated,evolved and functions today in the angiosperms.

In our previous work the presence of XET action wasdemonstrated in vivo in roots of representatives of all vas-cular plants down to one of the most primitive vascularland plant orders, the Selaginellales (Vissenberg et al.,2003). The conservation of the XTH amino acid sequenceand function throughout vascular plant evolution was alsodescribed in a detailed study of a heterologously expressedXTH of Selaginella kraussiana, Sk-XTH1 (Van Sandtet al., 2006). Like expansins (Sampedro et al., 2006),XTHs are thus part of an ancient cell wall reorganizingmachinery that originated even before the evolutionarydivergence of vascular plants (Vissenberg et al., 2003;Van Sandt et al., 2006).

As sequencing of the Physcomitrella patens genome isin progress searches were made for the presence ofPhyscomitrella XTH genes in silico and potential XTHsequences were analysed. Recently the primary cell wallcomposition of the bryophytes and charophytes wasdescribed (Popper and Fry, 2003). In accordance with thiswork the presence of XET activity in the three majorgroups of bryophytes and in charophytes was investigated,and some representatives of the chlorophytes, rhodophytesand phaeophytes were also included. Owing to autofluores-cence of the tissues, the in vivo assay used previously tostudy XET activity in roots (Vissenberg et al., 2000, 2001,2003, 2005), is unfeasible in bryophytes and algae. Toovercome this problem tissue prints on XET test paperswere used, as described by Fry (1997). The activity thusfound in vitro is merely indicative for its presence in vivo,as xyloglucans are absent in the Charophyta andChlorophyta (Popper and Fry, 2003). However thistechnique allows the localization of enzymes that arecapable of incorporating xyloglucan oligosaccharides intoa xyloglucan matrix and thus displaying XET activity.These enzymes can include XTHs or XTH-like enzymes.In accordance, searches were made for the presence of anXTH or XTH-like gene in one of the closest relatives ofthe vascular plants, Chara vulgaris. Combining these datawith recent findings on the phylogenetic relationship ofXTHs and family GH16 hydrolases, the possible originand evolution of the XTH–xyloglucan interacting mechan-ism itself as well as the evolution of PCW elongation ingeneral are discussed.

MATERIALS AND METHODS

Plant material

Physcomitrella patens (Hedw.) Bruch & Schimp. wasgrown in Petri dishes on minimal medium (Schaefer,2001) covered with cellophane discs. Plants were grown ina growth chamber (TCPS, Werchter, Belgium) at 26 8C,16-h light per day (Sylvania, cool white), with a photosyn-thetic photon flux of 50–80 mmol m22 s21. Seven-day-old

gametophytes were used to prepare RNA and tissuehomogenates.

Most bryophytes used for tissue printing were collectedin the wild in Hechtel-Eksel (Belgium), Phaeoceroscarolinianus (Michx.) Prosk. was collected in Boom(Belgium), Anthoceros agrestis Paton nom. cons. prop.near Ghent (Belgium), Chara vulgaris L. in the NationalBotanical Garden (Meise, Belgium) and marine algae werecollected in Wimereux (France). All bryophytes and algaewere directly processed in the lab upon arrival to minimizestress and changes in growth conditions.

Analysis of the Physcomitrella database

The Selaginella kraussiana XTH amino acid sequence,Sk-XHT1 (accession no. AY580314), was used in aprotein blast against the draft Physcomitrella patens data-base (http://www.cosmoss.org; Lang et al., 2005). Theretrieved sequences were analysed in expasy (http://www.expasy.org) and aligned using ClustalW (http://www.ebi.ac.uk/clustalw/).

3 0 RACE PCR

RNA from P. patens was prepared using a slightly modi-fied Plant Concert RNA protocol (Invitrogen). The RNAwas kept in 50 % isopropanol overnight, at 220 8C, toachieve optimal precipitation and a maximal extractionyield. Five micrograms of total RNA was reverse-transcribed using Superscript II RNase H-ReverseTranscriptase (Invitrogen) according to the manufacturer’sinstructions. The 30-end of the CAC43709 cDNA (http://www.cosmoss.org) was subsequently obtained by30 RACE using a gene-specific internal primer and anoligo-dT26. The full-length cDNA sequence of CAC43709was constructed in silico and subsequently amplified fromtotal cDNA using a specific primer set and a High Fidelity(Roche) proofreading enzyme mix.

Preparation of XET test paper

The XET and control test papers were made asdescribed by Fry (1997). A filter paper (Whatman no. 1)was dipped into a solution of 0.5 % 1-1-1trichloro-2-methylpropan-2-ol and 1 % 200-kDa tamarindxyloglucan, and dried overnight. The dried papers weresubsequently dipped into a 5 mM XLLGol-sulforhodamine(XGO-SRs) in 70 % acetone solution to become XETtest papers, or into a 5 mM trisaccharide-SR solution(¼non-XET substrate) to serve as control papers. Theywere all left to dry in the dark before use. Thetrisaccharide-SR preparation contained two main fluor-escent compounds (one major and one minor), both ofwhich ran on a silica-gel thin-layer chromatography plate(in BuOH/HOAc/H2O 2 : 1 : 1) between maltose-SR andmaltotriose-SR, suggesting that they are trisaccharideswith at least one residue smaller than Glc. The materialused for preparation of these SR derivatives was a minorfraction of small oligosaccharides obtained by cellulasedigestion of tamarind xyloglucan. Their exact structures

Van Sandt et al. — Study of XET Activity in Bryophytes and Algae40


are unknown, but one may possibly be Xyl-Glc-Glc. Theydo not act as XET acceptor substrates for a crude prep-aration of cauliflower enzymes (S. Fry, University ofEdinburgh, pers. comm.).

Dot-spot

Physcomitrella patens gametophytic tissue (7 g) washomogenized with several 15-s pulses (IKA Labortechnik,Staufen, Germany) in 300 mM sodium acetate, 20 mM

calcium chloride, 1 mM dithiothreitol at pH 5.5 adjustedwith sodium hydroxide (Iannetta and Fry, 1999). XETactivity was tested by spotting 5 mL of the homogenateonto both a control and an XET test paper, which werethen incubated in a sealed acetate envelope and kept over-night at room temperature. XET activity is defined as thetransfer of the non-labelled xyloglucans to fluorescentacceptor-substrates (XGO-SRs). Non-reacted fluorescentxyloglucan oligosaccharides and trisaccharides werewashed away after the assay in 90 % formic acid/ethanol/water (1 : 1 : 1) for 1 h followed by a 5-min wash in water.XET activity is seen as the remaining fluorescence on theXET test paper upon UV illumination. Images were takenusing an Olympus C-5050 ZOOM digital camera withidentical settings (1/10 shutter time, F 2.0 diaphragm) toallow comparison of the resulting fluorescence.

Tissue print

Both XET and control test papers used for the tissueprints were initially dipped into 25 mM MES bufferadjusted to pH 5.5. Fresh tissue of bryophytes and algaewas subsequently pressed onto the test papers (see above)and sealed into an acetate envelope. This ‘sandwich’ wasincubated in the dark at room temperature for 3 d under aconstant pressure of 3.2 g cm22 to ensure that the tissuewas kept in contact with the test papers. Pictures of theprint were taken before and after it was washed (asdescribed above), using daylight and UV illumination. Insome tissue prints, chlorophyll was transferred to the blot,especially at the sites were the tissue was cut. This con-taminant, however, was largely washed away in formicacid/ethanol/water. The remaining autofluorescence wasextremely weak and of a different colour than that of theSR substrates. It thus did not interfere with the obser-vations. The pigments of brown and red algae were not aseasily removed by washing and hence caused some pro-blems in analysing the presence of XET activity. Anoverlay of the tissue and fluorescence images allowed siteswith (high) leachable XET activity to be located exactly.

Searching a putative XTH or XTH-like transcript in Chara

RNA of Chara vulgaris was prepared and reversetranscribed as described for Physcomitrella above. Theresulting cDNA served as template in a PCR reactionusing a degenerate primer based upon variations presentin the catalytic domain of angiosperm XTHs and anoligo-dT26 primer [annealing temperature of 50 8C during45 s]. A 480-bp cDNA sequence was amplified, sequenced

and analysed in silico (http://www.expasy.org/tools/dna.html, http://elm.eu.org/). Its homology to XTHs and1,3-,1,4-b-D-endoglucanases was analysed using ClustalW(http://www.ebi.ac.uk/ clustalw/).

RESULTS

Putative XTH cDNAs in P. patens and XET activity

Blast analyses of the Physcomitrella genome with theamino acid sequence of the most primitive XTH thus farcharacterized, i.e. the lycopodiophyte XTH, Sk-XTH1(Van Sandt et al., 2006), identified one putative full-lengthXTH (CAC43710) and 17 fragmentary putative XTHESTs. An identical experiment using the differentArabidopsis XTH amino acid sequences gave an identicalset of hits. Some of the 18 hits differed in only a fewamino acids and probably originated from sequencingerrors. Seven ESTs were suggested to be part ofArabidopsis thaliana XTH precursor genes (Rensing et al.,2005; http://www.cosmoss.org). Yet only one EST(CAC43709; cDNA: AX172659) and one full-lengthsequence (CAC43710; cDNA: AX172661) included avariant of the catalytic domain, which is one of the criteriafor a gene to become annotated as ‘putative XTH’. ThecDNA sequence of AX172659 was completed using30 RACE PCR and the deduced amino acid sequenceswere analysed in silico.

Both moss amino acid sequences showed high homo-logy with known XTH amino acid sequences of numerousvascular plants. Most domains and some well-positionedamino acids, characteristic of vascular plant XTHs(Campbell and Braam, 1998; Johansson et al., 2003, 2004;Henriksson et al., 2003; Van Sandt et al., 2006), werefound in both moss cDNAs. Yet some notable differenceswere present in the amino acid composition of the cataly-tic domain. For clarity, in Fig. 1A both Physcomitrellaamino acid sequences were aligned with one representativevascular plant XTH, i.e. Sk-XTH1 from the lycopodio-phyte Selaginella kraussiana (Van Sandt et al., 2006). Asecretion signal at the N-terminus allowed both enzymesto be secreted in the apoplast (Fig. 1A, lower-case letters).The catalytic domain of both putative PhyscomitrellaXTHs (Fig. 1A, bold) differed in two (CAC43709,CEFDFEFLG) and three (CAC43710, YELDMEFLG)amino acids, respectively, compared with the ‘average’functional site DEIDFEFLG, conserved among mostXTHs of seed plants (Nishitani, 1997) and present infamily 16 glycoside hydrolases (Henrissat et al., 2001).However, both glutamic acid residues and the secondaspartic acid residue of the motif (xExDxExxx), whichplay a crucial role in the cleavage of the donor substrate infamily GH16 enzymes, were maintained. As in most vas-cular plant XTHs, the catalytic domain of both mossamino acid sequences was immediately followed by apotential N-glycosylation site (N-X-T/S, Shakin-Eshlemanet al., 1996) (Fig. 1A, dashed line). There was a notablevariation in the C-terminal part of both putativePhyscomitrella XTHs and in vascular plant XTHs.However, important residues involved in the maintenance

Van Sandt et al. — Study of XET Activity in Bryophytes and Algae 41


of three-dimensional stability (Fig. 1A, underlined) andthe recognition of the acceptor substrates in higher plantXTHs were also encoded by both moss cDNAs (Fig. 1A,superimposed). The conservation of most of the XTHidentity motifs made both Physcomitrella amino acidsequences putative XTH enzymes. Thus, the in silico datasuggested the presence of at least two potentialXTH-encoding cDNAs in the bryophyte P. patens.

A dot-spot of homogenized Physcomitrella tissue onXET test paper revealed a bright fluorescent spot inFig. 1B, resulting from the specific incorporation of fluor-escently labelled XyG oligosaccharides (X), whereas nofluorescence was seen on control test paper (C). This con-firmed the presence of at least one functional XTH inPhyscomitrella.

XET activity is present at growth sites in bryophytes

The presence of XET activity in other bryophytes wasassayed by tissue printing the gametophyte and/or sporo-phyte of 15 species, representing members of the threemajor groups of bryophytes, i.e. the Bryophyta, theAnthocerophyta and the Marchantiophyta (Table 1). Tolocalize specifically the sites of the tissue in whichenzymes, capable of xyloglucan transglucosylation, areexpressed, an image showing the tissue (left) and the

tissue print (right) is shown in Figs 2 and 3. Here the xylo-glucan transglucosylation, i.e. XET activity, was visible asan orange fluorescent spot under UV illumination.

The early diversification of the Bryophyta or mosses islinked to spore dispersion and this is reflected in the taxo-nomy of this group (Goffinet and Buck, 2004). Based onthe position of the gametangia and on the branching of thestem, however, two major groups can be distinguished,acrocarps and pleurocarps (Magdefrau, 1982; Buck andGoffinet, 2000). As growth patterns differ in the twogroups, XET activity was assayed in different acrocarp andpleurocarp species (Table 1, pleurocarpous mosses aredesignated with an asterisk). The expression of XET-displaying enzymes in the pleurocarp mosses correspondedto the site just beneath the apical meristem of the moss(Fig. 2A, see circle), whereas the other internal parts ofthe gametophyte displayed no visible XET activity on theprint. No incorporation of fluorescence was visible in thecontrol assay (Fig. 2B), indicating that the fluorescence inFig. 2A was indicative of the XET activity of XTHs orXTH-like enzymes. In acrocarp mosses the same patternof XET activity was visible when a young moss wasassayed (Fig. 2C, see circle). However, the XET signalnear the acrocarp apex was clearly weaker than thatobserved in pleurocarp mosses. This was probably causedby the high density of leaves covering the apex of most

FI G. 1. (A) Alignment of the amino acid sequence of a lycopodiophyte XTH, Sk-XTH1 and two Physcomitrella patens putative XTH amino acidsequences, CAC43709 and CAC43710. The alignment of the amino acid sequence of Sk-XTH1, CAC43709 and CAC43710 is shown in single lettercode. The predicted secretion signal peptide of each sequence is marked in lower-case letters, while the conserved catalytic site, shared with the activesite of the b-endoglucanase from Bacillus licheniformis, is presented with bold amino acid residues. The underlined (dashed) amino acids represent thepossible N-linked glycosylation sites. Conserved amino acids possibly involved in the stability of the enzyme and in acceptor-substrate recognition areunderlined. Tyrosines, putatively involved in the recognition of the acceptor substrate, are superscripted. (B) Dot-spot of homogenized Physcomitrellatissue on XET (left) and control (right) test paper. XET activity is seen as an orange spot on the XET test paper (left). As a control for specificity of

the assay, fluorescently labelled non-XET substrate was used, yielding no fluorescence when homogenized Physcomitrella tissue was spotted (right).



acrocarps, making it more difficult for the growing part tocome into contact with the test paper. Remarkably, noXET activity was visible near the apex when older acro-carp gametophytes, bearing sporophytes, were pressedonto the test paper (Fig. 2E). The remainder of both theyoung and the old acrocarp gametophytes did not displayXET activity (Fig. 2C, E), and nor did the control assays(Fig. 2D, F). Strong XET activity was, however, seen atthe sites where the maturing sporophyte capsule of bothacrocarp and pleurocarp mosses were pressed onto theXET test paper (Fig. 2E, see circle, example of an acro-carp moss). By contrast, young capsules showed no or

only very little XET activity (Fig. 2E, right). This differ-ence in XET activity, however, can be explained by thepresence of a calyptra, covering the young capsules. Oldercapsules completely lost this gametophytic cover (Fig. 2E,left), and were therefore in direct contact with the XETassay paper leaving a bright XET signal. No signal wasdetected at the site where the seta was pressed onto thepaper. In the control assay, the test paper revealed noincorporation of fluorescence (Fig. 2F).

The Sphagnales, or peat mosses, do not belong withinthe acrocarps or pleurocarps and are completely isolatedwithin the Bryophyta. The gametophyte of a peat moss

TABLE 1. List of the bryophyte and algal species assayed for XET activity

Family Species

BRYOPHYTAClass Sphagnopsida Sphagnaceae Sphagnum fimbriatum Wilson

Order SphagnalesSphagnum palustre L.

Class PolytrichopsidaOrder Polytrichales Polytrichaceae Atrichum undulatum (Hedw.) P.Beauv.

Class BryopsidaOrder Funariales Funariaceae Funaria hygrometrica Hedw.Order Dicranales Dicranaceae Dicranella heteromalla (Hedw.) Schimp.Order Bryales Mniaceae Mnium hornum Hedw.Order Hypnales Hypnaceae Brachythecium rutabulum (Hedw.) Schimp.*

Hypnum cupressiforme Hedw.*Vesicularia reticulata (Dozy & Molkenboer) Brotherus*

Thuidiaceae Thuidium tamariscinum (Hedw.) Schimp.*

ANTHOCEROTOPHYTAOrder Anthocerotales Anthocerotaceae Anthoceros agrestis Paton nom. cons. prop.

Phaeoceros carolinianus (Michx.) Prosk.

MARCHANTIOPHYTAClass Marchantiopsida Ricciaceae Riccia sp.

Order RiccialesOrder Fossombroniales Pelliaceae Pellia epiphylla (L.) Corda

Class Jungermanniopsida Geocalycaceae Lophocolea heterophylla (Schrad.) Dum.Order Jungermanniales

CHAROPHYTAClass Charophyceae

Order Charales Characeae Chara vulgaris L.

CHLOROPHYTAClass Ulvophyceae

Order Ulvales Ulvaceae Ulva linza L. (L.) J. Agardh. – E1Ulva lactuca L. – E2

Order Cladophorales Cladophoraceae Cladophora rupestris (L.) Kutzing – E3

RHODOPHYTAClass Florideophyceae

Order Gigartinales Gigartinaceae Chondrus crispus Stackhouse – G1Phyllophoraceae Gymnogongrus crenulatus (Turner) J. Agardh – G2Polyideaceae Polyides rotundus (Hudson) Greville. – G3

Order Gracilariales Gracilariaceae Gracilaria gracilis (Stackhouse) Steentoft, L.M. Irvine & Farnham – G4Order Plocamiales Plocamiaceae Plocamium cartilagineum (L.) Dixon – G5

HETEROKONTOPHYTAClass Phaeophyceae

Order Fucales Fucaceae Pelvetia canaliculata (L.) Decaisne & Thuret – I1Order Dictyotales Dictyotaceae Taonia atomaria (Woodward) J. Agardh – I2

Dictyota dichotoma (Hudson) Lamouroux – I3Order Laminariales Laminariaceae Lainaria saccharina (L.) Lmouroux – I4

* Pleurocarp mosses. Some algae are designated as in Fig. 3, e.g. E1, E2.References: Bryophytes: Buck and Goffinet (2000). Hornworts: Stotler and Crandall-Stotler (2005). Hepatics: Crandall-Stotler and Stotler (2000).

Charophyta and Chlorophyta: Lewis and McCourt (2004). Rhodophyta: Saunders and Hommers (2004). Heterokontophyta: Andersen (2004).



FI G. 2. Tissue print on XET (left) and control test paper (right) of representatives of the three major bryophyte lineages. XET activity is present on theXET test papers near the apex of the gametophyte of pleurocarpous Bryophyta: Thuidium sp. (A, encircled) and acrocarpous Bryophyta: Atrichumundulatum (Hedw.) P.Beauv. (C, encircled) and at the site of the mature sporophyte capsule: Atrichum undulatum (Hedw.) P.Beauv. (E). A morediffuse XET-generated signal is visible in Sphagnum fimbriatum (G). A faint spot of XET activity is present at the borders of the anthocerophyte game-tohytic thallus of Phaeoceros carolinianus (Michx.) Prosk (I, encircled). A bright spot of XET activity is present directly above the basal meristem, atthe site of cell elongation (J) of the sporophyte of Phaeoceros carolinianus (Michx.). This signal is only visible when the sporophyte was removedfrom the gametophyte and its protecting involucre (I,J). (K) Image of the elongation zone of the sporophyte; arrows point to short and longer cells (seelines representing the cross walls). In the gametophyte of both thalloid (Riccia sp., M) and leafy (Lophocolea heterophylla (Schrad.) Dum., O,encircled) Marchantiophyta an XET spot is visible near the apical cell(s). The images in B, D, F, H, L, N and P represent the fluorescence on

trisaccharide-SR control test papers of the XET assays on the left.



FI G. 3. Tissue print on XET (left) and control test paper (right) of representatives of the Charophyta, Chlorophyta, Rhodophyta and Phaeophyta. InChara vulgaris L. a fluorescent spot of XET activity is visible at the site where the apex and node are pressed onto the XET test paper (A). At highermagnification a clear XET signal is present at the sites of the branchlets (B). In the Chlorophyta XET activity is visible at the site of the holdfast ofUlva linza (Linnaeus) J. Agardh. in its tubular growth form (this species was previously named Enteromorpha linza) (E1, encircled). No signal wasfound in the other chlorohytes assayed: Ulva lactuca (E2) and Cladophora rupestris (L.) Kutzing (E3). In the assays of both Rhodophyta [G1:Chondrus crispus Stackhouse, G2: Gymnogongrus crenulatus (Turner) J. Agardh, G3: Polyides rotundus (Hudson) Greville, G4: Gracilaria gracilis(Stackhouse) Steentoft, L.M. Irvine & Farnham, G5: Plocamium cartilagineum (L.) Dixon] and Phaeophyta [I1: Pelvetia canaliculata (L.) Decaisne &Thuret, I2: Taonia atomaria (Woodward) J. Agardh, I3: Dictyota dichotoma (Hudson) Lamouroux, I4: Laminaria saccharina (L.) Lamouroux] a sub-stantial amount of autofluorescence is visible, but no XET signal is present on the XET test papers. On the right control prints on trisaccharide-SR test

papers of each group are shown (C, D, F, H and J).



displayed a more widespread XET signal, from the capitu-lum down to the youngest branches that are still stretchingout to their full length (Fig. 2G, H); this in contrast to theother groups of the Bryophyta.

Similar results were obtained in the Anthocerophyta.Pressing the gametophyte onto the test paper, a faint, butclear incorporation of XGO-SRs was visible at the edgesof the thallus (Fig. 2I, see circles), but absent from centralparts of the gametophyte (Fig. 2I). When the sporophyteswere embedded in the gametophyte and their basiscovered by the involucre, no XET signal was visible(Fig. 2I), but when removed from the gametophyte, thesporophyte foot generated a clear spot of XET activity onthe test paper (Fig. 2J, see circle). This site of high XETactivity corresponded to the zone above the basal meristemof the sporophyte where the epidermal cells elongate fromshort cells (arrow) to elongated cells (dashed arrow), ascan be seen in several cell files at high magnification(Fig. 2K). Some transverse cell walls were marked with aline. No incorporation of fluorescence was visible in thecontrol test paper impregnated with trisaccharide-SR(Fig. 2L).

In the Marchantiophyta both leafy and thalloid liver-worts were assayed. In both groups the gametophytesshowed clear XET activity. In thalloid liverworts anintense XET signal was visible at the edges of the gameto-phyte thallus, as seen in a Riccia species (Fig. 2M), but noXET activity was found at central parts of the gameto-phyte. In leafy liverworts, XET activity was seen near thesite of the gametophyte apex (Fig. 2O, see circle). Theremainder of the gametophyte showed no XET activity onthe test paper (Fig. 2N, P). Sporophytes were not assayedin marchantiophytes.

XET activity is present in green algae

The presence of extractable XET activity was alsostudied in the Charophyta. In simple charophytic algae,such as the Zygnematales, cell division is often notlocalized into meristematic regions. Also, the anatomy ofthese organisms is not particularly suitable for tissue print-ing. Therefore, Chara vulgaris was analysed. This specieshas a highly organized thallus with clear spatial segre-gation of meristematic activity and cell expansion. InChara clear XET activity was visible at the site where theapex was pressed onto the XET test paper, but also at sitescorresponding to other parts of the gametophyte (Fig. 3A,see circles). A clear fluorescent signal was present at thesite of the node and at the sites where the branchletswere pressed onto the test paper (Fig. 3B, see circle).The control assays confirmed the specificity of theXET-generated signal (Fig. 3C, D). The cell walls ofChara therefore clearly contain enzymes that act upon theexogenous xyloglucan donor and recognize the XyGacceptor substrates on the XET test paper.

Studying the other major lineage of green plants, theChlorophyta, the presence of enzymes displaying XETactivity was found in the ulvophycean algae Ulva linza,previously known as Enteromorpha linza. At the base,where the foot is attached to substrate, fluorescently

labelled xyloglucan was clearly incorporated in the testpaper (Fig. 3E, see circle). The absence of incorporationof non-XET substrates (Fig. 3F) confirmed the specificityof the XET activity in Chlorophyta. Other chlorophytes,however, displayed no XET activity (Fig. 3E).

Different representatives of the Rhodophyta (Fig. 3G)and Phaeophyta (Fig. 3I) were also tested for XETactivity. A substantial amount of autofluorescence wasvisible, both in experiments and in controls, hamperingthe detection of XET activity. However, none of thespecies assayed appeared to show a signal of XET activityon the prints (Fig. 3H, J).

Searching for an XTH or an XTH-like enzyme inChara vulgaris

The fluorescence found on the XET tissue print ofChara vulgaris indicated that there are enzymes expressedin the charophycean cell wall that are able to transglucosy-late xyloglucans in vitro. Based upon these findings RNAwas prepared and searches were made for the presence ofXTH or XTH-like transcripts in Chara. 30 RACE using adegenerate primer, based upon the variations present in thecatalytic domain of vascular plant XTHs, resulted in theamplification of a 480-bp fragment, named Chara2. Toidentify its homology with angiosperm XTHs, the deducedamino acid sequence was analysed in silico. As the evol-utionary connection between XTHs and (1,3-1,4)-b-D-glucan endohydrolases was recently demonstrated byStrohmeier et al. (2004), the relationship of Chara2 withboth enzyme groups was studied in an alignment(ClustalW) including two monocot XTHs of barley(HvXTH3, HvXTH4), two dicot XTHs of Arabidopsis(AtXTH4, AtXTH22) and two family 16 (1,3-1,4)-b-D-glucan endohydrolases (Fig. 4). Both groups ofenzymes shared a homologous catalytic site (shown initalics), which in most XTHs was immediately followedby an N-linked glycosylation site (underlined). Strohmeieret al. (2004) stated that one of the key differences betweenXTHs and (1,3-1,4)-b-D-glucan endohydrolases is theinsertion of three amino acids in XTHs, i.e. the PYX motif(boxed in Fig. 4). This motif is absent in the Chara2sequence and in the two (1,3-1,4)-b-D-glucan endohydro-lases; another key difference is the substitution of a meth-ionine in (1,3-1,4)-b-D-glucan endohydrolases by anaromatic amino acid residue, which is generally a tyrosinein XTHs. Interestingly, this substitution is present inChara2 (Fig. 4, arrow). In addition, the sequence ofChara2 shaded in grey shows more homology with thatof the corresponding acceptor binding loop sequence ofXTHs than do the endoglucanase sequences (Fig. 4, accep-tor binding loop is shaded in grey, homologous aminoacids are marked in white).

DISCUSSION

Thus far, XET action has been shown to be present in allvascular plants from the very ‘primitive’ lycopodiophyteSelaginella kraussiana up to ‘more evolved’ angiosperms(Vissenberg et al., 2003). Little is known about the



presence of cell-wall-modifying enzymes in even ‘earlier’land plants and algae. Thus far, extractable XET activitywas detected in the liverwort Marchantia and the mossMnium (Fry et al., 1992), a few partial putative XTHsequences were revealed in the Physcomitrella patensgenome (Rensing et al., 2005; http://www.cosmoss.org)and expansin sequences were found in mosses as well(Schipper et al., 2002; Yi et al., 2002). To our knowledge,no data concerning the presence of XET activity in algaehave been reported to date. To explore further the presenceof XET activity, a broad range of bryophytes were exami-ned and the potential origin of XET activity in green, redand brown algae was studied (Fig. 5).

The Bryophyta, Anthocerophyta and Marchantiophytarepresent the oldest lineages among extant land plants andtogether they form the bryophytes (Fig. 5). Given that thePhyscomitrella genome is in the process of beingsequenced, the XET search started with the Bryophyta,blasting the draft database (http://www.cosmoss.org) withSk-XTH1 (Van Sandt et al., 2006). This blast resultincluded all hits that were also obtained when blastingwith the different Arabidopsis XTHs. At least two of theresulting ESTs included all motifs that are essential forXET activity in vascular plants (Campbell and Braam,

1998; Henriksson et al., 2003; Johansson et al., 2003,2004; Van Sandt et al., 2006). A functional XET assay onPhyscomitrella homogenate was performed to prove thepresence of at least one functional XTH protein. XTH andits XET function are thus present in Physcomitrella patensand they are possibly encoded by a multi-gene family. AnXTH-related cell-wall-modifying machinery is thus prob-ably present throughout the mosses and perhaps even inevolutionary more primitive phyla.

Specific XET activity was detected in other Bryophytaand in the Anthocerophyta and Marchantiophyta.Furthermore, in both the gametophyte and the sporophyteof the Bryophyta and Anthocerophyta and in the gameto-phyte of the Marchantiophyta, a clear correlation betweenthe site of growth and the presence of XET activity wasdemonstrated. In the Bryophyta the pattern of XET activityin acrocarp mosses corresponds nicely with the develop-mental stage of the apical cell. A fluorescent XET signalis only present when a young gametophyte is pressed ontothe test paper. In older acrocarps the apical cell is used toform a gametangium, causing the gametophyte to ceasegrowth. In accordance, no XET activity was found at thesite of the gametophyte apex in sporophyte-bearing game-tophytes. Another illustration of the correlation between

FI G. 4. Alignment of the amino acid sequence of Chara2 with two monocot XTHs of barley (HvXTH3, HvXTH4), two dicot XTHs of Arabidopsis(AtXTH4, AtXTH22) and two family 16 (1,3-1,4)-b-D-glucan endohydrolases. The catalytic site shared by XTHs and (1,3-1,4)-b-D-glucan endohydro-lases is shown in italic and the PYX motif, a key difference between XTHs and (1,3-1,4)-b-D-glucan endohydrolases is boxed. The substitution ofmethionine in (1,3-1,4)-b-D-glucan endohydrolases to an aromatic amino acid residue, mostly a tyrosine, in XTHs is marked with an arrow, while theXTH acceptor binding loop and corresponding sequences in the Chara2 and (1,3-1,4)-b-D-glucan endohydrolases are shaded in grey. Homologous

amino acids are marked in white.



XET activity and growth was found in the sporophytes ofthe Anthocerophyta. These land plants have a near basalmeristem (Renzaglia and Vaugh, 2000; Shaw andRenzaglia, 2004), and XET activity occurs specifically atthe elongation site above the basal meristem of the sporo-phyte. In liverworts, considered to be the most ancientplant lineage (Kenrick and Crane, 1997; Bateman et al.,1998), a clear correlation between XET activity and thesites of growth was observed.

Although significant differences exist in cell wall com-position of the major land plant lineages, xyloglucan wasfound in the primary cell wall of all land plants, includingthe bryophytes (Popper and Fry, 2003, 2004). The conser-vation of the XET substrate throughout land plant evol-ution fits nicely with the conservation of XET functionwithin land plants. It is therefore likely that XET activityis part of an ancient cell-wall-modifying machinery thatoriginated even before the divergence of land plants.

The descent of the embryophytes from a charophyteancestor (Fig. 5) is among others supported by the pre-sence of cellulose-synthesizing rosettes in both groups(Hotchkiss and Brown, 1987). Charophytic algae wereshown to lack xyloglucan (Popper and Fry, 2003).Remarkably, an unambiguous XET signal was presentnear to or below the meristematic cells of both apex andbranchlets of the Chara vulgaris tissue print. Althoughxyloglucan was found to be absent in charophycean algae

(Popper and Fry, 2003), some Chara cell wall enzymesseem to be able to catalyse the incorporation of xyloglucanoligosaccharides into the xyloglucan matrix on the testpaper, and thus display XET activity. Recent studies havedemonstrated a structural connection between XTHs andxylan endohydrolases. Based upon these findings it wassuggested that XTHs could be active not only on cell wallxyloglucan but also on xylans (Strohmeier et al., 2004;Nishitani and Vissenberg, 2006). Interestingly, the enzy-matic digests of the Chara AIR (alcohol insoluble residue)resulted in products corresponding with the oligosacchari-des of xylan (Popper and Fry, 2004). The presence ofgenes coding for XTHs or XTH-like enzymes that in vivotransglucosylate xylans or other hemicelluloses in Charawas therefore studied. 30 RACE, using a degenerate primerbased upon the variations present in angiosperm XTHs,resulted in the amplification of a 480-bp fragment, namedChara2. To study its homology with XTHs, Chara2 wasaligned with angiosperm XTHs (monocots and dicots). Asrecent data suggest an evolutionary link between1,3-1,4-b-D-endoglucanases and XTHs (Strohmeier et al.,2004), 1,3-1,4-b-D-endoglucanases were included inthe alignment as well. In addition to a homologousthree-dimensional topology of the active site aminoacids, as seen for XTHs and endoxylanases,1,3-1,4-b-D-endoglucanases share the same amino acidcomposition with XTHs. Both enzyme families are

FI G. 5. Schematic view of part of the tree of life. The branching order of Phaeophytes, Rhodophytes and the green plants are based upon current DNAsequence data (Palmer et al., 2004). The phylogenetic relationships within the Chlorophyta and Charophyta are according to Lewis and McCourt(2004) and Graham et al. (2000). The branching order of the bryophytes remains uncertain (Shaw and Renzaglia, 2004). Note that the phaeophytesform a separate lineage, completely separate from the rhodophytes and green plants. Lineages studied for the presence of XET activity are shown in

black. Those exhibiting XET activity are underlined.



therefore thought to share a common ancestor (Strohmeieret al., 2004). Aligning Chara2 with representatives of bothenzyme families revealed that Chara2 shares featuresof both XTHs and 1,3-1,4-b-D-endoglucanases. Thekey differences between XTHs and 1,3-1,4-b-D-endoglucanases are the presence of a PYX motif in XTHsand the substitution of a methionine in 1,3-1,4-b-D-endoglucanases by a tyrosine in XTHs (Strohmeier et al.,2004). Chara2 has one of the two possible substitutions(M! Y), positioning it between both groups. The PYXmotif, however, is not necessary for the XET function, asAtXTH4, an XTH that was shown to display XET activity,lacks this motif (Campbell and Braam, 1999). Anadditional difference between XTHs and1,3-1,4-b-D-endoglucanases is a region in the protein thatforms the acceptor binding loop in XTHs. In Chara2 thisregion is more similar to that of XTHs than in1,3-1,4-b-D-endoglucanases (Fig. 4, shaded). Again, thispositions Chara2 between both groups of enzymes.Together these data suggest that Chara2 is possibly theC-terminal end of an enzyme that is an intermediatebetween 1,3-1,4-b-D-endoglucanases, present in microbialorganisms, and XTHs, seen in vascular plants. This is alsoreflected in the positioning of the Chara2 amino acidsequence within XTHs and 1,3-1,4-b-D-endoglucanases ina phylogenetic tree (data not shown). The Chara XTH-likeenzyme and its interacting donor substrate therefore prob-ably evolved together to an optimal enzyme–substrateinteracting model as is seen in higher plants. The evol-ution to a XyG/XTH interacting mechanism was possiblyone of the crucial events allowing the development of landplants.

Remarkably, a tissue print of Ulva linza, an ulvophy-cean alga belonging to the Chlorophyte lineage of thegreen plants, also showed a clear XET signal. The fluor-escent spot corresponded to a small region above the hold-fast of the algae. This holdfast is formed by the basal celldividing into 3–4 holdfast cells, which elongate andundergo further division (Kim et al., 1991). A clear corre-lation between cell elongation and transglucosylationactivity is therefore even found in the Chlorophyte lineageof green plants. Analysis of the cell wall components fromUlva lactuca and Ulva rigida suggested the presence ofcellulose microfibrils associated with XyG-like polysac-charides (Lahaye et al., 1994). The structure of this sul-fated glucuronorhamnoxyloglucan (ulvan) is, however,markedly different from that of higher plants (Lahaye andRay, 1995). In other studies a 1,3-1,4-b-D-glucan endohy-drolase digest of the Ulva AIR was shown to containglucose and xylose (Popper and Fry, 2003), indicating thepresence of a mixed linkage glucan with xylose substi-tutions in the Ulva cell wall. In accordance with the find-ings in the Charophyta, an ancient chlorophyte XTH-likeenzyme possibly interacts with the mixed linkagexyloglucan-like polysaccharide as donor substrate in Ulvalinza. Again these findings are supported by the evolution-ary connection between 1,3-1,4-b-D-glucan endohydro-lases and XTHs.

It is possible that in the green algae transglucosylatingenzymes with a broader substrate range were/are present

that gave rise to the XTHs with a higher substrate speci-ficity. Recently the reverse story was suggested in mono-cots, where XTHs are thought to have lost their substratespecificity and now transglucosylate 1,3-1,4-b-D-glucansas well during growth and cell elongation (Strohmeieret al., 2004). Similar to Chara2, these monocot XTHs(HvXTH4) miss some typical XTH features which poss-ibly play a role in narrowing the substrate specificity(Fig. 4, arrows) and have a less conserved amino acidcomposition of their acceptor binding loop (Fig. 4,shaded). This could be proof of the close relationship andpossible interconversions of XTHs and1,3-1,4-b-D-endoglucanases by small mutations. Inaddition, a mannan transglycosylase was detected inseveral plant species (Schroder et al., 2004), but nosequences of responsible enzymes are yet known.

In contrast to Ulva linza, no XET activity was found inUlva lactuca. This difference in the presence of XETactivity is striking. It can be explained by the fact thatboth organisms have a distinct morphology and growthpattern, being either tubular or planar. The Ulva lactucaspecimen studied had a planar form where growth occursin the entire plant body (http://www.algaebase.org). XETactivity would therefore probably cause a very diffuse andhence undetectable XET signal, whereas growth and XETactivity is more concentrated and detectable in the holdfastof the tubular growth form, as found in Ulva linza. Theabsence of a detectable XET signal in Cladophora rupes-tris, another Ulvophycean, supports this suggestion as theslender filaments could leave only a very faint and thusinvisible XET signal on the test paper. This limitation ofthe technique to visualize XET activity in filamentous orunicellular organisms, as mentioned above, preventedfurther analysis of the presence of XET-related transgluco-sylation activity in the other major groups of chlorophytesand charophytes.

The finding of XET-related activity in a chlorophyticspecies opens new ideas on the understanding of the evol-ution of green plants and gives new insights into thedevelopment of their cell wall. The XTH-modifyingmachinery therefore probably originated in the ancestorsof both charophytes and chlorophytes, before the split ofthe green plants. The presence of XyG and preference ofthis hemicellulose as an XTH substrate, as seen in dicots,is probably the result of selecting an optimal enzyme–substrate interacting mechanism to allow efficient cell wallelongation. A future strategy allowing the study ofXET-related activity in thread-like and unicellular organ-isms, such as Mesostigma (see Fig. 5), could furtherclarify the phylogenetic relationship of the charophytesand chlorophytes and would reveal more details on theevolution of XET activity in green plants.

The presence of XET activity in the major lineages ofthe green plants raised the question of the presence ofXET-related growth mechanism in red and brown algae.No XET activity was detected on tissue prints of differentred and brown algae. The colour of the autofluorescencecaused by algae pigments differed enough from that of theSR substrates to allow the detection of possible XETactivity. Furthermore, the absence of XET activity is in



complete agreement with the composition of their primarycell walls. Although both phaeophytes and rhodophytescontain cellulose fibrils, no homologue of XyG, mixedlinkage-b-D-glucans and xylans can be found within theircell wall, suggesting another mechanism for cell wallexpansion (Graham and Wilcox, 2000).

In summary, the data presented herein have revealed aclear correlation between growth (or cell elongation) andthe presence of XET activity in all three major groups ofbryophytes, and the presence of at least two potentialXTH-encoding cDNAs in the Physcomitrella genome hasbeen demonstrated. For the first time, it has been shownthat XET activity is also present at sites of growth inCharophyta and Chlorophyta, suggesting that XET origi-nated even before the evolutionary divergence of theChlorobionta. In accordance, part of a transcript in Charathat possibly encodes an ancestral XTH enzyme wasidentified, and the structural and evolutionary link betweenXTHs, endo-xylanases and 1-3,1-4-b-D-endoglucanaseswas discussed, explaining the substrate-tolerant behaviourof this ancient transglucosylating enzyme. As no XETactivity was detected in the Phaeophyta and Rhodophyta,XET activity is probably a feature unique to green plants.

ACKNOWLEDGEMENTS

V.V.S. is funded by a PhD grant of the Institute for thePromotion of Innovation through Science and Technologyin Flanders (IWT – Vlaanderen). K.V. is a PostdoctoralFellow of the Fund for Scientific Research – Flanders(FWO – Vlaanderen). This research was partially fundedby a University of Antwerp grant (UA-BOF) and a grantfrom the Fund for Scientific Research – Flanders (FWO –Vlaanderen), grant G.0101.04. We thank S. C. Fry for thelabelled oligosaccharides, D. De Beer for his help infinding Phaeoceros carolinianus, Professor S. Hoste,Chairman of the Flemish Study group of Bryology andLichenology, for his help in the collection of Anthocerosagrestis, Professor W. De Smet for the collection ofmarine algae and the Fund for Scientific Research –Flanders (FWO – Vlaanderen) for financial support.

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XET Activity is Found Near Sites of Growth and Cell Elongation in ...

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