plant and microbial xyloglucanases: structure and phylogeny405550/fulltext01.pdf · 2011-03-22 ·...

Plant and microbial xyloglucanases:

Function, Structure and Phylogeny

Jens Eklöf

Doctoral thesis

Royal Institute of Technology

School of Biotechnology

Division of Glycoscience

Stockholm 2011

Plant and microbial xyloglucanases: Function, Structure and Phylogeny

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ISBN 978‐91‐7415‐932‐5

ISSN 1654‐2312

Trita‐BIO Report 2011:7

©Jens Eklöf, Stockholm 2011

Universitetsservice US AB, Stockholm

Jens Eklöf

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Jens Eklöf (2011): Plant and microbial xyloglucanases: Function, Structure and Phylogeny School of Biotechnology, Royal Institute of Technology (KTH), Stockholm, Sweden

Abstract

In this thesis, enzymes acting on the primary cell wall hemicellulose xyloglucan are studied.

Xyloglucans are ubiquitous in land plants which make them an important polysaccharide to utilise for

microbes and a potentially interesting raw material for various industries. The function of

xyloglucans in plants is mainly to improve primary cell wall characteristics by coating and tethering

cellulose microfibrils together. Some plants also utilise xyloglucans as storage polysaccharides in

their seeds.

In microbes, a variety of different enzymes for degrading xyloglucans have been found. In this thesis,

the structure‐function relationship of three different microbial endo‐xyloglucanases from glycoside

hydrolase families 5, 12 and 44 are probed and reveal details of the natural diversity found in

xyloglucanases. Hopefully, a better understanding of how xyloglucanases recognise and degrade

their substrate can lead to improved saccharification processes of plant matter, finding uses in for

example biofuel production.

In plants, xyloglucans are modified in muro by the xyloglucan transglycosylase/hydrolase (XTH) gene

products. Interestingly, closely related XTH gene products catalyse either transglycosylation (XET

activity) or hydrolysis (XEH activity) with dramatically different effects on xyloglucan and on cell wall

characteristics. The strict transglycosylases transfer xyloglucan segments between individual

xyloglucan molecules while the hydrolases degrade xyloglucan into oligosaccharides. Here, we

describe and determine, a major determinant of transglycosylation versus hydrolysis in XTH gene

products by solving and comparing the first 3D structure of an XEH, Tm‐NXG1 and a XET, PttXET16‐

34. The XEH activity was hypothesised, and later confirmed to be restricted to subset of the XTH

gene products. The in situ localisation of XEH activity in roots and hypocotyls of Arabidopsis was also

visualised for the first time. Furthermore, an evolutionary scheme for how XTH gene products

developed from bacterial β‐1,3;1,4 glucanases was also presented based on the characterisation of a

novel plant endo‐glucanase, PtEG16‐1. The EG16s are proposed to predate XTH gene products and

are with activity on both xyloglucan and β‐1,3;1,4 glucans an “intermediate” in the evolution from β‐

1,3;1,4 glucanases to XTH gene products.

Keywords, xyloglucan, XTH, XET, XEH, xyloglucanase, plant cell wall, phylogeny, GH16, endo‐glucanase


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Sammanfattning

I den här avhandlingen undersöks enzymer som modifierar hemicellulosan xyloglukan. Xyloglukaner

finns i alla landlevande växter där de och binder till, täcker och korslänkar cellulosafibriller i den

primära cellväggen. Funktionen av xyloglukan är att spatialt separera cellulosafibriller, men ändå

behålla cellväggens styrka. I cellväggen modifieras xyloglukan av enzymer från xyloglukan

transglykosylas/hydrolas (XTH) genfamiljen. Dessa närbesläktade enzymer, som antingen katalyserar

transglykosylering (XET‐aktivitet) eller hydrolys (XEH‐aktivitet) av xyloglukan har vitt skilda effekter

på xyloglukan och därför också på cellväggens egenskaper. Strikta transglykosylaser utbyter

xyloglukanbitar mellan olika xyloglukanmolekyler medan hydrolaserna klyver ner xyloglukan till

oligosackarider.

Mikrober som lever på och bryter ner växtmaterial och således också xyloglukan har utvecklat olika

typer av xyloglukanaser. Tre mikrobiella xyloglukanaser, från glykosidhydrolasfamiljerna 5, 12 och

44, vars 3D‐strukturer och biokemiska karaktärisering demonstreras i denna avhandling påvisar en

del av den naturliga variationen av xyloglukanaser. En bättre förståelse av dessa enzymer och hur de

känner igen sina substrat kan förhoppningsvis leda till förbättrad nedbrytning av växtmaterial för

t.ex. biobränslen.

Även växter är i behov av xyloglukanaser. Den första 3D‐strukturen av en XEH, Tm‐NXG1 kopplar en

strukturell skillnad, mellan transglykosylaser och hydrolaser, till en funktionell skillnad, och kan

därigenom lokalisera hydrolas aktivitet i XTH‐genprodukter till några väl avgränsade enzymer.

Vävnadsspecificiteten i XEH‐aktivitet hos modellorganismen backtrav, Arabidopsis thaliana,

demonstreras även för första gången. Dessutom, föreslås hur XTH‐genprodukterna har utvecklats

från bakteriella β‐1,3;1,4‐glukanaser genom upptäckten och karaktäriseringen av en ny typ av växt‐

endo‐glukanas, PtEG16‐1. PtEG16‐1 indelas i en ny grupp av växtenzymer som tros vara äldre än XTH‐

genprodukter och har både β‐1,3;1,4‐glukanas aktivitet och xyloglukanas aktivitet samt har likheter i

aminosyrasekvens med både bakteriella β‐1,3;1,4‐glukanaser och XTH‐genprodukter.

Jens Eklöf

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List of publications included in thesis

I Ariza A*, Eklöf JM*, Spadiut O*, Offen WA, Roberts SM, Wilson KS, Brumer H, Davies GJ Structure and Activity of a Paenibacillus polymyxa Xyloglucanase from Glycoside Hydrolase Family 44. (Manuscript)

II Gloster TM, Ibatullin FM, Macauley K, Eklöf JM, Roberts S, Turkenburg JP, Bjørnvad ME, Jørgensen PL, Danielsen S, Johansen KS, Borchert TV, Wilson KS, Brumer H, Davies GJ (2007) Characterization and Three‐dimensional Structures of Two Distinct Bacterial Xyloglucanases from Families GH5 and GH12. J. Biol. Chem. 282: 19177‐19189

III Baumann MJ, Eklöf JM, Michel G, Kallas AM, Teeri TT, Czjzek M, Brumer H, III (2007) Structural Evidence for the Evolution of Xyloglucanase Activity from Xyloglucan Endo‐Transglycosylases: Biological Implications for Cell Wall Metabolism. Plant Cell 19: 1947‐1963

IV Kaewthai N*, Eklöf JM*, Gendre D*, Ibatullin FM, Ezcurra I, Bhalerao RP, Brumer H Group III‐A XTH Genes Encode Predominant Xyloglucan endo‐hydrolases Active in Expanding Tissues of Arabidopsis thaliana. (Manuscript)

V Maris A, Kaewthai N*, Eklöf JM*, Miller JG, Brumer H, Fry SC, Verbelen JP, Vissenberg K

(2011) Differences in Enzymic Properties of Five Recombinant Xyloglucan endotransglucosylase/hydrolase (XTH) Proteins of Arabidopsis thaliana. J. Exp. Bot. 62: 261‐271

VI Eklöf JM, Brumer H, An endo β‐1,4 glucanse, PtEG16‐1 from black cottonwood (Populus trichocarpa) represents an evolutionary link between bacterial lichenases and XTH gene products. (Manuscript)

*Authors contributed equally to the work


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Author’s contributions

Paper I: Performed part of the experimental work primarily on polysaccharides and inhibition

studies. I also made significant contributions to the writing of the paper.

Paper II: Performed the experimental work concerning polysaccharide specificity and pH optimum.

Paper III: Took part in almost all parts of the paper except the crystallography. The experimental in

vitro work was done together with Dr. Martin Baumann and the phylogenetic analyses were

performed together with Dr. Gurvan Michel.

Paper IV: Performed the production, purification and characterisation AtXTH31 in vitro.

Paper V: Performed the production, purification and characterisation AtXTH13 in vitro and

contributed in the writing of the paper.

Paper VI: This work was performed by me; from idea, to cloning and characterisation and

phylogenetic analyses.

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Related publications not included in this thesis

1. Eklöf JM, Brumer H (2010) The XTH Gene Family: An Update on Enzyme Structure, Function, and Phylogeny in Xyloglucan Remodeling. Plant Physiol. 153: 456‐466 (Included, in part as a chapter in the introduction.)

2. Eklöf JM*, Tan TC*, Divne C, Brumer H (2009) The crystal structure of the outer membrane

lipoprotein YbhC from Escherichia coli sheds new light on the phylogeny of carbohydrate esterase family 8. Proteins 76: 1029‐1036

3. Mark P, Baumann MJ, Eklöf JM, Gullfot F, Michel G, Kallas AM, Teeri TT, Brumer H, Czjzek M (2009) 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: Struct. Funct. Bioinf. 75: 820‐836

4. Nordgren N, Eklöf JM, Zhou Q, Brumer H, Rutland MW (2008) Top‐down grafting of

xyloglucan to gold monitored by QCM‐D and AFM: Enzymatic activity and interactions with cellulose. Biomacromolecules 9: 942‐948

*Authors contributed equally to the work


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Content

Introduction ............................................................................................................................................. 1 Plants & Man ....................................................................................................................................... 2 The Origin & Early Evolution of Plants ................................................................................................ 2 The Embryophytes ............................................................................................................................... 3 The Plant Cell Wall ............................................................................................................................... 5 The primary cell wall ....................................................................................................................... 5 The secondary cell wall ................................................................................................................... 6 Cell wall growth ............................................................................................................................... 7

Carbohydrate Active enZymes (CAZymes) .......................................................................................... 9 Reaction mechanism of glycoside hydrolases ............................................................................... 10

Xyloglucan ......................................................................................................................................... 13 Xyloglucan structure ...................................................................................................................... 14 The biosynthesis of xyloglucan ...................................................................................................... 15 The function of xyloglucan ............................................................................................................ 17 The evolution of xyloglucan .......................................................................................................... 17

Xyloglucan Degradation .................................................................................................................... 18 The Xyloglucan endo‐transglycosylase/hydrolase (XTH) Gene Products .......................................... 20 Molecular phylogeny of XTH gene products ................................................................................. 21 The structural basis of XET versus XEH activity ............................................................................. 24 How XETs and XEHs recognise their natural substrates ................................................................ 29 Alternate substrates, and activities, for XTH gene products ......................................................... 31 Other transglycosylases in plants .................................................................................................. 33 A future for “old‐fashioned” enzymology in these modern times of high‐volume functional genomics? ...................................................................................................................................... 34

Results ................................................................................................................................................... 37 Microbial Xyloglucan Degrading Enzymes ......................................................................................... 38 Aim of investigation ...................................................................................................................... 38 Structural convergent evolution of xyloglucanases ...................................................................... 38 Probing substrate specificity in xyloglucanases ............................................................................ 39 What makes a xyloglucanase? ...................................................................................................... 39

Plant GH16 Enzymes .......................................................................................................................... 41 Aim of study .................................................................................................................................. 41 Activity of plant GH16 enzymes .................................................................................................... 41 In vivo localisation of XEH activity in Group III‐A knockouts in Arabidopsis ................................. 43 Structure function relationships in XTH gene products ................................................................ 44

Activity of plant endo β‐1,4 glucanases, EG16s ............................................................................. 45 The phylogeny of plant GH16 enzymes ......................................................................................... 45 Evolution of XTH gene products and the shift in substrate specificity ......................................... 47

Conclusions ............................................................................................................................................ 50 Acknowledgements ............................................................................................................................... 51 References ............................................................................................................................................. 52 Appendix ................................................................................................................................................ 63

Jens Eklöf

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Introduction


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Plants & Man

One life form has probably changed the world more than any other. Since the first plants left their

aquatic habitats for land, almost 500 million years ago, they completely changed Earth, conquering

and dominating just about all ecological niches. Plants have also played a major role in the human

evolution. The domestication of food crops, such as fig and barley, was an absolute requirement for

the development of the human, sedentary agriculture society that first appeared in the fertile crest

around 10000 BCE (Salamini et al., 2002). This sedentary culture led, with time, to the development

of the written language and the world we know today.

Still today, plants play important roles in our lives, as food, materials and as a source of energy. Of

Sweden’s export value, about 12% comes from wood related products and adding up to about 3% of

Sweden’s GDP (http://www.skogsindustrierna.org).

Our dependence of plants is likely to increase. One of the major challenges of the future lies in

securing food for a growing world population and at the same time exchange fossil fuels for

sustainable alternatives, of which plant biomass is likely to be one. Solving these truly

multidisciplinary issues will require changes on many levels. To better understand how plants

function is one of these issues, and might lead to better crops, new materials or improved efficiency

in biomass conversion for biofuels.

The Origin & Early Evolution of Plants

Niklas defined plants as “photosynthetic eukaryotes”, including very distantly related organisms

under the umbrella of plants (Niklas, 2000). A key developmental step in plant evolution leading to

the first green alga was the endosymbiosis of a photosynthetic cyanobacterium by a unicellular

eukaryote which lead to the first photosynthetic eukaryote as hinted by Schimper in the 19th century

(Schimper, 1883). With time, the internalised cyanobacterium lost many genes either completely or

via transfer to the nucleus eventually becoming dependent on its host for survival. These host‐

dependent cyanobacteria are the chloroplast, or the plastid in which photosynthesis takes place in all

plants. One or more of these endosymbiotic events lead to the emergence of three different

lineages of photosynthetic organisms namely the glaucophytes, the red algae and the green algae

(Keeling, 2004). The green algae (Viridiplantae) in turn divided into the marine living chlorophytes

and the freshwater dwelling streptophytes (charophytes) (Fig. 1a). The early adaptation of

streptophytes to fresh water habitats was instrumental in their eventual land colonisation. The

transition from freshwater to land habitats was probably done through a series of steps where the

adaptation of living in pools with periodic desiccation ultimately lead to plants that could live in

habitats without constant water supply (Becker and Marin, 2009). Marine dwelling green algae on

Jens Eklöf

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the other hand had a longer step transitioning from a saline marine environment to the freshwater

rain‐based land (Becker and Marin, 2009).

Figure 1. The tree of plants. In a, all plants originating from an ancestral eukaryotic endosymbiotic event of a cyanobacterium believed to have happened over 1500 million years ago (MYA) (Palmer et al., 2004) are shown. The tree has an emphasis on Viridiplantae and streptophytes. In b, a tree showing the relationship within the Embryophyta is shown with a charophycean green alga as an ancestral species.

The question of which order within the Streptophyta that finally colonised land about 480 MYA

(Kenrick and Crane, 1997; Qiu and Palmer, 1999) has and continues to be an enigma and a constant

reason for debate within the scientific community. The closest living relatives of land plants have

historically been placed either in the order of Charales or Coleochaetales using either morphological

or sequence data (discussed in Finet et al. 2010). A recent extensive analysis of 77 nuclear genes

suggested that the last common ancestor of the embryophytes was from the order of Coleochatales

supported both by high Bayesian posterior probabilities and boot strap values using Bayesian‐ and

maximum‐likelihood methods respectively (Finet et al., 2010). However, the issue of the origin of

embryophytes will not be settled until there are representative genomic sequences or at least more

extensive EST libraries of streptophytes.

The Embryophytes

The Charales and the Coleochaetales have advanced body plans similar in many respects to the land

living embryophytes. Features such as plasmodesmata, phragmoplasts and apical meristems were

developed within the streptophytes before land colonisation (Graham et al., 2000). One of the major

differences between the charophycean green algae and embryophytes are their life cycles. The

Charales and Coleochaetales are haplobiontic where only the gametophyte (1n) has a multicellular

phase. Land plants are on the other hand diplobiontic with multicellular gametophytes and

sporophytes (2n) alike. The oldest, still living, lineages of embryophytes, the bryophytes (liverworts,


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mosses and hornworts) appeared around 480 MYA. The bryophytes still have the gametophyte as

the main vegetative growth phase with the sporophyte being dependent on the gametophyte for

nutrition.

With time new groups of embryophytes (Fig. 1b) emerged with new inventions that made them more

competitive for nutrients and light. Below, highlights of important events in embryophyte evolution

are listed:

• Lycophytes (club mosses): the first paleobotanical evidence of lycophytes, Cooksonia and

Rhynia is from 415 MYA. They had water conductive tissue and the sporophyte generation

dominated over the gametophyte generation (Gerrienne et al., 2006). The water conductive

tissues allowed plants to grow taller, in this manner giving them an advantage in the

competition for sunlight.

• Monoliophytes (ferns) evolved around 380 MYA (Pryer et al., 2004; Schneider et al., 2004)

and have true leaves with a branched vein structure called megaphylls (fronds) that allow the

growth of larger leaves.

• Gymnosperms (conifers, cycads, Gnetales and Ginko) developed the first seeds ca. 360 MYA

(Gerrienne et al., 2004) and dispersion of pollen is mostly air‐ and not water‐dependent as

opposed to more basal plants.

• Angiosperms appeared ca. 140 MYA (Hughes, 1994; Moore et al., 2007; Bell et al., 2010) and

developed carpels, flowers and protected seeds. The endosperms of angiosperms are triploid

(3n) as compared to gymnosperms (haploid, 1n) (Stuessy, 2004).

Once the angiosperms appeared on Earth they soon spread and dominated the landscape already

around 90 MYA. The number of angiosperm species virtually exploded with more than 250 000

species known today being roughly ten times higher than the total number of species for all other

groups of land plants (Palmer et al., 2004; De Bodt et al., 2005; Crepet and Niklas, 2009). The

angiosperms are divided into several groups, but the two main lineages are the monocotyledons,

which contain the grasses and dicotyledons, that comprise almost 200 000 species of the worlds

flowers, shrubs and trees.

Jens Eklöf

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The Plant Cell Wall

Cell walls have developed in almost every major clade of unicellular organisms (Niklas, 2004). The

polymers that build these walls differ between different lineages but they all share the common

function of protecting the cell. All plants cells have a polysaccharide‐based wall, most likely because

carbon is the least limiting resource for plants while cell walls of other organisms can be protein‐

based as in the S‐layer of Archea (Sara and Sleytr, 2000) or a copolymer of polysaccharides and

peptides as in the peptidoglycan found in most bacteria.

The primary cell wall

The primary cell wall is thin, usually between 0.1 and 10 µm thick and determines the cell shape and

size. It must be mechanically strong enough to support the protoplast and protect it from turgor

pressure, whilst at the same time being plastic enough to allow the cell to grow without bursting

(Cosgrove, 2005; Jarvis, 2011). In addition, the primary cell wall is also the first line of defence

against pathogens and a source of signalling molecules (Lagaert et al., 2009).

The Viridiplantae (green algae and land plants), have a complex polysaccharide‐based primary cell

wall surrounding the protoplast. The wall can be described as a hydrogel containing up to 90% water

(Redgwell et al., 1997; Redgwell et al., 2008; Rondeau‐Mouro et al., 2008) where crystalline cellulose

microfibrils construct a fortifying network coated and cross‐linked by hemicelluloses with pectins as

matrix polysaccharides (Fig. 2a) (Carpita and Gibeaut, 1993; Carpita and McCann, 2000). Primary cell

walls also contain structural glycoproteins e.g. arabinogalactan proteins (AGPs) and hydroxyproline‐

rich proteins (HPRPs) alongside catalytically active or non‐active proteins for cell wall remodelling or

degradation, e.g. expansins, xyloglucan endo‐transglycosylases/hydrolases (XTHs) gene products and

pectin methylesterases (PMEs) to mention a few (Micheli, 2001; Cosgrove, 2005).

Structurally, primary cell walls of the economically important spermatophytes have traditionally been

divided into two types, Type I and Type II (Carpita and Gibeaut, 1993). Recent progress in the cell

wall composition of more basal embryophytes reviewed in Popper and Tuohy (2010) indicates that

Type I‐like cell walls are found in most embryophytes while Type II cell walls are mainly found in

members of the grassy monocots in the Poales and some closely related species. In Type I walls,

xyloglucans are the main hemicelluloses and they also have a large proportion of pectic

polysaccharides. While similar in structure, Type II walls have increased levels of

glucuronoarabinoxylan (GAX) and mixed‐linked glucan (MLG) but reduced levels of xyloglucans and

pectins (Carpita and Gibeaut, 1993; Vogel, 2008). Both GAX and MLG are classified as hemicelluloses

and can hydrogen‐bond to cellulose. In maize‐coleoptiles, MLG coats cellulose microfibrils while


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different domains of GAX either bind cellulose or behave like a matrix polymer, similar to the

predicted function of pectin (Carpita et al., 2001). It should be noted that primary cell walls display

large variations in temporal, tissue and species dependent manners (Knox, 2008).

Figure 2. Models of the plant cell walls. a, the primary cell wall where cellulose microfibrils coated by hemicelluloses form a strong yet pliable cell wall. b, the secondary cell wall with the thin primary cell wall containing randomly oriented cellulose microfibrils on the outside (P). The secondary cell wall has a layered structure (S1‐S3) with layers having different microfibrillar angles.

Plants have a special layer gluing adjacent cells together called the middle lamella. This pectin‐rich

middle lamella cross‐links cells by a pectinaceous network where Ca2+ brigdes between carboxylates

of homogalacturonan (HG) mediate the cross‐links (Vincken et al., 2003). The pectic polysaccharide

rhamnogalacturonan II (RGII) can also be cross‐linked through boron bridges between two apiose

residues (Vincken et al., 2003) but is mainly found in the primary cell wall. In certain tracheophyte

tissues, a third cell wall layer called the secondary cell wall is present.

The secondary cell wall

Early embryophytes and bryophytes were not limited to a life close to the ground solely by the

dependence on water diffusion. Physical constraints probably also prevented them from growing

taller. It was not until the evolution of the secondary cell wall and xylem in tracheophytes that plants

could compete for sunlight by rising higher above their substrate.

Many important products such as paper, timber and wood are mainly secondary cell walls from

gymnosperm‐ and angiosperm trees called soft and hardwood respectively. Both hardwood and

softwood contains 40‐50 % cellulose (dry weight), up to 35 % hemicelluloses and up to 25 % lignin

with small amounts of pectins, proteins and extractives. The main hemicelluloses in secondary cell

walls of softwoods are mannans while the main hemicelluloses in hardwoods are xylans. The

deposition of the secondary cell wall takes place between the primary cell wall and the plasma

Jens Eklöf

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membrane and is usually divided into three layers called S1, S2 and S3 with different cellulose

microfibrillar angles (Fig. 2b).

The secondary cell wall is usually lignified. Lignin is a radical‐formed hydrophobic polymer made

from three main monomers of p‐hydroxycinnamyl alchohols: p‐coumaryl‐, H; coniferyl‐, G; sinapyl‐

alchohols, (S) (Vanholme et al., 2010). The capacity to synthesise S‐monolignols was previously

believed to be restricted to angiosperms (Ralph et al., 2004) but recent findings confirm their

presence in more basal species, albeit from different pathways (reviewed in (Weng and Chapple,

2010)).

Cell wall growth

Cell wall growth is governed by changes in turgor pressure and cell wall‐loosening agents. The turgor

pressure is maintained by the uptake of water into the vacuole and is around 0.3‐0.9 MPa (3‐9 times

the atmospheric pressure; (Cosgrove, 1993)). Because of the high pressure, the cell wall extension

needs to be tightly regulated to prevent cells from bursting. The process of cell wall growth is called

polymer creep and involves slow sliding of individual microfibrils and their associated

polysaccharides within the cell wall creating an increased surface area (Marga et al., 2005).

Simultaneously, new cell wall material is synthesised and secreted to maintain the cell wall strength.

The cell enlargement can be dramatic. Xylem vessels for example can increase their volume more

than 30 000 times compared to their meristematic initials (Cosgrove, 2005).

The primary cell wall can be seen as an external cell organ where the cell has good control over

apoplastic pH and the proteins and materials it sends into it. This allows the cell to control both the

direction and rate of growth. Firstly, the direction of growth is controlled by the orientation of

cellulose microfibrils (anisotropic growth). Secondly, one of the few protein groups with proven cell

wall loosening ability, the expansins, can be secreted and targeted to certain parts of the cell wall

(Cosgrove, 2000). The α‐expansins have increased cell stress‐relaxing abilities at lower pHs than

normally found in cell walls and are probably the main contributor in acid growth. The mechanism

by which expansins loosen the cell wall is not known but it has been speculated that they break

hydrogen bonds between cellulose microfibrils and other cell wall polysaccharides thereby

facilitating polymer creep (Cosgrove, 2000). Thirdly, a range of other enzymes are secreted into the

cell wall during growth. Endo‐ or exo‐acting glycoside hydrolases can alter the properties of

polysaccharides and pectin methylesterases (PMEs) can fortify cell walls by calcium mediated pectin

bridges (egg boxes). PMEs have fairly high pH optima and are activated first in later stages of cell

wall extension when the pH rises (Micheli, 2001). The xyloglucan endo‐transglycosylase /hydrolase

(XTH) gene products are an example of glycoside hydrolases/transglycosylases involved in cell wall


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loosening and strengthening. These proteins exhibit either xyloglucan endo‐transglycosylase (XET, EC

2.4.1.207) or xyloglucan endo‐hydrolase (XEH, EC 3.2.1.151) activity (Eklöf and Brumer, 2010).

Experiments on pea stems and suspension cultures of tobacco cells showed that externally added

high Mr xyloglucan suppressed cell elongation, probably due to the incorporation of xyloglucan into

the cell wall matrix by XETs. Upon the addition of xylogluco‐oligosaccharides (XGOs) to the same

system, XETs caused a depolymerisation of the native xyloglucan leading to accelerated elongation

(Takeda et al., 2002; Kaida et al., 2010). This demonstrates XETs ability to cause either strengthening

or loosening of the cell wall, depending on available substrates. Albeit, experimental proof that

plants use XGOs as growth modulators has not yet been presented. The cell wall‐strengthening

effects of XETs has however been shown in Arabidopsis (Maris et al., 2009). Finally, non‐enzymatic

cleavage of polysaccharides might also lead to cell wall loosening. Hydrogen peroxide or superoxide

can be decomposed catalytically into the potent radical •OH by metal ions such as Cu+ or Fe2+ (Fry et

al., 2002; Swanson and Gilroy, 2010) through the Fenton reaction.

In some tissue, another type of cell wall extension takes place. In trichoblasts (certain root epidermal

cells) and pollen tubes, a local weakening of the cell wall caused by acidification leads to a

phenomenon called tip growth. Tip growth involves a tip focused Ca2+ gradient, polarised actin

polymers, and tip‐directed vesicle trafficking (reviewed by (Cole and Fowler, 2006)).

Jens Eklöf

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Carbohydrate Active enZymes (CAZymes)

The diversity in carbohydrates far exceeds any other biopolymer. In nature, over 30 different sugars

(Marth, 2008) have been found which can be further modified by non‐carbohydrates, such as

sulphate, phosphate and acyl esters. Only assuming unmodified carbohydrates, the number of

theoretical hexasaccharide isomers from D‐hexoses reach a staggering 1012. Therefore the enzymes

responsible for their biosynthesis, modification and degradation face a daunting task. The work of

classifying carbohydrate active enzymes began around 1990 by Bernard Henrissat, comparing

different glycoside hydrolases (Henrissat et al., 1989; Henrissat, 1991). The resulting database, CAZy

is an excellent example of a successful bioinformatic project and today it is an instrumental tool for

all glyco‐scientists, with approximately 3000 downloaded pages daily, showing the key role sequence

classification has in carbohydrate research (Davies and Sinnott, 2008).

Figure 3. The year by year growth of GH ORFs in the CAZy database. Figure from Davies &

Sinnott, 2008.

Bernard Henrissat began the CAZy project by ordering 291 sequences into 35 glycoside hydrolase

families, GH1‐GH35, using the unusual method, hydrophobic cluster analysis (HCA; (Gaboriaud et al.,

1987)). From the original classification of 35 GH families, CAZy has continuously grown and

contained almost 40 000 ORFs of GHs ordered into 112 GH families in 2008 (Fig. 3; (Davies and

Sinnott, 2008)). The database is constantly being updated and now contains 122 GH families

(January 2011). Of these 122 GH families, GH21, 40, 41, 60 and 69 (Cottrell et al., 2005; Smith et al.,

2005) have been discontinued due to lack of hydrolytic activity on glycosidic bonds while others like


10

GH61 is still maintained even though evidence is mounting that they are not glycoside hydrolases

(Harris et al., 2010; Vaaje‐Kolstad et al., 2010) but involved in biomass conversion by other means.

Following the original initiative to classify GHs, other types of carbohydrate‐acting proteins have

been added to CAZy. The new groups are glycosyltransferases (GTs), polysaccharide lyases (PLs),

carbohydrate esterases (CEs) and carbohydrate binding modules (CBMs).

The power of the CAZy database lies in that the classification into GH families contains more

information than what can be derived from an EC number (NC‐IUBMB,

http://www.chem.qmul.ac.uk/iubmb/enzyme/). The EC numbers 3.2.1.x account for all GH activities

as the first three numbers indicate that an enzyme hydrolyses glycosidic or thio‐glycosidic linkages.

The shortcomings of EC numbers are that they only account for one reaction and seldom reveal

anything about the mechanism of an enzyme. For GHs, who often have broad substrate specificities,

a single EC number is not sufficient. Instead CAZy classifies enzymes according to their amino acid

sequence and fold. This classification enables the user to infer protein fold, often reaction

mechanism and catalytic amino‐acids from other family members as well as getting hints to enzyme

activity.

Reaction mechanism of glycoside hydrolases

The glycosidic bond, especially the β‐1,4 bond between two glucose residues, is the strongest bond

found in biopolymers with a calculated half‐life in excess of 4 million years. Glycoside hydrolases

face a daunting task trying to hydrolyse these bonds but have in certain cases managed to accelerate

the reaction by an impressive 1017‐fold (Wolfenden et al., 1998).

A comprehensive understanding of how enzymes accelerate reactions has not yet been reached and

many factors influence enzyme rates. In the 1930s, JBS Haldane built on the lock and key simile, but

said that the key doesn’t perfectly match the lock and exercises a certain strain on it (Haldane, 1930).

In the 40’s, the Nobel laureate Linus Pauling stated that enzymes must bind their substrate in a

constrained conformation which corresponds to an activated complex (Pauling, 1946). Still today

Linus Pauling’s idea about transition state stabilisation is generally accepted as a major contributor to

enzyme catalysis but there are also other factors. Enzymes distort reactants either physically or

electronically around the site of catalysis and also bring reactants into contact distance of each other

and catalytic residues (Menger, 2005). However, substrate interactions tens of Ångström away from

the catalytic residues can drastically alter the rate of catalysis.

Concerning glycoside hydrolases, Koshland first laid the basis of how GHs accomplish hydrolysis

already in 1953 (Koshland, 1953). Since then much has been published on the mechanisms of

Jens Eklöf

11

different glycoside hydrolases but most of the early theories still hold true (Vocadlo and Davies,

2008). Even though several exceptions are known (Vuong and Wilson, 2010), essentially all GH

families use one of two reaction mechanisms, either the inverting mechanism or the retaining

mechanism (Rye and Withers, 2000) reflecting the configuration of the anomeric oxygen of the

product compared to the substrate.

The inverting mechanism is a one step mechanism leading to overall inversion of the configuration of

the anomeric carbon. It uses a general base to activate a water molecule by extracting a proton and

an acid to facilitate the departure of the leaving group by protonation. The transition state is

proposed to be oxocarbenium‐ion‐like (Figure 4a; (Vocadlo et al., 2001)). The catalytic residues are

usually aspartates or glutamates and even though the distance between them have been shown to

vary considerably (Zechel and Withers, 2000) a general rule of thumb is that they are ~10 Å apart.


12

Figure 4. General mechanisms of glycoside hydrolases. a, the inverting mechanism; b, the retaining mechanism;. c, the substrate‐assisted mechanism; d, the syn or anti protonation of the leaving group.

Jens Eklöf

13

The other common mechanism used by glycoside hydrolases is the retaining mechanism (Fig. 4b). It

is a two step mechanism involving a nucleophile and an acid‐base functionality. Similar to the

inverting glycoside hydrolases, the catalytic residues are usually aspartates or glutamates but with

some variation as in sialidases and trans‐sialidases of GH33 and GH34, where a tyrosine acts as the

nucleophile (Watts et al., 2003). In GH18, 20, 25, 56, 84, 85 and 103, substrate‐assisted catalysis is

utilised by employing an acetoamide group from the substrate itself as the nucleophile (Fig. 4c;

(Macauley et al., 2005)). In the retaining mechanism the first step is an attack by the nucleophile on

the anomeric carbon of a sugar ring resulting in a glycosyl‐enzyme intermediate. The departure of

the leaving group is assisted by protonation from the acid‐base in a syn or anti fashion (Fig. 4d). In

the second step the acid‐base activates a water molecule that subsequently attacks the anomeric

carbon releasing the product with retention of the stereochemistry at the anomeric carbon and

restoring the active site. For a more in‐depth discussion on the mechanistic properties of glycoside

hydrolases the reader is referred to a recent review by Vocadlo and Davies, 2008.

Xyloglucan

Xyloglucan is the main hemicellulose in the primary cell wall of most land plants where it coats and

tethers adjacent cellulose microfibrils (Nishitani, 1998). In the primary cell walls of dicots it can

represent as much as 20‐25 % of the dry mass (Ebringerova, 2006). Xyloglucan has also been found

in tension wood fibers in poplar where it seems to be important for the gravitropic response

(Nishikubo et al., 2007; Baba et al., 2009). In certain plants, including the ornamental plant

nasturtium (Tropaeolum majus) and the tropical tree tamarind (Tamarindus indica) xyloglucan has

been recruited as a seed storage polysaccharide, supplying the growing embryo with both hexoses

and pentoses (Kooiman, 1960; Reid, 1985; Buckeridge et al., 2000; Buckeridge, 2010).


14

Figure 5. Common structural motifs of xyloglucan and xylogluco‐oligosaccharides (XGOs). Both a and b depict the same XLFG cellotetraose‐based XGO. Each glucosyl in the glucan backbone is denoted by the sidechains it carries. A glucosyl with an α‐1,6 xylosyl substitution is called X. If the xylosyl is further substituted by a β‐1,2 galactosyl unit the branch is called L. In the primary cell walls of angiosperms the F branch is common and is an α‐1,2‐L‐fucosyl substitution on the galactosyl. Unsubstituted glucosyl units are found at every fourth glucosyl unit in the backbone and are denoted G (XGO nomenclature from Fry et al., 1993)

Xyloglucan structure

Structurally, xyloglucans are branched glucans constituted by a range of different oligosaccharide‐

monomers with a common cellotetraose (β‐1,4‐linked glucan) backbone (Fig. 5). To account for the

different branching patterns of the xylogluco‐oligosaccharides (XGOs), Fry et al., (1993) came up with

a unifying nomenclature, naming sidechains of individual glucosyl units. The first two or three

glucosyl units, counting from the non‐reducing end are substituted by α‐1,6 xylosyl residues and

denoted X with the reducing end glucosyl unit unsubstituted (G). These XXGG or XXXG‐motifs form

the basis for all other xyloglucan monomers (Vincken et al., 1997). The middle two X motifs of XXXG

can then be further substituted by β‐1,4 galactosyl residues and are denoted L. Another common

structural motif in the primary cell wall xyloglucan of higher plants contains an α‐1,2‐L‐fucosyl

residue on the galactosyl closest to the reducing end (F; Fig. 5). The most common type of monomers

in land plants are thus XXXG, XXLG, XLXG, XXFG, XLLG and XLFG (Hoffman et al., 2005; Peña et al.,

2008; Hsieh and Harris, 2009). In the oldest lineages of the land plants, i.e. liverworts and mosses,

fucosylated xyloglucan is not found. Instead, a more pectin‐like anionic xyloglucan with galacturonic

acid residues has been found (Peña et al., 2008). Other deviations from the standard xyloglucan

monomers are found in the lycophytes where arabinose can replace galactose (Peña et al., 2008), in

the solanaeous plants where arabinose replaces fucose (York et al., 1996) and in seed storage

xyloglucans that lack fucose (Faik et al., 2000).

Jens Eklöf

15

The branching pattern of xyloglucans influences the physico‐chemical properties of the

polysaccharides. A homo‐xyloglucan made from XXXG motifs had low water solubility while XLLG‐

based homo‐xyloglucan and native xyloglucan with mixed monomer composition has much higher

water solubility (unpublished results, and (Gullfot et al., 2009). While xyloglucan monomers are

believed to be randomly distributed in the polymer there is a least one case where xyloglucan has a

domain‐like structure (Tine et al., 2003; Tine et al., 2006). Xyloglucan may also be linked covalently

to other polysaccharides. A study using Arabidopsis cell cultures has indicated that as much as 50 %

of the xyloglucan in these walls is covalently linked intra‐cellularly to the pectic polysaccharide

rhamnogalacturonan I (Popper and Fry, 2008).

The biosynthesis of xyloglucan

In plants, the hemicelluloses and pectins are synthesised in the Golgi apparatus and exported to the

apoplast by exocytosis (Scheller and Ulvskov, 2010) and only cellulose and callose are made in the

plasma membrane (Somerville, 2006; Guerriero et al., 2010; Carpita, 2011) (Fig. 6). The rather poor

understanding of polysaccharide biosynthesis by membrane‐bound glycosyltransferases is, at least in

part, due to the fact that membrane‐bound proteins are more difficult to study compared to soluble

proteins and might also require the assembly of complexes or post‐translational modifications for

activity. The enzymes involved in xyloglucan biosynthesis are fairly well mapped compared to other

plant cell wall polysaccharides. The backbone of xyloglucan is synthesised by enzymes belonging to

the cellulose synthase like C (CslC) group. Arabidopsis has five CslC genes of which only CSLC4 has

been directly implicated in synthesising the β‐1,4 glucan backbone of xyloglucan. The α‐xylosylation

branching is made by two α‐1,6 glycosyltransferases from family 34 (GT34; (Cavalier et al., 2008)).

Three more GT34s could be involved in xyloglucan xylosylation as a third GT34, XXT5 has also been

suggested to be involved in xyloglucan α‐xylosylation in Arabidopsis (Zabotina et al., 2009). Other

enzymes involved in xyloglucan assembly were found when monitoring fucose epitopes in plant cell

wall mutants. Using this methodology, an α‐fucosyl transferase and a β‐galactosyl transferase

specific to the third X in the XXXG‐motif were found (Madson et al., 2003; Li et al., 2004). The other

enzymes involved in the biosynthesis of xyloglucan are still unidentified.


16

Figure 6. The biosynthesis of polysaccharides and the vesicular trafficking to the primary cell wall. Hemicelluloses and pectins are made in the Golgi apparatus and secreted into the cell wall by exocytosis while the biosynthetic machinery for cellulose and callose are located in the plasma membrane. Figure from Cosgrove (2005).

Localisation studies have recently shown the specific Golgi localisation of known glycosyltransferases

involved in xyloglucan biosynthesis in tobacco BY‐2 cells. The first branching enzymes, the α‐

xylosyltransferases are mainly found in the cis‐ and medial Golgi regions while the β‐galactosyl‐ and

α‐fucosyltransferases are found from the medial to the trans Golgi compartments (Chevalier et al.,

2010).

Despite this knowledge, the mode of assembly of xyloglucan still remains an enigma. Is it assembled

from XGOs to xyloglucan or are the branch‐building enzymes working on longer glucan chains? To

me personally, it seems more plausible that xyloglucan is assembled from monomers, perhaps lipid

anchored, by a hitherto unknown enzyme. If so, a link between the xyloglucan and the β‐1,3;1,4

glucan (MLG) biosynthesis of the Poaceae can be postulated. Arabidopsis walls do not contain any

MLG but when Poaceae specific CslH or CslF genes are expressed in Arabidopsis, MLG can be

detected in the cell wall (Burton et al., 2006; Doblin et al., 2009). This can either be due to a dual

function of these genes making both β‐1,3 and β‐1,4 glucosidic bonds or more likely (in my mind),

these two gene families can use the existing biosynthetic machinery of the CslCs (responsible for

biosynthesis of the xyloglucan backbone) and connect shorter β‐1,4 glucan oligos by β‐1,3 bonds.

They could do so by being located in the cis Golgi, before α‐xylosyltransferases or by making

complexes with the CslCs. A compelling circumstantial evidence of this hypothesis is the similarity in

length of the repeating units of XGOs and MLG.

Jens Eklöf

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The function of xyloglucan

Despite xyloglucan’s ascribed role as an integral part of contemporary cell wall models, cardinal work

by Cavalier et al. (2008) showed that Arabidopsis plants lacking two α‐xylosyltransferase genes had

no detectable xyloglucan in their cell wall whilst only displaying a mild dwarf phenotype with

aberrant root hairs (Cavalier et al., 2008). Recent analyses of this xxt1/xxt2 double mutant indicated

that the cell walls are weakened and that xyloglucan is needed for the spatial distribution of cellulose

microfibrils and to prevent the formation of microfibrilar aggregates (Anderson et al., 2010). This

shows that xyloglucan is not an absolute requirement in land plants (under laboratory conditions)

and that there is at least a partial functional redundancy of different plant cell wall polysaccharides.

XGOs have been shown to negatively influence growth indirectly by antagonising the effect of auxin

(Lorences et al., 1990; Fry et al., 1993) but they can also increase cell elongation probably by XET

mediated xyloglucan depolymerisation (Takeda et al., 2002; Kaida et al., 2010).

The evolution of xyloglucan

Despite not being essential to Arabidopsis, xyloglucan could have been a key component in the

transition from water to land (Popper and Fry, 2003). Previously not considered present in the

charophycean algae (Popper and Fry, 2003) recent antibody‐based microarray experiments and

methylation analyses by GC‐MS suggest small amounts of xyloglucan in specific cells in both Chara

corallina and Spirogyra sp. (Moller et al., 2007; Ikegaya et al., 2008; Domozych et al., 2009;

Domozych et al., 2010). In support of the presence of a xyloglucan‐like polysaccharide in

charophycean green algae is the finding of a CslC gene in Chara globularis inferred to be involved in

making the backbone glucan of xyloglucan (Del Bem and Vincentz, 2010). Caution is warranted not

to over‐analyse antibody detection data of polysaccharides as antibodies can detect epitopes

potentially present in more than one polysaccharide and polysaccharides can also be masked by

other polysaccharides in the cell wall (Marcus et al., 2008).

If charophycean green algae indeed have small amounts of xyloglucan, I believe the recruitment of

xyloglucan to all primary cell walls and in larger quantities was seminal to early land colonising plants

rather than acquiring the ability to synthesise xyloglucan. This would be analogous to the pectic

polysaccharide RGII found in all land plants but only in very low levels in bryophytes compared to

tracheophytes. Higher levels of RGII together with the invention of lignin are believed to be

important events in the tracheophyte evolution (Matsunaga et al., 2004; Popper and Tuohy, 2010).


18

Xyloglucan Degradation

With the realisation that peak oil is already upon us or at least around the corner, together with

national security decrees of energy independence have lead to a surge of interest in biomass

conversion. Even though emphasis has been on degradation of recalcitrant cellulose, lignocellulose

materials and starch, research has also been conducted on hemicellulose degradation (Gilbert et al.,

2008; Vlasenko et al., 2010). Nature’s large arsenal of xyloglucan degrading enzymes, from a range

of different GH families are now being studied in detail and papers within this thesis have made real

contributions to the understanding of some of them.

The initial attack in xyloglucan degradation is made by endo‐acting xyloglucanases residing in the

retaining families GH5, 7, 12, 16 and 44 as well as in the inverting GH74 that depolymerise high

molecular mass xyloglucan into XGOs (Juhász et al., 2005; Gilbert et al., 2008). While most of these

xyloglucanases act by cleaving xyloglucan after an unsubstituted glucosyl unit, G (Fig. 7a; EC

3.2.1.151, www.chem.qmul.ac.uk/iubmb/enzyme) notable exception have been found in GH44 and

GH74. A GH44 enzyme (Paper I) has been shown to preferably cleave xyloglucan with an X motif in

the ‐1 subsite and a G in the +1 subsite while the GH74 enzyme uses this mode of cleavage to a lesser

extent (Yaoi et al., 2005). Other unusual xyloglucan cleavage patterns have been observed for

various GH74 enzymes. Some prefer cleavage between two X motifs (Desmet et al., 2007) whilst two

GH74 enzymes have been shown to be reducing end‐specific cellobiohydrolases (EC 3.2.1.150; (Yaoi

and Mitsuishi, 2002; Bauer et al., 2005)) releasing XG, LG or FG from xyloglucan. The exo‐acting

activity is believed to be due to a loop insertion closing off the positive subsites (Yaoi et al., 2007). A

similar mechanism for the exo‐activity of a ruminal GH5 enzyme was recently proposed (Wong et al.,

2010).

Sometimes, the mode of action on xyloglucan is less clear. A GH74 enzyme from Chrysosporium

lucknowense has shown mixed endo and exo‐activity, initially attacking a xyloglucan chain in an endo

fashion but at a later stage mainly working in an exo fashion, producing XXXG‐based XGOs

(unpublished data Eklöf; (Grishutin et al., 2004)). A mixed type of degradation pattern has also been

reported for a GH7 cellobiohydrolase previously believed to be acting exclusively on the reducing end

of cellulose chains. A more in‐depth analysis of this enzyme has however shown that it can also

initiate new cuts in an endo fashion (Kurašjin and Väljamäe, 2011).

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Figure 7. The degradation of high Mw xyloglucan to monosaccharides. The first step in xyloglucan degradation is to depolymerise xyloglucan into XGOs by endo‐acting xyloglucanases (a). In b, some of the exo‐acting side chain‐trimming enzymes are shown together with the XGO nomenclature for the respective xyloglucan units (Fry et al., 1993). In c, the degradation of XXXG into monosaccharides by the sequential action of α‐xylosidases and β‐glucosidases as observed in Arabidopsis and bacteria (Iglesias et al., 2006; Larsbrink et al., 2011).

After conversion of high molecular mass xyloglucan by endo‐acting xyloglucanases into XGOs, exo‐

acting enzymes debranch the XGOs (Fig. 7). In Arabidopsis the apoplastic degradation of XGOs has

been studied and there it seems like side chains are stripped off by α‐1,2‐L‐fucosidases (GH95, EC

3.2.1.63) and β‐galactosidases (GH1, 2, 35, 42 and 43; EC 3.2.1.23) to create xylosylated XGOs

(Iglesias et al., 2006). These are then hydrolysed into monosaccharides starting from the non‐

reducing end by the concerted and sequential actions of α‐xylosidases (GH31; EC 3.2.1.X) and β‐

glucosidases (GH1 and GH3; EC 3.2.1.21) (Buckeridge et al., 2000). In other species, other de‐

branching, exo‐acting enzymes are most likely present acting on XGOs, for example α‐L‐

arabinofuranosidases (GH3, 10, 43, 51, 54 and 62; EC 3.2.1.55).


20

The Xyloglucan endotransglycosylase/hydrolase (XTH) Gene Products1

In contrast to microbial endo‐acting xyloglucanases that are found in numerous GH families (Gilbert

et al., 2008; Vlasenko et al., 2010), plants enzymes responsible for the cleavage and/or

rearrangement of the xyloglucan backbone in muro have so far only been identified in GH16.

Although this limited distribution certainly belies their importance. The xyloglucan endo‐

transglycosylase/hydrolase (XTH)2 genes encode proteins that can potentially have two distinct

catalytic activities, with radically different effects on xyloglucan: Xyloglucan endo‐transglycosylase

(XET) activity (formally, xyloglucan:xyloglucosyl transferase; EC 2.4.1.207) results in the (non‐

hydrolytic) cleavage and ligation of xyloglucan chains, whereas xyloglucan endo‐hydrolase (XEH)

activity (formally, xyloglucan‐specific endo‐β‐1,4‐glucanase; EC 3.2.151) yields irreversible chain

shortening. Since their initial discovery in the early 1990’s, XETs have figured prominently in plant

cell wall models, due to the believed potential of these enzymes to cause transient matrix cleavage

without hydrolysis, thus providing a potential molecular mechanism for controlled, turgor‐driven wall

expansion (Rose et al., 2002).

Indeed, XETs have been implicated in both wall‐loosening and wall‐strengthening, gravitropic

responses, as well as in incorporating nascent xyloglucan into the wall during biosynthesis (Rose et

al., 2002; Cosgrove, 2005). And, while the biomechanical roles of XETs are still being debated

(Cosgrove, 2005; Van Sandt et al., 2007), it is doubtless that XTH gene products are important:

higher plants maintain large XTH gene families (comprised of 20‐60 genes, Fig. 8B), whose members

are actively transcribed in tissue‐, time‐, and stimulus‐dependent contexts (Rose et al., 2002;

Yokoyama et al., 2004; Becnel et al., 2006; Mellerowicz and Sundberg, 2008; Miedes and Lorences,

2009).

To understand the physiological effects of individual XTH gene products in diverse processes such as

seed germination, organogenesis, cell expansion, and fruit ripening, it is essential to understand their

1 This section is a modified version of a review article, where the author of this thesis is the primary author, published in a Focus issue of Plant Physiology (www.plantphysiol.org; Copyright American Society of Plant Biologists). Eklöf JM, Brumer H (2010) The XTH Gene Family: An Update on Enzyme Structure, Function, and Phylogeny in Xyloglucan Remodeling. Plant Physiol. 153: 456‐466 2XTH has alternately been defined as xyloglucan endo‐transglucosylase/hydrolase (Rose et al., 2002), although this is formally incorrect: an entire glycan chain is transferred, not a single glucosyl residue; compare cyclomaltodextrin glucanotransferase, EC 2.4.1.19; 4‐α‐glucanotransferase, EC 2.4.1.25; 1,4‐α‐glucan 6‐α‐glucosyltransferase, EC 2.4.1.24; etc.

Jens Eklöf

21

biochemistry. I.e., it is necessary to know whether a regulated transcript encodes a strict XET devoid

of kinetically relevant hydrolytic activity, a strict XEH with no capacity to rearrange the xyloglucan

network except through irreversible degradation, or an enzyme with mixed function.

In the last decade there have been significant advances in the understanding of protein structure‐

function relationships encoded by the XTH gene family, including the first three‐dimensional

structures of a plant XET (Johansson et al., 2004) and a plant XEH (paper II here referred to as

(Baumann et al., 2007)).

Molecular phylogeny of XTH gene products

An evolving view of families, subfamilies, clades, and clans

The proteins encoded by XTH genes comprise a subfamily of Glycoside Hydrolase family 16 (GH16) in

the Carbohydrate‐Active enZYmes (CAZy) classification (Cantarel et al., 2009), which groups enzymes

on the basis of structural and mechanistic similarity (Davies and Henrissat, 1995; Davies et al., 2005).

GH16 enzymes display a diversity of substrate specificities, with family members cleaving β‐1,3 or β‐

1,4 bonds in various glucans and galactans. GH16 enzymes are also distantly related to the GH7

cellulases within Clan GH‐B3 (Michel et al., 2001; Eklöf, 2010). Within GH16, the XTH gene products

are most closely related to bacterial β‐1,3;1,4‐glucan hydrolases and fungal Crh proteins implicated

in cross linking chitin to β‐1,6 and β‐1,3 glucans (Cabib et al., 2008) (Fig. 8a).

The molecular phylogeny of XTH genes and gene products, which was originally divided into three

major Groups (I, II, and III) (Campbell and Braam, 1999), has undergone continual revision as a

consequence of an ever‐increasing body of sequence information. Indeed, just over ten years ago,

there were ca. 50 XTH gene product sequences (Campbell and Braam, 1999). As a result of

numerous plant genome sequencing projects (http://www.phytozome.org/), this number has

increased to several hundred, with the result that original phylogenetic differences are beginning to

become blurred. Indeed, already in 2004 a comparison of the first two public plant genomes, rice (a

monocot) and Arabidopsis (a dicot), indicated that Groups I and II had become indistinguishable

(Yokoyama et al., 2004). This observation has been mirrored by the analysis of other large data sets

(Baumann et al., 2007; Michailidis et al., 2009).

3 Clans are groups of GH families with a conserved three‐dimensional structure and catalytic machinery despite low amino acid sequence similarity and differing substrate specificity; see http://www.cazy.org/


22

Figure 8. The evolution and distribution of XTH and EG16 genes in publicly available genomes. a, the proposed evolution of Clan B containing GH7 and GH16 (updated from (Michel et al., 2001)). b, a simplified tree showing the representatives of the Viridiplantae (top) and a diagram showing the distribution of XTH and EG16 genes into groups (bottom). The number of genes from each organism is shown on top of each column. The groups are coloured as follows: Group I, black; Group II, light gray; Group III‐A, white; Group III‐B, darkgrey; EG16, tilted stripes. The presence of an EG16 in Charophyta was shown in Paper VI. Genomes accessed in February 2011 from http://www.phytozome.org.

Jens Eklöf

23

The validation of a subfamily rests in statistical robustness, as well as the ability of such groupings to

predict enzyme characteristics and/or in vivo functional differences. Within the composite Group I/II

there are a number of statistically robust clades, although there is no evidence (thus far) that these

clades harbour significantly different activities. Indeed, heterologously expressed XTH genes from

this group have all exhibited exclusively XET activity (Supplementary Table S1, available online from

(Eklöf and Brumer, 2010)), although in most cases donor and acceptor substrate specificities have not

been extensively analysed. For XTH phylogenies to have a predictive power, an emerging picture is

that finer‐grain considerations of clade and sub‐clade groupings should be made, based both on high‐

level sequence clustering algorithms and rigorous biochemical data (Baumann et al., 2007; Atkinson

et al., 2009; Del Bem and Vincentz, 2010).

As a case in point, historical Group III (Campbell and Braam, 1999) can be divided in two, supported

by both sequence analysis and catalytic measurements. The archetypal XEH, Tm‐NXG1 from

nasturtium (Tropaeolum majus), and VaXGH from azuki bean (Vigna angularis) of Group III‐A (Group

nomenclature according to Baumann et al., (2007)) are thus far the only XTH gene products with

demonstrable hydrolytic (XEH) activity (Edwards et al., 1986; Fanutti et al., 1993, 1996; Tabuchi et al.,

2001; Baumann et al., 2007). In contrast, heterologously expressed XTH genes in Group III‐B,

AtXTH27 (Campbell and Braam, 1999), SlXTH5 (Saladie et al., 2006), and HvXTH8 (Kaewthai et al.,

2010) show predominantly or exclusively XET activity, thus validating a functional distinction

between the A and B clades.

Recently, a new clade of plant GH16 enzymes has been described (Fig. 8b, Paper VI). These genes,

called GH16 endo glucanases (EG16s) seem to predate the XTH gene product present already in

charophycean green algae. The product of an EG16 gene from black cottonwood, Populus

trichocarpa showed activity on both β‐1,3;1,4 glucan and xyloglucan, and was suggested to have

shared a last common ancestor with XTH gene products, giving clues to the evolution of XTH gene

products from bacterial lichenases (Paper VI).

Insight from plant genome sequencing

As a result of the 23 publicly available land plant genomes (http://www.phytozome.org/), the

outlines of the XTH gene product family evolution can be visualised (Fig. 8b). Xyloglucan and XTH

genes are not present in all plants. The unicellular green algae of the division Chlorophyta are not

likely to contain xyloglucan (Frei and Preston, 1961), and three available genome sequences reveal

that these organisms do not possess XTH genes (Fig. 8b). In the charophycean green algae more

closely related to land plants, both XET activity, GH16 enzymes and small amounts of xyloglucan have

been found (Popper and Fry, 2003; Ikegaya et al., 2008; Domozych et al., 2009; Domozych et al.,


24

2010). For example, in Chara vulgaris, one of the closest living relatives to the early land plants, a

transcript with high sequence similarity to GH16 β‐1,3;1,4‐glucanases has been found (Van Sandt et

al., 2007). It is tempting to speculate that such a sequence might be the ancestor of an early XTH

gene, due to the similarities between GH16 β‐1,3;1,4 glucanases and XTH gene products (Fig. 8a and

Paper VI).

The oldest plants currently known to have both xyloglucan (Popper and Fry, 2003; Popper, 2008) and

XTH genes are the bryophyte Physcomitrella patens and the early vascular plants (lycophytes)

Selaginella kraussiana (Van Sandt et al., 2006) and Selaginella moellendorffii. The relative abundance

of XTH gene products in the genomes of P. patens and S. moellendorffii (Fig. 8b) indicate that Group I

is likely to be the original XTH gene product subfamily and that Group II and III‐B arose subsequently

in separate events. Particularly notable in these early plants is the lack of the Group III‐A, which

harbours the predominant XEH enzymes Tm‐NXG1 (Paper III), AtXTH31 (Paper IV) and VaXGH (Fig.

8b). These observations are in accordance with Baumann et al., (2007), who suggested that XEH

activity may have developed as a gain of function in an ancestral XET, and also that an “ancestral

group” of sequences (part of Group I) clustered closest to a bacterial β‐1,3;1,4 glucanase.

The structural basis of XET versus XEH activity

A key outstanding question in the functional genomics of XTH genes is, what determines whether a

protein will have XET or XEH activity, or some combination of the two? In 2004, the first

experimentally‐determined three‐dimensional structure of a XET was reported (Johansson et al.,

2004), that of the Populus tremula x tremuloides (hydrid aspen) XET16‐34 (PDB ID 1un1), which

opened the door to addressing this question through “structural biology.” PttXET16‐34, a member of

Group I/II that is highly up‐regulated in wood‐forming tissues, is notably devoid of measurable

hydrolytic activity across a range of xyloglucan concentrations (Kallas et al., 2005; Baumann et al.,

2007). Shortly thereafter, this first XET structure was followed by the structure of the archetypal XEH

from nasturtium, Tm‐NXG1 of Group III‐A (PDB ID 2uwa) (Baumann et al., 2007). Together, these two

structures were instrumental in providing first atomic‐level insight into the molecular mechanisms of

cell wall remodelling and degradation by XTH gene products.

Jens Eklöf

25

Figure 9. Structures of XTH gene products and a closely related GH16 β‐1,3;1,4 glucanase. a, surface representation of Ptt‐XET16‐34 (PDB ID 1un1, 1umz) in silver (C‐terminal extension in copper) with a xylogluco‐oligosaccharide, XLLGXLLG bound (glucosyl backbone in blue and xylosyl and galactosyl units in gold). b, surface representation of a β‐1,3;1,4 glucanase (PDB ID 1u0a) in silver with a loop narrowing the negative subsites in copper with a β‐1,3;1,4 gluco‐tetrasaccharide (G4G4G3G). c, a cartoon representation of Ptt‐XET16‐34 showing the C terminal extension (copper) and catalytic amino acids (green) with XLLGXLLG bound. d, overlay of a XET, Ptt‐XET16‐34 (silver with red loops) and a XEH, Tm‐NXG1 (gold with blue loops; PDB ID 2uwa, 2vh9).

Global features of enzyme structures revealed by X‐ray crystallography

Both the PttXET16‐34 and Tm‐NXG1 structures display the β‐jellyroll fold common to all members of

GH16, but with notable differences that reflect the specialisation of these enzymes toward their


26

highly branched substrate. Compared to the GH16 β‐1,3;1,4‐glucanases, which hydrolyse

unsubstituted polyglucose chains (Planas, 2000), the XETs and XEHs have a much wider substrate

binding cleft due to a major loop deletion in the negative subsites4 of the active‐site cleft (Fig. 9a cf.

Fig. 9b). In the positive subsites, a C‐terminal extension, which has been long recognised as a

distinguishing feature of XET sequences in GH16 ((Campbell and Braam, 1999), InterPro entry

IPR010713), elongates the substrate binding cleft by providing an extra β‐strand at the end of an α‐

helix (Fig. 9c) (Johansson et al., 2004). Strikingly, this motif follows a disorganised leader sequence

extending across the underside of the tertiary structure (Fig. 9c), which may present a hurdle in

protein folding that contributes to the difficulties encountered in the recombinant production of

these enzymes in prokaryotic and eukaryotic hosts (Kaewthai et al., 2010).

The experimentally determined PttXET16‐34 (Johansson et al., 2004) and Tm‐NXG1 (Baumann et al.,

2007) tertiary structures also reveal the structural importance of highly conserved cysteine residues

(Campbell and Braam, 1999), which help to stabilise the C‐terminal extension by formation of two

disulfide bonds. Finally, the PttXET16‐34 structure highlight the structural importance of the N‐

glycan, the core of which interacts tightly with the polypeptide chain (Johansson et al., 2004).

Removal of this N‐glycan, either through site‐directed mutagenesis or enzymatic treatment, greatly

reduces proper protein folding and stability in a number of XETs (Campbell and Braam, 1999; Kallas

et al., 2005). This N‐glycosylation site is conserved in all Group I/II sequences, but is notably absent

in nearly all Group III‐A enzymes such as Tm‐NXG1 (Baumann et al., 2007). In Group III‐B enzymes,

the N‐glycosylation site is shifted toward the C‐terminus, thus placing it on the other side of the

active site cleft (Kallas et al., 2005).

The catalytic amino acids: identification and roles

Although the inclusion of XETs, xyloglucan:xyloglucosyl transferases (EC 2.4.1.207) in a Glycoside

Hydrolase family may seem incongruous, the ability of XETs to catalyse glycosyl transfer is a logical

consequence of the canonical “retaining” catalytic mechanism employed by all members of GH16

(Planas, 2000). A key feature of the retaining mechanism (Fig. 10) is the formation of a covalent

glycosyl‐enzyme intermediate, which can be broken‐down by water, yielding hydrolysis (XEH

activity), or an incoming saccharide substrate, yielding transglycosylation (XET activity; (Sinnott,

1990; Baumann et al., 2007)). Definitive proof of the covalent nature of the glycosyl‐enzyme has

4 The negative subsites are those which bind the polysaccharide from the point of cleavage toward the non‐reducing end of the polysaccharide, according to the nomenclature of (Davies et al., 1997); see Fig. 10.

Jens Eklöf

27

been obtained through kinetic trapping and direct observation of this intermediate in PttXET16‐34 by

mass spectrometry (Piens et al., 2008).

Figure 10. A schematic representation of the mechanism used by XETs and XEHs. Left, xyloglucan binds to XETs and XEHs in both negative and positive subsites (glucosyl units of xyloglucan in blue and xylosyl units in orange). After binding, the substrate is cleaved resulting in an enzyme‐glycosyl intermediate. In the last step the glycosyl‐enzyme is broken down by an incoming acceptor, either by water (XEH activity) or by the non‐reducing end of a xyloglucan molecule (XET activity).

The catalytic motif in all XTH gene products is 83(H/W/R)-(D/N)-E-(I/L/F/V)-D-

(F/I/L/M)-E-(F/L)-(L/M)-G92 (catalytic residues in bold, most frequent amino acids

underlined; PttXET16‐34 numbering). The first glutamate (E85 in PttXET16‐34) is known as the

“catalytic nucleophile,” which attacks the anomeric carbon of the glucose ring in subsite ‐1 to yield

the glycosyl‐enzyme in the first step of the catalytic reaction. E89 functions alternately as a catalytic

general acid to protonate the xyloglucan chain fragment departing the positive subsites, and as a

catalytic general base to activate the incoming acceptor substrate (water, XG, or XG oligosaccharide).

The specific role of D87, which lies on the same face of the beta‐strand as E85 and E89 (Fig. 9c,

highlighted in green), is currently unknown, although hydrogen bonding patterns (Johansson et al.,

2004; Mark et al., 2009) suggest that this residue might modulate the pKa of the catalytic nucleophile,

as has been previously suggested for other GH16 enzymes (Planas, 2000).

The identities of these catalytic residues, conserved in all GH16 enzymes, are known from elegant

mutagenesis, kinetic, and structural studies on bacterial endo‐glucanases (reviewed in (Planas, 2000;

Eklöf, 2010)). Indeed, the catalytic residues in the ExDxE motif of Bacillus sp. endo‐glucanases are

exactly superimposable onto those of both PttXET16‐34 and Tm‐NXG1 (Johansson et al., 2004;

Baumann et al., 2007). Confirmation of the identity of the catalytic nucleophile in GH16 XETs and

XEHs comes from the observation that mutation of the corresponding glutamate in both PttXET16‐34

and Tm‐NXG1 results in the complete loss of the wild‐type activity; these nucleophile variants are

nonetheless able to perform “glycosynthase” reactions, i.e. the re‐synthesis of xyloglucans from

artificial donor substrates (α‐fluoroglycosides of XG oligosaccharides; (Piens et al., 2007; Gullfot et


28

al., 2009)). Furthermore, mutation of E85 in PttXET16‐34 to alanine prevents formation of the

covalent glycosyl‐enzyme (Piens et al., 2008). The hydrophobic sidechains of the intervening

residues in the catalytic motif do not play a direct role in catalysis, as these point into the interior of

the β‐jellyroll structure, where they fulfil a structural function.

Differences in loop structures lining the active‐site cleft affect XET versus XEH activity.

Initial comparisons of the PttXET16‐34 structure with the Bacillus sp. β‐1,3;1,4‐glucanase in fact

revealed very little about the determinants of transglycosylation versus hydrolysis in XTH gene

products, due to the exact superposition of many key active‐site residues (Johansson et al., 2004).

The stability of the PttXET16‐34 glycosyl‐enzyme to hydrolysis in dilute aqueous solution (56 M H2O)

is indeed remarkable: kinetic trapping studies placed the hydrolytic half‐life of this intermediate at

ca. 3 hours (khydr = 0.005 min‐1) in the complete absence of an XG acceptor substrate (which could not

occur in vivo), while the addition of XG oligosaccharides rapidly turned‐over the glycosyl enzyme (in <

2 min) (Piens et al., 2008). This behaviour stands in stark contrast to the measured hydrolytic rate of

Tm‐NXG1 on tamarind xyloglucan (apparent kcat 5.5 min‐1), whose glycosyl‐enzyme is thus rapidly

broken‐down by water (Baumann et al., 2007). The three‐dimensional structures of these two

enzymes revealed that much of this difference in catalytic properties is the result of subtle variations

in active‐site loop structures, as follows.

Overall, the structures of PttXET16‐34 and the archetypal XEH Tm‐NXG1, as indicated by

superposition of their backbone Cα traces, are very similar. The obvious differences between these

enzymes lay in two loops lining one side of the substrate binding cleft; Loop 1 lines the ‐1 and ‐2

subsites while Loop 2 lines the +1 and +2 subsites (Fig. 9d). A three amino acid insertion in Loop 1 is

common to members of both Groups III‐A and III‐B, relative to Group I/II members, while the length

of the insertion in Loop 2 varies between Groups III‐A and III‐B: Group III‐B possesses a two amino

acids insert (Xaa-Gly) in Loop 2 compared to Group I/II, while this insert in Group III‐A is five

amino acids long, with the consensus sequence (Gly-Arg)-Asn-Ile-Ile-Gly.

The importance of the length of Loop 2 in tuning the balance between XET and XEH activity was

demonstrated through the generation of a chimeric enzyme, Tm‐NXG1‐ΔYNIIG (Baumann et al.,

2007). This chimera, in which the shorter Loop 2 of the PttXET16‐34 was presented on the Tm‐NXG1

scaffold (PDB ID 2uwb), showed intermediate kinetics: With respect to the Tm‐NXG1 wild‐type, XEH

activity was reduced two‐fold, while XET activity was doubled to reach half that of PttXET16‐34 at

saturating substrate conditions (Baumann et al., 2007). Thus, while loop 2 is one of the important

determinants for hydrolysis versus transglycosylation, there are clearly other yet unidentified factors

governing the fate of the glycosyl‐enzyme intermediate. In particular, further protein engineering

Jens Eklöf

29

studies are needed to completely remove the hydrolytic capacity of the Tm‐NXG1 protein scaffold,

thereby converting it into a strict (or overwhelmingly predominant) XET.

Molecular dynamics simulations based on crystallographic complexes of PttXET16‐34 and Tm‐NXG1

with XG oligosaccharides (Mark et al., 2009) and kinetic measurements (Saura‐Valls et al., 2008)

indicate that PttXET16‐34 may bind XG substrates more tightly in the positive subsites, while Tm‐

NXG1 binds these substrates more strongly in the negative subsites. Small changes in protein‐

substrate interactions, together with subtle changes in protein dynamics, are thus likely to be

additional contributors affecting XET versus XEH activity. Indeed, there are certain parallels to be

explored between xyloglucan activity in GH16 and the diverse starch hydrolases and

transglycosylases (van der Maarel et al., 2002). These include the potential of induced‐fit

mechanisms to properly orient the active‐site residues and conformational changes to effect leaving

group departure and acceptor substrate binding (Uitdehaag et al., 2000).

How XETs and XEHs recognise their natural substrates

Length of the active site

In addition to possessing a wide active‐site cleft, which is capable of accommodating brush‐like

xyloglucan chains (Fig. 9a), GH16 XETs and XEHs have a number of structural features that form

specific interactions with the polysaccharide to bring it “in register” for cleavage by the catalytic

amino acids. Ligand structures of PttXET16‐34 with substrate bound in the positive subsites (PDB ID

1umz) and Tm‐NXG1 with substrate bound in the negative subsites (PDB ID 2vh9) provide visual

evidence that XTH gene products have a 35 Å long active site cleft, comprised of primary subsites for

seven backbone glucosyl units (‐4 to +3, Fig. 9a & 9c) (Mark et al., 2009). The glucan backbone binds

tightly in the cleft, primarily through hydrophobic stacking interactions mediated by three

tryptophans and a tyrosine in the ‐3 to +2 subsites (Trp 19, 174, 179 and Tyr75, PttXET16‐34

numbering). Composite models from molecular dynamics simulations suggest that the unbranched

glucosyl unit in the ‐1 subsite is forced into a strained skew‐boat conformation in the enzyme‐

substrate complex (Mark et al., 2009) as a prerequisite for catalysis (Davies et al., 1998).

While structural and modelling data only show that XETs and XEHs have 7 primary subsites, it has

been shown that longer substrates give a slight increase in transglycosylation rate (Purugganan et al.,

1997; Saura‐Valls et al., 2008)(corresponding to a transition state binding energy of only 0.5 kcal/mol

for e.g., XXXG3 vs. XXXG2 (Saura‐Valls et al., 2008)). This increase cannot be accounted for by direct

enzyme‐substrate interactions and may be due to entropic reasons, i.e., better ordering of sugar

residues near the ends of the active‐site cleft in longer substrates. Nonetheless, it should be clarified

that, contrary to earlier suggestions (Rose et al., 2002), there is no firm evidence to suggest that


30

GH16 XETs and XEHs generally require substrates longer than the minimal donor XXXGXXXG (Tabuchi

et al., 1997; Baumann et al., 2007; Ibatullin et al., 2008).

Contributions of branching glycosyl units to substrate recognition and catalysis.

The effect of branching substitutions on the xyloglucan backbone on the interaction of PttXET16‐34

with its substrate has been assessed using a library of 18 synthetic xylogluco‐oligosaccharides (Saura‐

Valls et al., 2008). The design and generation of this library alone was a tour de force in carbohydrate

synthesis (Faure et al., 2006), as was the determination of full kinetic parameters (kcat and Km values)

for nearly all substrates under initial‐rate conditions (Saura‐Valls et al., 2006; Saura‐Valls et al., 2008).

The assay employed a highly sensitive capillary electrophoresis method to analyse the products from

the XET catalysed glycosyl transfer from donor substrates onto a fluorescent 8‐aminonaphthalene‐

1,3,6‐trisulfonate (ANTS)‐labelled acceptor substrate, e.g.: XXXGXXXG (donor) + XXXG‐ANTS

(acceptor) → XXXGXXXG‐ANTS + XXXG. Variation of the donor backbone length and the pattern of α‐

1,6‐xylose substitution allowed elucidation of the specific contribution of individual glycosyl units to

transition state stabilisation (ΔΔG≠) in the substrate binding cleft of PttXET16‐34.

Interestingly, the singular influence of many α‐1,6‐xylosyl units is small: xylose substitution of the

glucosyl units in subsites ‐4, ‐3, ‐2, +1, and +3 (Fig. 9a & 9c) affected the specificity parameter kcat/Km

by a factor of 2 or less, which equates to <0.5 kcal/mol of transition‐state stabilisation (see Schemes

4‐6 in (Saura‐Valls et al., 2008)). The xylosyl unit in the +2’ subsite (i.e., attached to Glc(+2)) is

however crucial to activity, contributing ca. 4 kcal/mol to catalysis, which equates to an increase in

the kcat/Km (rate) value more than 500‐fold over substrates lacking this residue (Saura‐Valls et al.,

2008). The obvious importance of Xyl(+2’) reflects earlier studies on Tm‐NXG1 from Group III‐A

(Fanutti et al., 1996) and XET preparations extracted from pea (Pisum sativum) (Fry et al., 1992;

Lorences and Fry, 1993), thus highlighting that Xyl(+2’) recognition is a major specificity determinant

common to both XETs and XEHs of GH16. Interestingly, despite species‐ and tissue‐ dependent

heterogeneities in xyloglucan sidechain decorations, a xylosyl unit is always present two glycosyl unit

after an unsubstituted glucosyl unit (Hoffman et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009).

And while Xyl(+2’) clearly dominates in the XG‐XET interaction, it should be borne in mind that the

additive effects of several xylosyl units on the glucan backbone are ultimately responsible for XG

specificity in both the positive and negative subsites of XTH gene products.

Due to the complexities of obtaining pure donor substrates with further specific sidechain

elaboration, the contributions of pendant galactosyl, fucosyl, and arabinosyl units has not been

examined to the same extent as xylosylation. In a few cases, galactosylation and

(fuco)galactosylation have also been shown to influence XET activity, but to a limited extent (Fry et

Jens Eklöf

31

al., 1992; Lorences and Fry, 1993; Maris et al., 2009; Maris et al., 2011). For example, substrate

competition assays of a pea XET preparation indicated an acceptor preference of XLLG > XXXG >

XXLG, although the difference in catalytic rates between the best and least good substrates was ca.

2‐fold (Fry et al., 1992), again reflecting a minimal contribution to transition state binding.

Interestingly, a very crude protein preparation from Arabidopsis showed reduced XET activity on

xyloglucan extracted from mur3 plants, which contain only XXXG repeats lacking galactosyl and

(fuco)galactosyl sidechains (Peña et al., 2004). As with the pea XET study, however, the analysis of

unpurified enzyme mixtures requires that these results be interpreted with some caution. In the

archetypal XEH, Tm‐NXG1, the presence of galactosylation had little to no effect on the rate of

hydrolysis when comparing XXXG‐ versus XLLG‐based substrates. In contrast, xylosylation

(comparing GGGG and XXXG substrates) boosted specificity, kcat/Km, over 2 orders of magnitude

(Ibatullin et al., 2008).

These specificity data can be nicely mapped onto the experimental three‐dimensional structures of

XET and XEH active sites (Mark et al., 2009). As a specific example, the +2’ xylose‐binding subsite

discussed above is a hydrophobic pocket comprised of a tyrosine and two arginine sidechains

(Arg116, Tyr250 and Arg258 in PttXET16‐34). The C‐terminal extension, unique to the XTH gene

products, comprises two thirds of this hydrophobic pocket by contributing with the strictly conserved

Tyr250 and Arg258 (Fig. 9a & 9c, see (Mark et al., 2009) for full structural details and (Baumann et al.,

2007) for sequence alignments). The third amino acid in the hydrophobic pocket, Arg116, is

conserved within Groups I and III, whereas in Group II, Arg116 is replaced by a glutamine; the

functional significance of this change, if any, is currently unknown. As is typical for carbohydrate

active‐enzymes (CAZymes) and binding modules (CBMs), XTH gene products employ a multitude of

hydrophobic ring‐stacking interactions and hydrogen bonds with sugar hydroxyl groups to recognise

and bind their substrates. A complete survey of all active‐site interactions is, however both

challenging and tedious (Mark et al., 2009): the minimal substrate XXXGXXXG spanning the active

site is comprised of 14 monosaccharide units, presenting a remarkable 38 hydroxyl groups and 28

glycosidic and ring‐oxygens, each with 2‐3 hydrogen‐bond donor/acceptor sites each!

Alternate substrates, and activities, for XTH gene products

Hetero‐transglycosylation gives linear conjugates of unsubstituted β‐glucans and xyloglucan

The grassy species of the order Poales have cell walls with relatively low amounts of xyloglucan

compared to the dicotyledons (Fincher, 2009; Hsieh and Harris, 2009). Despite this, the XTH gene

family of all angiosperms are of similar sizes (Fig. 8b) and XTH gene transcripts in the Poales species

Hordeum vulgare (barley) are abundant in cell wall remodelling libraries (Strohmeier et al., 2004).


32

Work by Hrmova, Fincher, and co‐workers has shown that a pure barley XET has the quantifiable

ability to conjugate xyloglucan with other β‐glucans (Hrmova et al., 2007). Thus, HvXET6 was able to

use the substrate donor/acceptor pairs xyloglucan/cello‐oligosaccharides and β‐1,3:1,4‐

glucan/xyloglucan‐oligosaccharides at rates of 0.1% and 0.2% of the wild‐type

(xyloglucan/xyloglucan‐oligosaccharides) activity, respectively. The enzyme was also able to use

hydroxyethylcellulose and sulphuric acid‐swollen cellulose as glycosyl donors with xyloglucan‐

oligosaccharide acceptors at 44% and 5% of the wild‐type rate, respectively. These observations

prompted the proposition that XTH isoforms may build xyloglucan‐(β‐1,3:1,4‐glucan) links that fortify

the cell wall in grass species (Hrmova et al., 2007). Given the comparatively low activities observed

for hetero‐transglycosylation, supporting this hypothesis with in vivo data may present a significant

experimental challenge.

Low‐level hetero‐transglycosylation activities may, in fact, be a general feature of XTH gene products,

due to inherent active‐site promiscuity. Work preceeding the study described above detected

xyloglucan/β‐glucan conjugation activity in crude extracts from germinating nasturtium seeds

(Mohand and Farkas, 2006). Although the enzyme responsible for this activity was not identified,

later studies indicated that pure nasturtium Tm‐NXG1 could use cello‐oligosaccharide acceptors to

generate preparative amounts of xyloglucan conjugates (Baumann et al., 2007). Likewise, the subsite

mapping studies described above showed that PttXET16‐34 could use both GGGG•XXXG and

XXXG•GGGG (and shorter congeners) as glycosyl donors (“•” indicates the point of cleavage), thus

indicating that xylosylation is not an absolute requirement for activity (Saura‐Valls et al., 2008).

Indeed, the selectivity (kcat/Km value) for XXXG•GGGG was ca. 0.1% of the value of the “wild‐type”

substrate XXXG•XXXG, which mirrors the results described above for HvXET6, while that for

GGGG•XXXG was 65% of the XXXG•XXXG value. Futhermore, on the basis of experimental structural

data (Johansson et al., 2004; Baumann et al., 2007; Mark et al., 2009), there is no steric reason why

XETs and XEHs would exclude skinnier substrates from their active‐site clefts, although the loss of Xyl

sidechain binding, especially Xyl(+2’), would be expected to penalise catalysis, as described above.

Notably, C‐6 oxidation of xyloglucan backbone Glc residues to glucuronyl residues using the reagent

TEMPO destroys the ability of the polysaccharide to act as a donor substrate in the first step of the

XET reaction (Fig. 10) (Takeda et al., 2008). Structural analysis indicates that the presence of a uronic

acid may disrupt hydrogen bonding patterns implicit in binding the donor substrate and forcing it

into a skew‐boat conformation in the Michealis (E•S) complex (Mark et al., 2009). The observation

that TEMPO‐oxidised xyloglucans retain the ability to act as glycosyl acceptors is rationalised by the

observation that Glc(+4) extends beyond the end of the active‐site cleft (Johansson et al., 2004).

Jens Eklöf

33

The case for a specific “mixed‐linkage β‐glucan:xyloglucan endotransglucosylase (MXE)”

An enzyme activity equivalent to one proposed for barley HvXET6, i.e mixed‐linkage β‐

glucan:xyloglucan endotransglucosylase (MXE), has been found in Equisetum species (horsetails) as

well as in the closest living ancestors of land plants, the Charophyta (Fry et al., 2008). MXG activity in

the species tested was indeed significant, in some cases reaching equivalent values for

xyloglucan:xyloglucosyl transferase (XET, EC 2.4.1.207) activity in crude extracts. In contrast, MXE

activity was deemed negligible in a range of land plants tested, including a selection of bryophytes,

pteridophytes, angiosperms and a lycopodiophyte. For example, an extract of local Yorkshire fog

grass (Holcus lanatus) exhibited a relative MXG/XET activity ratio of 0.17% (Fry et al., 2008), similar to

HvXET6 from barley (Hrmova et al., 2009) – also a member of the order Poales.

A series of biochemical arguments have been put forth to suggest that the observed MXE activity is

not due to an XTH gene product. However, definitive proof, as well as the possibility to engineer this

activity into cereal crops to improve cell wall properties (Fry et al., 2008), will have to await the

cloning and heterologous expression of the gene encoding this activity, as well as full kinetic

characterisation of the recombinant enzyme. Cautious optimism is warranted, since similar enzyme

discovery experiments using specific transglycosylation assays have identified “microbial XETs,”

which have later turned out to be predominant (xylo)glucan endo‐hydrolases (Nielsen, 2000).

Indeed, any number of endo‐β‐glucanases employing the canonical retaining mechanism could

possess MXE side activity, although the low apparent Km value (4 μM) for the alditol acceptor

substrate XXXGol certainly speaks for the existence of a specific enzyme (Fry et al., 2008). If not an

XTH gene product, it will be interesting to see if this enzyme belongs to Glycoside Hydrolase Family

17 (which includes β‐1,3(4)‐endo‐glucanases (Varghese et al., 1994)), another endoglucanase family

(Gilbert et al., 2008), or eventually defines a new CAZy family (Cantarel et al., 2009).

Other transglycosylases in plants

The idea that transglycosylases could play a key role in primary cell wall extension was first

articulated in the mid‐1970’s (Albersheim, 1976). Since that time, XETs i.e. xyloglucan:xyloglucosyl

transferases, have received by far the most attention in this respect, although as highlighted above,

other activities are beginning to be explored. In particular, the potential for transglycosylation has

been revealed in a tomato endo‐β‐mannanase (Schröder et al., 2009) from Glycoside Hydrolase

Family 5, members of which use the canonical retaining mechanism (Gilbert et al., 2008). Indeed, the

transglycosylation potential of GH5 enzymes has been long known (Harjunpää et al., 1999), although

a full 3‐D structure/kinetic analysis is sorely needed to highlight key active‐site features (Bourgault et

al., 2005) that might allow functional prediction based on sequence phylogeny.


34

It is noteworthy that surveys of available plant genomes reveal many families of retaining glycoside

hydrolases (Henrissat et al., 2001; Geisler‐Lee et al., 2006), the numerous members of which all have

the implicit potential to catalyse transglycosylation reactions (Sinnott, 1990). Taking Arabidopsis as

an example, 23 GH families use the retaining mechanism (see http://www.cazy.org/Genomes.html).

Of these, 15 GH families are likely to be exo‐enzymes, which act on terminal monosaccharide units.

Discounting predicted starch‐ and chitin‐cleaving enzymes, four known retaining GH families contain

demonstrated endo‐polysaccharidases: GH5 (mannanases, glucanases, galactanases), GH10

(xylanases), GH16 (xyloglucanases), and GH17 (β‐1,3 glucanases). In contrast, for example, the 25

GH9 members (predicted endo‐glucanases (Urbanowicz et al., 2007)) and 67 GH28 members

(predicted galacturonidases) from Arabidopsis utilise the inverting glycosidase mechanism, which

only allows polysaccharide hydrolysis (Sinnott, 1990). It will be interesting to see if future functional

studies uncover new CAZy families in plants with polysaccharide re‐arrangement activities.

A future for “oldfashioned” enzymology in these modern times of highvolume functional

genomics?

The relation of in vitro enzyme kinetics and structure‐function studies to effects in vivo can prove to

be a powerful tool in functional genomics studies, although such extrapolations are sometimes

viewed with scepticism. It is arguably better to know as many details of catalysis as possible when

postulating the effect of an enzyme on plant physiology, and this is certainly true in the case of the

disparate activities of polysaccharide hydrolysis and transglycosylation. For example, a XET which

does not measurably hydrolyse xyloglucan in in vitro assays containing 56 M H2O (Supplementary

Table S1 (Eklöf and Brumer, 2010)) is unlikely to do so in the hydrogel matrix of the cell wall, where

the water activity may be even lower. Likewise, the observation that nasturtium xyloglucanase 1

(Tm‐NXG1) is hydrolytically inefficient (Ibatullin et al., 2008) and only transglycosylates at very high

XG oligosaccharide concentrations (Baumann et al., 2007) might imply a product inhibition/feedback

mechanism during seed germination.

Such relationships, however, require careful quantitation of kinetic rates across a range of substrate

concentrations. Where possible, competing reactions, such as hydrolysis and transglycosylation

should be measured directly in a single assay mixture (e.g., (Harjunpää et al., 1999; Baumann et al.,

2007)). Comparatively insensitive and semi‐quantitative assays, such as those based on viscometry,

should be avoided in favour of strictly quantitative assays that yield true catalytic turnover numbers,

e.g., radiometric and fluorometric assays of transglycosylation (Fry et al., 1992; Saura‐Valls et al.,

2006; Hrmova et al., 2007) and reducing sugar assays of hydrolysis (Kallas et al., 2005; Hrmova et al.,

2009). “Chemical biology” can also play a key role in coupling fundamental enzymology with

phenotypic and functional genomics studies: in situ fluorescence assays are currently available for

Jens Eklöf

35

both XET (Vissenberg et al., 2000) and XEH (Ibatullin et al., 2009) activity, and similar methodology

has recently been used to examine plant GH9 endo‐glucanase (KORRIGAN1 homolog) over‐

expression in vivo (Takahashi et al., 2009).


36

Jens Eklöf

37

Results

“The temptation to form premature theories is the bane of our profession”

Sherlock Holmes, The Valley of Fear


38

Microbial Xyloglucan Degrading Enzymes

Aim of investigation

The saccharification of non‐starch cellulose‐based plant substrates for biofuels suffers from the

recalcitrance of the cell wall (Gilbert et al., 2008; Gilbert, 2010). Cellulose microfibrils in plant cell

walls are coated by hemicelluloses which encumber access of cellulases to their substrate (Vincken et

al., 1994). Xyloglucanases represent one class of “hemicellulases” that could be used as accessory

enzymes in the saccharification processes to facilitate access to cellulose. A better understanding of

the biochemical properties of these enzymes and how they recognise their substrates are important

for an improved theoretical knowledge of biosaccharification. Finding new enzymes with desired

properties allow making tailor made xyloglucans for various biotechnological industrial applications

such as artificial blood vessels (Bodin et al., 2007), adhesives or barrier coatings (Mishra and

Malhotra, 2009; Kochumalayil et al., 2010).

Structural convergent evolution of xyloglucanases

Xyloglucanases are found in several GH families including GH5, 7, 12, 16, 44 and 74 (Gilbert et al.,

2008; Vlasenko et al., 2010). In Paper I, ligand structures of a GH44 endo‐xyloglucanase, PpXG44 are

presented together with biochemical characterisation. In Paper II, the first structural representatives

of GH5 and GH12 endo‐xyloglucanases are presented together with biochemical characterisation.

These two papers together with Papers III & VI represent structurally unrelated xyloglucanases from

different GH families that have evolved specificity towards xyloglucan. This convergent evolution

highlights the importance of harnessing this polysaccharide in Nature (Fig. 11).

Figure 11. Ligand structures of three bacterial xyloglucanases.

On top, overlay of ligand structures of PpXG44 in complex with the inhibitor Glc‐Glc‐Oxazine (‐3 to ‐1) and a GXGGG xylogluco‐oligosaccharide (+1 to +5).

Bottom from left, PpXG5 in complex with GXXG in the ‐4 to ‐1subsites and BlXG12 in complex with GXXG XX from ‐4 to +2.

Jens Eklöf

39

Probing substrate specificity in xyloglucanases

While polysaccharide specificity is useful to determine what an enzyme is active on the use of

specific aryl β‐glycosides can elucidate what determines that specificity. In Papers I & II, the

comparison of GGGG, XXXG and XLLG‐CNP (2‐chloro‐4‐nitrophenol) showed to what extent these

different xyloglucanases can harness xyloside and galactoside substituents of xyloglucan for catalysis.

Of the three enzymes, only PpXG5 utilised the side chain decorations for catalysis by increasing

kcat/Km 85‐fold due to xylosylation. In BlXG12 and PpXG44, xylosylation decreased kcat/Km ca. 5‐6‐fold.

PpXG5 also benefited from galactosylation but to a minor extent (Table 1).

Table 1. Substrate specificity of PpXG5, BlXG12 and PpXG44 on various aryl β glycosides.

While both PpXG5 and BlXG12 cleave xyloglucan after unsubstituted glucosyl units, PpXG44 displayed

an unusual cleavage pattern on XXXGXXXG. The main products were XXX and GXXXG and this is the

first xyloglucanase shown to preferentially cut xyloglucan with an X motif in the ‐1 subsite and a G in

+1. When the xylose on the non‐reducing end of XXXG2 was cleaved off by an α‐xylosidase, the

cleavage pattern changed and PpXG44 cleaved GXXGXXXG at roughly equal rates at GXXG as after

GXX, reflecting the affinity for an unsubstituted glucosyl unit in the ‐4 subsite as well as the

importance of a +5 stacking interaction seen in the crystal structure (Fig. 11). By cleaving xyloglucan

with a G in the +1 subsite, G motifs would be present in the ‐4 and +5 subsites due to the regular

spacing of this motif in xyloglucan. Harnessing the ‐4 and +5 subsites optimally probably makes this

cleavage mode even more pronounced on polymeric xyloglucan. In light of this data, the kinetic

parameters for the XGO‐CNPs of PpXG44 should be seen as the data for a less favoured cleavage

mode since a xyloside substituent is present in the ‐4 subsite that would clash with the protein if the

glucan chain would take the preferred trajectory.

What makes a xyloglucanase?

A xyloglucanase could either be defined as an enzyme that have a higher catalytic efficiency on

xyloglucan compared to other polysaccharides or an enzyme that can utilise the sidechain

decorations to improve catalysis as seen for PpXG5. The problem with using polysaccharides for this


40

distinction is that the solubility of unsubstituted β‐1,4 glucan is poor. Indeed, already for

cellohexaose, the solubility is in the µM range and comparing rates of hydrolysis on solid cellulose

and water soluble xyloglucan to determine specificity is not feasible. Instead natural water soluble

glucans such as MLG and glucomannans or artificial polysaccharides such as hydroxyethyl cellulose

(HEC) or carboxymethyl cellulose (CMC) are used. Even though they are water soluble, they

represent a version of xyloglucan that can be seen as Nature’s water soluble cellulose.

For all three enzymes, xyloglucan is the best substrate under the conditions tested and the enzymes

can therefore be called xyloglucanases. These three enzymes span a wide range of xyloglucanases

with PpXG5 representing a specific and fast xyloglucanase. BlXG12 is a rather slow xyloglucanase

that can hydrolyse several polysaccharides at comparable rates. PpXG44 is intermediate and can, as

BlXG12 be seen as “xyloglucan tolerant”.

Jens Eklöf

41

Plant GH16 Enzymes

Aim of study

One of the long term goals in the Carbohydrate Enzymology group at KTH has been to decipher the

underlying mechanisms for transglycosylation (XET activty) versus hydrolysis (XEH activity) in closely

related XTH gene products. In line with the previous objective was an intent to map XET and XEH

activity on a phylogenetic tree to improve functional annotation of XTH gene products.

Understanding the mechanism of transglycosylation versus hydrolysis is not merely of an academic

interest into the enzymology of transglycosylation. Since these two different activities have

drastically different effects on xyloglucan they also differentially influence plant cell wall

characteristics and ultimately, plant growth.

Alongside the above mentioned goals was a personal interest in the evolution of the XTH gene

products which led to the discovery of the EG16s (vide infra).

Activity of plant GH16 enzymes

A transglycosylating activity similar to the XETs of the XTH gene products was for a long time only the

brainchild of Peter Albersheim (Albersheim, 1976), proposed as a possible actor in plant cell wall

modification. In the early 1990’s, XET activity was independently demonstrated experimentally by

three groups (Farkas et al., 1992; Fry et al., 1992; Nishitani and Tominaga, 1992). Another XTH gene

product, Tm‐NXG1 displayed mainly hydrolytic activity on xyloglucan but with some

transglycosylating activity (Edwards et al., 1986; Fanutti et al., 1996).

Since XTH gene products had been shown to exhibit XET, XEH or a mixed type of activity a method to

measure both XET and XEH activity at the same time for a quantitative comparison was needed for

the investigation of transglycosylation versus hydrolysis. Available methods could only measure XET

or XEH activity (Fry et al., 1992; Kallas et al., 2005; Saura‐Valls et al., 2006; Hrmova et al., 2007) and

no assay could measure these two activities simultaneously. For this purpose a high‐performance

anion‐exchange chromatography with pulsed amperometric detection (HPAEC‐PAD) based method

using a minimal substrate, a Glc8‐based XGO2, was developed in Paper III. In a transglycosylating

event, an XGO2 is cleaved to give an XGO1 and a glycosyl‐enzyme intermediate as the first step (Fig.

12). In the final step, a second XGO2 breaks the glycosyl‐enzyme intermediate making an XGO3. The

products of a transglycosylation event are therefore one XGO1 and one XGO3 from two XGO2. In a

hydrolytic event, an XGO2 is hydrolysed to two XGO1 (Fig. 12). Hence, the method relies on the

simultaneous quantification of XGO1 and XGO3 separated by HPAEC‐PAD. Transglycosylation (XET

activity) is quantified by the amount of XGO3 built up. Hydrolysis (XEH activity) is calculated from the


42

total amount of XGO1 subtracted by the amount of XGO3 and divided by 2 since two XGO1 are made

in each XEH reaction (Figure 12).

Figure 12. Outline of the HPAEC‐PAD assay for simultaneous measurement of XET and XEH activity. A XET reaction results in an XGO3 and an XGO1 from two XGO2, while a XEH reaction hydrolyses an XGO2 into two XGO1.

This assay was used in Papers III‐V to determine the kinetic parameters for XET and XEH activity of

four different XTH gene products (Table 2). A few things should be kept in mind when interpreting

data from the XGO2‐based assay. The assay might give the impression that the rate of

transglycosylation is higher than it actually would be in polymeric xyloglucan. In XGO2 the ratio

between cleavable bonds and non‐reducing ends is 1:1. In tamarind xyloglucan (Mw ca. 1 MDa) the

ration is close to 800:1. This means that the concentration of non‐reducing ends in a typical assay is

1 µM (tamarind xyloglucan 1 g/L) and makes transglycosylation events rare in the hydrolytic Tm‐

NXG1 and AtXTH31 with the apparent Km being 500 and 4500 times above that concentration

respectively (Table 2). Comparing the kinetic constant between the characterised XETs and XEHs the

results are fairly similar. Notably, the differences in the XETs are mainly reflected in the apparent Km,

while the XEHs vary mainly in apparent kcat. However, with a sample size of two enzymes in each

group it is not necessarily a general trend.

Table 2. Apparent kinetic constants of XTH gene products on XGO2.

The substrate specificity of the XTH gene products, Tm‐NXG1 (Paper III), AtXTH31 (Paper IV),

AtXTH12‐13 and AtXTH17‐19 (Paper V) were tested on several polysaccharide substrates. They were

Jens Eklöf

43

all shown to be very specific to xyloglucan. While the hydrolytic enzymes Tm‐NXG1 and AtXTH31 had

no measurable activity on other polysaccharides under the conditions tested, the AtXTHs in Paper V

all showed some activity with different cellulose derivatives as donor substrates and with XGOs as

acceptors, but virtually no activity at all on MLG. Interpreting the ability of the AtXTHs to use various

artificial water soluble cellulose derivatives as donor substrates into an in vivo function of XTH gene

products is difficult. Xyloglucan can be seen as Nature’s water soluble cellulose derivative and made

soluble by the addition of short oligosaccharide‐branches instead of the chemical groups in

hydroxyethyl cellulose (HEC), carboxymethyl cellulose (CMC) and water soluble cellulose acetate

(WSCA).

The donor specificities of the AtXTHs in Paper V was also probed with a variety of different [3H]XGO

derivatives and with [3H]cellohexaitol. The various donor substrates showed that further

substitutions on a minimal XGO, XXXG, has minor effects to the rate of transglycosylation while the

non‐xylosylated cellohexaitol, was a poor substrates with severely reduced activity. These results are

in line with a previous study on Ptt‐XET16‐34 (Saura‐Valls et al., 2008), where a library of XGOs was

probed to determine the contribution to catalysis of individual monosaccharide substituents.

Despite the relatively low activity with cello‐oligosaccharides as acceptors, XGO1‐cello‐

oligosaccharide products could be prepared using xyloglucan as the acceptor with the mainly

hydrolytic Tm‐NXG1 (cello‐oligosaccharide [S] 8 mM; Paper III). The effect in the negative subsites of

xyloside and galactoside substituents have been assessed with chromogenic substrates on Tm‐NXG1

(Ibatullin et al., 2008). The galactosylated XLLG‐CNP showed that double galactosylation had a slight

detrimental effect to hydrolysis compared to XXXG‐CNP (kcat/Km reduced by a factor of 2). GGGG‐

CNP showed more than a 100‐fold reduction in kcat/Km, again indicating that the specificity for

xyloglucan over cellulose in both the negative and the positive subsites in XTH gene products is

mainly an effect of xylosylation (Ibatullin et al., 2008; Saura‐Valls et al., 2008).

In vivo localisation of XEH activity in Group IIIA knockouts in Arabidopsis

In Paper IV, two orthologs in Group III‐A of the hydrolytic Tm‐NXG1, AtXTH31 and AtXTH32 were

identified in Arabidopsis. To investigate the impact of the hydrolytic Group III‐A members on the

phenotype of Arabidopsis, a T‐DNA double knockout line of AtXTH31 and AtXTH32 was created.

Somewhat disappointing, but not necessarily surprising, these plant lines showed no obvious

phenotype. To investigate these plants in more depth, XET and XEH activity was compared between

the wild‐type and the double knockout plants in an in situ assay using fluorescent and fluorogenic

probes. The incorporation of XXXG‐sulforhodamine (XET activity) into the cell walls was almost

identical in roots and etiolated hypocotyls of the double knockout and wild‐type lines. The only

observed differences were found just below the cotyledons where the XET activity signal was


44

stronger in wild‐type plants. The XEH activity, visualised for the first time in situ in Arabidopsis

(Paper IV) by the cleavage of XXXG‐resorufin in roots and etiolated hypocotyls, showed significant

differences between the wild type and double knockout plants. In general, XEH activity was severely

reduced in the roots of knockout plants with the largest signal loss observed in root hairs, where the

strong signal of the wild type plant was almost completely abolished in the double knockout plant

(Fig. 13). The strongest XEH signal in the root however, came from the root tip and was unchanged

between wild type and knockout plants. It should be noted that this activity could come from other

yet undisclosed plant xyloglucanases or created by the sequential action of α‐xylosidases and β‐

glucosidases (Fig. 13). Interestingly, the pH in the apoplast drops during root hair initiation and

growth (Bibikova et al., 1998). At these lower pH values, just below pH 5 is where AtXTH31, Tm‐

NXG1 and expansins are most active (Papers III & IV; (Cosgrove, 2005)).

Figure 13. In situ XEH activity in wild‐type and double knockout lines. a, XEH activity in wild‐type; b, XEH activity in the double knockout line.

In the etiolated hypocotyls, XEH activity was mainly seen in a region just below the cotyledons and

even though it seemed that XEH activity was reduced in the double knockout plants, large variation

in signal makes accurate conclusions hard to draw.

Structure function relationships in XTH gene products

The first structure of a predominantly hydrolytic XEH, Tm‐NXG1, was solved and presented in Paper

III. This structure together with the previously solved 3D‐structure of a strict XET, Ptt‐XET16‐34

(Johansson et al., 2004) allowed, for the first time, molecular insights into the details of

transglycosylation versus hydrolysis in XTH gene products. The structures were similar but two loops

lining the active site cleft were longer in the hydrolytic Tm‐NXG1 (Fig. 9d). Grafting the shorter loops

of Ptt‐XET16‐34 onto the scaffold of Tm‐NXG1 yielded two loop mutants. Disappointingly, only the

Jens Eklöf

45

mutant with the XET loop lining the positive subsites expressed and could be analysed. This chimera,

Tm‐NXG1‐ΔYNIIG, intriguingly showed intermediate characteristics with a 6‐fold reduction in the

hydrolytic rate of the scaffold Tm‐NXG1 and a doubling of the transglycosylating rate under

saturating substrate conditions (Table 2). This loop was therefore concluded to be a major

determinant of transglycosylation versus hydrolysis in XTH gene products, but not the only

determinant. The 3D‐structure of the loop in the chimera was also determined to verify that the

loop resembled the parental loop of Ptt‐XET16‐34 (Paper III).

Activity of plant endo β1,4 glucanases, EG16s

The endo β‐1,4 glucanases, EG16s, represent a new clade of plant GH16 enzymes. The first and only

characterised member of this clade is PtEG16‐1 from black cottonwood, Populus trichocarpa (Paper

VI). The sequence similarities of PtEG16‐1 to both bacterial GH16 lichenases and plant XTH gene

products made it likely that PtEG16‐1 would act on at least one of these polysaccharides. Despite

being assayed against a range of β‐1,3 and β‐1,4 glucans and galactans, the only polysaccharides on

which PtEG16‐1 showed activity was the two β‐1,3;1,4 glucans, barley β glucan (MLG) and lichenan,

and the β‐1,4 glucans, xyloglucan, HEC and CMC (Paper VI).

A homology model of PtEG16‐1 revealed that as many as 6 cysteines might be surface exposed.

Evidence that some of these cysteines are surface exposed comes from the heterologously expressed

and purified PtEG16‐1. In the absence of reducing agents such as TCEP, DTT, β‐mercaptoethanol or

GSH, no activity could be detected suggesting that free cysteines are important for PtEG16‐1 activity.

The production and purification of an EG16 protein from the moss P. patens was published in 2010

(PpXTH32 (Yokoyama et al., 2010); PpEG16‐3 in Fig. 14). No activity could however be demonstrated

for this protein.

The phylogeny of plant GH16 enzymes

The XTH gene products have traditionally been divided into 3‐4 different Groups (Nishitani, 1995;

Campbell and Braam, 1999; Yokoyama et al., 2004). In Paper III, a phylogenetic tree was constructed

that together with structure function studies could pinpoint the XET and XEH activities to different

groups or subgroups of previous trees (Fig. 14; Paper VI). A longer loop in Tm‐NXG1 (an XEH), shown

to be important for the partition of the glycosyl‐enzyme intermediate towards hydrolysis was

restricted to a subset of Group III. This subset was renamed Group III‐A and hypothesised to be XEHs

(Paper III). The hypothesis that Group III‐A members were hydrolytic, based on a single

characterised enzyme has since been further supported by AtXTH31 (Paper IV) a second hydrolytic

Group III‐A member. A third hydrolytic XTH gene product, VaXGH from Vigna angularis (Tabuchi et

al., 2001) was also placed in Group III‐A based on a short peptide sequence (Eklöf and Brumer, 2010),


46

lending strong support to the notion that Group III‐A XTH gene products indeed display mainly XEH

activity. Over 20 members of Groups I/II and III‐B have been characterised an all have shown only

XET activity, devoid of hydrolytic activity on xyloglucan (reviewed in (Eklöf and Brumer, 2010)).

Group III‐A XEHs were also hypothesised to have arisen as a gain of function from a Group III‐B XET

(Paper III). In 2010, Group III‐A was first thought to have appeared in the angiosperms (Eklöf and

Brumer, 2010) making them ca. 140 million years old, but later the same year their appearance was

moved back to ca. 360 MYA due to findings of gymnosperm Group III‐A mRNAs (Paper VI; (Del Bem

and Vincentz, 2010).

Jens Eklöf

47

Figure 14. A rooted tree of a plant GH16 enzymes. A rooted phylogenetic tree showing the relationship between bacterial lichenases, EG16s and the XTH gene products. Node confidence values are given by posterior probabilities multiplied by 100, supplemented with bootstrap values from maximum likelihood calculations at selected nodes (Paper VI). Gymnosperm Group III‐A members are represented with their TIGR TA identifiers.

Evolution of XTH gene products and the shift in substrate specificity

The discovery of the EG16s who share sequence features with both bacterial lichenases and XTH

gene products shed light on the evolution of the XTH gene products (Fig. 15). In Paper III, an

ancestral clade of XTH genes products from Group I clustered closer to a lichenases. With the

inclusion of EG16s in the analysis, the ancestral XTH gene product group was less conclusive and

could be either Group I or Group III‐B (Fig. 14; Paper VI).


48

Figure 15. Proposed evolution of plant GH16 enzymes. A bacterial lichenase gene was transferred to an early plant genome through lateral gene transfer. More than 480 MYA, a loop narrowing the negative subsites in bacterial lichenases was lost in the plant enzyme, enabling binding of branched xyloglucan. The XTH gene product C‐terminus is believed to have appeared before the first of land plants 480 MYA. While land plants maintain large XTH gene families, they only have a few, most often one EG16 gene, indicating several gene duplication events of XTH genes but not of EG16 genes (Paper VI). Also note how the glucan backbone (blue) has changed trajectory from lichenases to XTH gene products.

Combining the phylogenetic analysis with available 3D‐structures of lichenases, XTH gene products

and a homology model of PtEG16‐1 allowed us to postulate a hypothesis of the evolution from

lichenase to the XTH gene products and how the substrate specificity changed (Fig. 15). A bacterial

lichenase gene probably transferred into the nucleus of an early plant through lateral gene transfer.

Before plants colonised land, the last common ancestor of XTH gene products and EG16s had a loop

truncation event, which rendered a wider negative subsite cleft. This loop truncation was most likely

instrumental in the evolution towards accepting the branched polysaccharide xyloglucan, which does

not easily fit in the narrow negative subsites of lichenases. This loop in lichenases contributes with

many substrate interactions on MLG. These were lost in the last common ancestor of XTH gene

products and EG16s, with detrimental effects on MLG hydrolysis. The new protein surface unveiled

Jens Eklöf

49

by the loop truncation was most likely not very specific towards xyloglucan, and interactions in the

negative subsites with xyloglucan were probably minor.

The positive subsite interactions were not affected by the loop truncation and both xyloglucan and

MLG would have a cellobiose‐based moiety in the +1 and +2 subsites, conferring at least some

substrate interactions with xyloglucan. In the XTH gene products, the C‐terminal extension with a

conserved Xyl+2’ subsite that increases the rate of transglycosylation more than 500‐fold in the XET,

Ptt‐XET16‐34 (Saura‐Valls et al., 2008) appeared. Weak substrate interactions in the negative subsite

in combination with strong positive subsite interactions were probably the reason for the specific

transglycosylation activity of XTH gene products. Support of the importance of the positive subsites

in both the formation of the glycosyl‐enzyme intermediate and in consequent transglycosylation is

Ptt‐XET16‐34’s inability to cleave XGO‐CNP substrates (Johansson et al., 2004).

As the negative subsites evolved towards xyloglucan specificity in certain Group III XTH gene products

and due to a loop insertion in the positive subsites, the members of Group III‐A gained the ability to

cleave xyloglucan (and XGO‐CNP). Compared to the EG16s, these Group III‐A xyloglucanases were

highly specific (Papers III, IV & VI). In conclusion, the unusual strict transglycosylation activity of XETs

evolved from a hydrolytic EG16‐like enzyme. The EG16‐like enzyme had gained the possibility to

hydrolyse xyloglucan via a loop truncation in the negative subsites. Group III‐A members regained

the ability to hydrolyse xyloglucan by stronger substrate interactions in the negative subsites (Mark

et al., 2009) and by a loop insertion in the positive subsites.

This hypothesis of the change in substrate specificity from MLG to xyloglucan has not yet been

experimentally proven. Still, the hypothesis of how the change of the activity from lichenases to

EG16s, XETs and XEHs finds support in the papers presented or cited in this thesis.


50

Conclusions In this thesis two types of xyloglucanases have been described. The first type, represented by PpXG5

and the Group III‐A XTH gene products (Tm‐NXG1 and AtXTH31), are highly specific xyloglucanases

that can harness the xyloside substituent of xyloglucan to improve catalysis. The second type,

represented by PpXG44 and BlXG12, can be seen as xyloglucan tolerant where the xylosyl units, at

least in the negative subsites, are detrimental to catalysis. The specificity is higher for unsubstituted

glucans that due to solubility issues cannot be assayed. Since polymeric xyloglucan is the best

substrate under the conditions tested, they are included under the umbrella of xyloglucanases.

A proposed route for the structural and functional evolution of the strict plant xyloglucan endo‐

transglycosylases from bacterial lichenases via the GH16 endo‐glucanases (EG16s) has also been

described. The first structure of a GH16 XEH, Tm‐NXG1, allowed first glimpse into the molecular

mechanism of transglycosylation versus hydrolysis within XTH gene products. Through rational

design, the XEH activity could be pinpointed to a small subclade of XTH gene products and the

hypothesised XEH activity of Group III‐A was confirmed by the characterisation AtXTH31. While XETs

evolved from a hydrolytic EG16 by the addition of a long C‐terminal insertion, extending the positive

subsites, XEHs in Group III‐A evolved from a XET by a loop insertion in the positive subsites.

Localisation of the XEH activity in situ in Arabidopsis, and the comparison of wild‐type and plant lines

lacking Group III‐A XTH gene products (AtXTH31 and AtXTH32 genes), revealed that Group III‐A

enzymes are the major contributors to XEH activity in several Arabidopsis tissues.

Jens Eklöf

51

Acknowledgements After 10 years, I’m finally getting ready to graduate from KTH. It is not only with joy that I conclude

my time here with my doctoral thesis. After this long time, the school has almost become a part of

me, although that does not mean that I’m longing for KTH‐jumpsuit parties...

Of these 10 years roughly 5 have been spent at the Division of Glycoscience, formerly known as

Wood biotechnology. When I first arrived as a diploma student I was greeted by Prof. Tuula Teeri but

I started my work with Prof. Harry Brumer and under supervision of Dr. Martin Baumann. Somewhat

later, I got accepted as a PhD student with Harry and had the wealth of working with Martin for a

little while longer. Harry, I’m truly grateful for my time here and having had the fortune of working

with you. Martin, I think I owe you a great deal and I am happy to call you my friend.

However, I didn’t just hung around Harry and Martin during my time here. Many are those, whom I

would like to thank for making my time here at KTH a pleasant one. Firstly, I would like to

acknowledge past and present members in Harry’s group: Fredrika, a scientist of great potential who

decided to focus here enormous energy on things that actually contribute to society by starting up

Simris Alg AB; Johan, a great travel companion in tropical heat in both Japan and Oslo ; Nomchit, we

were a dynamic duo on the AtXTH31 project!; Farid, a great chemist who have a large part in most

publications coming from our group (See you in St Petersburg soon); Oliver, a very efficient postdoc

who I really enjoyed working and testing the local pubs with; Niklas, stop fiddling with your fancy

equipment and do some real research with us instead; and all the rest of you, Lage, Gustav, Maria,

Sassa, Ana Catarina, Chunlin, the Stefans, Yang and Magnus.

I’ve also had a great time with the people sitting in my room. Cortwa, Erik, Felicia, Johanna and

Gustav, you’re most likely what I’ll miss most when I leave KTH. Hopefully, I’ll still get invited to after

work beer whenever Ela or Nuria decides to organise it. Mohammad and Marcus, thanks for all the

ice‐cold fishing/hunting trips we made and I hope the anchor has come to good use. For those who

I’ve worked and published withm, whose papers did not make it in to this thesis, Thank you!

There are also less obvious acknowledgements to be made. For a couple of years now I’ve gone to

Munich with some friends around the end of September. I especially remember the weekend we

spent there when I had just broken my leg. So thank you Gustav, Niklas, Ted, Johan, Olle and let’s

not forget our host Alex for truly memorable days.

Finally I would like to thank you Lydia for taking care of our home for these last couple of intense

months. I fear though, that I will have ample opportunities to repay you in the near future.


52

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Appendix

Original articles (Appendices I‐VI)

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