University of Groningen Characterization of 4,6-α ... · glucanotransferase enzymes and their...

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University of Groningen Characterization of 4,6-α-glucanotransferase enzymes and their functional role in Lactobacillus reuteri Bai, Yuxiang IMPORTANT NOTE: You are advised to consult the publisher's version (publisher's PDF) if you wish to cite from it. Please check the document version below. Document Version Publisher's PDF, also known as Version of record Publication date: 2016 Link to publication in University of Groningen/UMCG research database Citation for published version (APA): Bai, Y. (2016). Characterization of 4,6--glucanotransferase enzymes and their functional role in Lactobacillus reuteri. [Groningen]: University of Groningen. Copyright Other than for strictly personal use, it is not permitted to download or to forward/distribute the text or part of it without the consent of the author(s) and/or copyright holder(s), unless the work is under an open content license (like Creative Commons). Take-down policy If you believe that this document breaches copyright please contact us providing details, and we will remove access to the work immediately and investigate your claim. Downloaded from the University of Groningen/UMCG research database (Pure): http://www.rug.nl/research/portal. For technical reasons the number of authors shown on this cover page is limited to 10 maximum. Download date: 20-01-2020

Transcript of University of Groningen Characterization of 4,6-α ... · glucanotransferase enzymes and their...

Page 1: University of Groningen Characterization of 4,6-α ... · glucanotransferase enzymes and their functional role in Lactobacillus reuteri. PhD thesis . to obtain the degree of PhD at

University of Groningen

Characterization of 4,6-α-glucanotransferase enzymes and their functional role inLactobacillus reuteriBai, Yuxiang

IMPORTANT NOTE: You are advised to consult the publisher's version (publisher's PDF) if you wish to cite fromit. Please check the document version below.

Document VersionPublisher's PDF, also known as Version of record

Publication date:2016

Link to publication in University of Groningen/UMCG research database

Citation for published version (APA):Bai, Y. (2016). Characterization of 4,6--glucanotransferase enzymes and their functional role inLactobacillus reuteri. [Groningen]: University of Groningen.

CopyrightOther than for strictly personal use, it is not permitted to download or to forward/distribute the text or part of it without the consent of theauthor(s) and/or copyright holder(s), unless the work is under an open content license (like Creative Commons).

Take-down policyIf you believe that this document breaches copyright please contact us providing details, and we will remove access to the work immediatelyand investigate your claim.

Downloaded from the University of Groningen/UMCG research database (Pure): http://www.rug.nl/research/portal. For technical reasons thenumber of authors shown on this cover page is limited to 10 maximum.

Download date: 20-01-2020

Page 2: University of Groningen Characterization of 4,6-α ... · glucanotransferase enzymes and their functional role in Lactobacillus reuteri. PhD thesis . to obtain the degree of PhD at

Characterization of 4,6-α-glucanotransferase enzymes

and their functional role in Lactobacillus reuteri

Yuxiang Bai

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Cover design Tjaard Pijning

Printed by Ipskamp Drukkers, Enschede

ISBN printed: 978-90-367-8706-2

ISBN digital: 978-90-367-8705-5

The work described in this thesis was carried out in the Microbial Physiology Group of the Groningen Biomolecular Sciences and Biotechnology Institute (GBB) in the University of Groningen and was financially supported by the China Scholarship Council, the University of Groningen, and by the TKI Agri&Food program as coordinated by the Carbohydrate Competence Center (CCC-ABC; www.cccresearch.nl).

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Characterization of 4,6-α- glucanotransferase enzymes and their

functional role in Lactobacillus reuteri

PhD thesis

to obtain the degree of PhD at the University of Groningen on the authority of the

Rector Magnificus Prof. E. Sterken and in accordance with

the decision by the College of Deans.

This thesis will be defended in public on

Friday 18 March 2016 at 12.45 hours

by

Yuxiang Bai

born on 27 September 1986 in Jiangsu, China

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Supervisor Prof. dr. L. Dijkhuizen Co-supervisor Dr. R.M. van der Kaaij Assessment Committee Prof. T. Desmet Prof. D.B. Janssen Prof. M.J.E.C. van der Maarel

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Contents

CHAPTER 1 7-44

4,6-ΑLPHA-GLUCANOTRANSFERASES, STARCH-ACTING ENZYMES FROM LACTIC ACID BACTERIA

CHAPTER 2 45-66

CHARACTERIZATION OF THE 4,6-ΑLPHA-GLUCANOTRANSFERASE GTFB ENZYME OF LACTOBACILLUS REUTERI 121 ISOLATED FROM INCLUSION BODIES

CHAPTER 3 67-94

BIOCHEMICAL CHARACTERIZATION OF LACTOBACILLUS REUTERI GLYCOSIDE HYDROLASE FAMILY 70 GTFB TYPE OF 4,6-ΑLPHA-GLUCANOTRANSFERASE ENZYMES THAT SYNTHESIZE SOLUBLE DIETARY STARCH FIBERS

CHAPTER 4 95-128

LACTOBACILLUS REUTERI STRAINS CONVERT STARCH AND MALTODEXTRINS INTO HOMO-EXOPOLYSACCHARIDES USING A CELL-ASSOCIATED 4,6-ΑLPHA-GLUCANOTRANSFERASE ENZYME

CHAPTER 5 129-156

STRUCTURAL BASIS OF THE ROLE OF STARCH AND SUCROSE IN HOMO-EXOPOLYSACCHARIDE FORMATION IN LACTOBACILLUS REUTERI 35-5

CHAPTER 6 157-204 CRYSTAL STRUCTURE OF THE 4,6-ΑLPHA-GLUCANOTRANSFERASE GTFB: EVIDENCE THAT DIETARY CHANGES TRIGGERED THE EVOLUTION OF GLUCANSUCRASES FROM ΑLPHA-AMYLASES

CHAPTER 7 205-210

SUMMARY AND THESIS DISCUSSION

SAMENVATTING EN DISCUSSIE 211-216

ACKNOWLEDGEMENTS 217-222

CURRICULUM VITAE 223-228

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Chapter 1

4,6-α-Glucanotransferases, starch-acting enzymes from lactic

acid bacteria

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Introduction Starch, which exists as storage carbohydrate in seeds, roots, tubers and plants, is a major dietary carbohydrate in our life (1). It consists of α-glucan with (α1→4)-linked backbone chains and α1→6 branching points, in the form of amylose and amylopectin. Amylose is a roughly linear polysaccharide containing approximately 99% α1→4 and less than 1% α1→6 glycosidic linkages, while amylopectin has approximately 95% α1→4 and 5% α1→6 linkages (2). According to the distribution of amylose and amylopectin, starches are divided into three groups: waxy starch (<15% amylose), normal starch (16-35% amylose), and high-amylose starch (>35% amylose) (3).

Apart from the raw starches, starch derivatives produced by physical or chemical treatments are now widely used (4). For instance, pre-gelatinization improves the cold water dispersibility of starch, and thus can be applied in preparation of instant convenience food (5). Acid hydrolyzed starches have lower molecular mass and therefore lower viscosity; they are applied in food coatings and batter (4). Etherification and esterification can markedly alter the physiochemical properties of starches by introducing functional groups (6-9).

In addition, many starch acting enzymes that are active under industrial conditions such as high viscosity and high temperature have been commerciallized and are now in use in a number of industrial processes (10, 11). This biological modification of starch draws a lot of attention from industry. Thermostable α-amylase enzymes of the glycoside hydrolase (GH) family 13 (EC 3.2.1.1) for instance are used for starch liquefaction in the syrup producing industry. They are also widely applied in the brewing industry to make products that partially replace the expensive malt, in the baking industry to improve the rheological properties of dough, in the paper industry to produce modified starch, and in the textile industry to remove starch (12-14). Glucoamylase (EC 3.2.1.3) also plays a vital role in the syrup producing industry, in the saccharification step following the liquefaction process (Fig. 1) (15). Moreover, isoamylase (EC 3.2.1.68) and pullulanase type I and II (EC 3.2.1.41), which hydrolyze α1→6 branching bonds within starch, constitute another important group classified as debranching enzymes (Fig. 1) (16-18).

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Apart from these hydrolyzing enzymes, several glucanotransferases have also found industrial applications in starch conversion. These enzymes cleave α1→4 glycosidic bond of the donor molecule and transfer the cleaved glucose residue or aglycon segment to an acceptor molecule, forming a new glycosidic bond. Cyclodextrin glucanotransferase (CGTase, EC 2.4.1.19) converts starch into cyclodextrins normally consisting of 6, 7, or 8 glucose residues (Fig. 1) (19). Because of the unique physiochemical properties of their rigid barrel structure, cyclodextrins are widely applied in the food, textile, cosmetic and pharmaceutical industry to improve the solubility and stability of guest molecules by partial or entire encapsulation (20, 21). 4-α-Glucan branching enzyme (EC 2.4.1.18) hydrolyzes an α1→4 bond within a chain and connects the non-reducing end to an α1→4-linked chain forming a branched α1→6 bond (Fig. 1). It plays an important role in amylopectin synthesis in plants along with starch synthases (22). 4-α-Glucanotransferase (4-α-GTase, EC 2.4.1.25) catalyzes the intermolecular disproportionation reaction by transferring a segment of α1→4 glucan to the non-reducing end of the acceptor α-glucan, forming a new α1→4 bond (Fig. 1). Following 4-α-GTase treatment, the modified starch is capable of forming gels with thermoreversible gelling property (23). Dextran dextrinase (DDase, EC 2.4.1.2) from Gluconobacter oxydans catalyzes the transfer of an α1→4 linked glucose unit from the non-reducing end of a donor to the non-reducing end of an acceptor molecule forming an α1→6 linkage. It is regarded as an interesting alternative for industrial dextran production from maltodextrins instead of sucrose (24). However, DDase cannot process unhydrolyzed starch (25).

4,6-α-Glucanotransferase enzymes (4,6-α-GTases, EC 2.4.1.B34 http://www.brenda-enzymes.org/) recently have been characterized (26). They predominately cleave an α1→4 bond from the non-reducing end of starch or maltodextrins and transfer the cleaved glucose unit to the non-reducing end of the acceptor mostly by formation of an α1→6 bond and occasionally an α1→4 bond, forming isomalto-/malto- oligosaccharides (IMMO) and polysaccharides (IMMP) (Fig. 1). Unlike DDase, 4,6-α-GTases do not have branching specificity on dextrin substrate and disproportion activity on isomaltooligosaccharides, but are able to process unhydrolyzed starch molecules such as amylose and side chains of amylopectin (Chapter 4), showing promising commercial value for the food industry (27, 28). Using protein blasts in the NCBI database

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(http://www.ncbi.nlm.nih.gov/BLAST/) with the GtfB enzyme as a query, approx. 28 characterized and putative 4,6-α-GTases are retrieved. All of them are exclusively present in lactic acid bacteria. Interestingly, these 4,6-α-GTases do not belong to the well-known starch-acting GH families such as GH13, 57, or 77 but constitute a GH70 subfamily; family GH70 mainly consists of sucrose-acting glucansucrases (GS) (29-31).

Fig. 1. Schematic representation of the reactions of several starch-acting enzymes including α-amylase, glucoamylase, isoamylase/pullulanase, cyclodextrin glucanotransferase, branching enzyme, 4-α-glucanotransferase and 4,6-α-glucanotransferase.

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Starch-acting enzymes in lactic acid bacteria (LAB) Fermentation by lactic acid bacteria (LAB) is a popular approach in food processing to increase the nutritional value, and to improve the taste and flavor of food. LAB strains are widely distributed in fermented meals, beverages, sourdoughs, cooked rice, fish products and digestive tract of animals as well as plants (32-37). Some of the LAB strains, especially of the genus lactobacilli, are able to degrade and utilize starch as carbon- and energy source during fermentation, using a dedicated set of enzymes acting on starch (38). Several of these enzymes have been expressed heterologously and studied (3). By biochemical characterization of these enzymes, amylopullulanase, pullulanase and α-amylase were observed intracellularly and/or extracellularly in Lactobacilli such as Lactobacillus amylophilus, Lactobacillus acidophilus, Lactobacillus cellobiosus, Lactobacillus plantarum, Leuconostoc lactis ssp. lactis and Streptococcus bovis (39-44). These known LAB enzymes all catalyze starch-hydrolyzing reactions by cleaving the α1→4 bonds and/or α1→6 branching points.

Some LAB are known to produce extracellular polysaccharides (EPS) including homopolysaccharides (composed of glucose or fructose units, respectively) and heteropolysaccharides (composed of mainly glucose, fructose, rhamnose and some charged groups) (45-47). Genes involved in heteropolysaccharide synthesis are located either on plasmids or on the chromosome, and are organized in clusters. They carry genes involved in regulation, polymerization and chain length determination, biosynthesis of the repeating unit, polymerization and export (48, 49). Homopolysaccharides, such as β-fructans and α-glucans, are synthesized from sucrose by individual enzymes such as fructansucrases including inulosucrase (EC 2.4.1.9), levansucrase (EC 2.4.1.10), and GS including dextransucrase (EC 2.4.1.5) and alternansucrase (EC 2.4.1.140) (50-52).

These GS convert sucrose into α-glucans with α1→2, α1→3, α1→4 or/and α1→6 glucosidic bonds and constitute family GH70 (47, 50, 52). In nature, GS are extracellular enzymes contributing to the EPS formation in host strains (53, 54). Interestingly, all reported GS have been exclusively isolated from LAB strains including Streptococcus, Leuconostoc, Weissella, and Lactobacillus (29, 55-57). The homo-exopolysaccharides (homo-EPS) synthesized by GS in LAB such as Streptococcus mutans play a clear role in occurrence of dental caries; GS three-

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dimensional structures therefore are studied in order to design structure-based inhibitors for anti-caries drugs (55, 58).

Like GS, 4,6-α-GTases are also exclusively found in LAB strains. According to the primary structure of 4,6-α-GTases, they are classified into family GH70. The first 4,6-α-GTase representative is GtfB from Lactobacillus reuteri 121; it is also the first starch-acting glucanotransferase characterized from LAB strains. Besides gtfB, L. reuteri 121 also harbors genes encoding levansucrase, inulosucrase and reuteransucrase (29, 53, 59-61). Recently, more 4,6-α-GTases, such as GtfW from L. reuteri DSM20016 and GtfML4 from L. reuteri ML1, were characterized, thus expanding this subfamily of GH70 (28).

GH70 and clan GH-H GH13, GH70 and GH77 constitute clan GH-H, also called the α-amylase superfamily, according to their protein sequences and the basic three-dimensional folds of their catalytic modules, rather than on their specificity of action (62-64). Of these families constituting clan GH-H, family GH13, including hydrolases, transglycosidases and isomerases, with around 30 known specificities, is most complex (www.cazy.org) (65, 66). Family GH77 is the least complex, all 25 characterized members are 4-α-glucanotransferases (4-α-GTases, amylomaltases). They are of similar specificity, cleaving an α1→4 bond and transfering a segment of the α1→4 linked chain to the non-reducing end of an acceptor substrate forming a new α1→4 bond (http://www.cazy.org/). (67-70). All members of clan GH-H harbor a basic (β/α)8-barrel, a TIM barrel, in the catalytic domain. The barrel consists of eight β-strands and eight α-helices that are alternately arranged (62). The C-terminal ends of β-strands join to the N-terminal ends of the following α-helices by irregular loops that also link the C-terminal ends of α-helices and the N-terminal ends of β-strands (Fig. 2A). In addition, the enzymes in clan GH-H catalyze reactions by a common mechanism that involves a nucleophilic aspartate residue at the C-terminal end of β-strand 4, an acid/base catalyst glutamic acid residue at the end of β-strand 5, and a putative transition state stabilizer aspartate residue located a few residues beyond the C-terminal end of β-strand 7.

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For GH13 enzymes, loop 3 connecting the β-strand 3 to the helix 3 of the barrel is generally long compared to other loops, and appears to fold as an independent unit that even can be considered as a domain. In some cases, such as the branching enzymes, loop 3 is shorter and does not have the characteristic of a domain. Some of the GH13 members are more complex and have extra domains, e.g. an additional N-terminal domain in isoamylase (71) or domain D and E in CGTase (Fig. 3) (19, 72, 73). For CGTase, domain E is confirmed as starch binding domain; domain D is exclusively found in CGTases but its function has remained unclear (72).

Unlike GH13 enzymes, domain C is missing from GH77 amylomaltases. Another remarkable characteristic of amylomaltase structures is the presence of subdomain B2 that contains large α-helices (Fig. 3). This subdomain B2 is unique in GH77 enzymes. It only partially overlaps with the N-terminal domain of isoamylase (67). Additionally, the active-site cleft of amylomaltase is different from the open cleft of GH13 members as it is partially covered by a long extended loop (250s loop formed by residues 247-255 in subdomain B1 for Thermus aquaticus) (67). This loop is flexible, which may contribute to substrate binding and prevent the formation of small cyclic products because of the steric hindrance to the active site.

For GH70, three-dimensional structures of several GS have been elucidated, including Gtf180-∆N from L. reuteri 180 (PDB: 3KLK), GtfA-∆N from L. reuteri 121 (PDB: 4AMC), Gtf-SI from S. mutans (PDB: 3AIE) and ∆N123-GBD-CD2 of the (1→2) branching glucansucrase DSR-E from L. mesenteroides NRRL B-1299 (PDB: 3TTQ) (55, 74-76). The domain organization is similar to that of GH13 enzymes as the catalytic cores of all GS consist of domains A, B, and C. However, GS also contain two extra domains IV, V, and a variable N-terminus (Fig. 3). Interestingly, domains A, B, IV and V are all built up by two discontinuous segments of polypeptide chains. Only domain C is composed of a contiguous polypeptide. The polypeptide chain of the full glucansucrase follows an order of V, IV, B, A, C, A, B, IV, and V from the N- to C-terminus. Besides the domain organization, the characteristic (β/α)8-barrel of clan GH-H in GH70 is also different from that of GH13 and GH77. A circular permutation of the (β/α)8-barrel predicted by the primary structure was confirmed by the 3D structure

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analysis (Fig. 2B). The barrel is initialized with the α-helix that is equivalent to the α-helix 3 of the GH13 or GH77 enzymes and ends with the β-strand equivalent to β-strand 3 of the GH13 or GH77 enzymes, following an order as shown in Fig. 2B.

Fig. 2. Schematic diagram of the (A) normal barrel of GH13 and GH77 and (B) the permuted barrel of family GH70; gray arrows represent β-strands and blue cylinders represent α-helices. The order of helices is numbered. For GH70 proteins, this order is 34567812.

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Fig. 3. Domain arrangements of clan GH-H proteins (A) human pancreatic α-amylase (PDB: 1HNY), (B) Bacillus circulans cyclodextrin glucanotransferase (CGTase) (PDB: 1CGT) of GH13, (C) Lactobacillus reuteri glucansucrase (PDB: 3KLK) of GH70 and (D) Thermus aquaticus amylomaltase (4-α-glucanotransferase) of GH77 (PDB: 1CWY).

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4,6-α-Glucanotransferase and GH70 Family GH70 mainly consists of GS that are classified as dextransucrases, mutansucrases, alternansucrases, reuteransucrases, and branching dextransucrases, according to the linkage specificity of the products formed. Although the GH70 GS are structurally closely related to the members of GH13 and GH77, their substrate specificity is completely different. These GS enzymes are inactive on starches but active on sucrose, synthesizing oligo- and/or polysaccharides.

The occurrence of 4,6-α-GTases in family GH70 implies that all families of the GH-H clan in fact possess catalytic activity on starches and maltodextrins. This also provides insights into the evolutionary pathway between families GH13 and GH70. A phylogenetic tree based on the alignment of the catalytic core amino acid sequences of GS (http://www.cazy.org) and 4,6-α-GTases is shown in Fig. 4. Putative 4,6-α-GTases such as Gtf106B were identified in the non-redundant protein sequence database by a protein blast search (http://www.ncbi.nlm.nih.gov). The maximum-likelihood tree shows that the 4,6-α-GTases are more closely related to the GS from L. reuteri compared to those from Leuconostoc and Streptococcus. However, unlike GS, 4,6-α-GTases are completely inactive on sucrose, with the exception of Gtf106B that has faint activity which is detectable after 27 h incubation (77).

These GS from different species clearly cluster within their host genera of Streptococcus, Lactobacillus and Leuconostoc (78-84). Along with the time of occurrence and the order of specific events, these clusters may indicate that the gtf genes from Streptococcus were acquired from other genera like Lactobacillus via horizontal gene transfer (85). By the acquisition of Gtfs, Streptococcus mutans became capable to form EPS, resulting in the formation of cariogenic dental biofilms (86). However, the characterized and putative 4,6-α-GTases are only found within the family Lactobacillaceae including Lactobacillus and Pediococcus, and so far no 4,6-α-GTase gene has been found in Streptococcus, Leuconostoc and other genera of LAB.

Alignment of the conserved sequences of the characterized 4,6-α-GTases, GtfB, GtfML4 and GtfW, with those of typical GS showed not only similarities but also characteristic differences in their primary structures (Fig. 5). The three catalytic

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residues (the nucleophilic aspartate, the acid/base glutamate and the transition state stabilizing aspartate) constituting the catalytic site in GS are also present in 4,6-α-GTases, e. g. D1015, E1053 and D1125 in GtfB of L. reuteri 121 (26, 58). They are in loops following β-strands β4, β5 and β7, in homology regions II, III, and IV of GS respectively (Fig. 2). Mutating D1015 or E1053 in GtfB resulted in loss of activity (26), indicating that 4,6-α-GTases and GS have the same catalytic residues. In view of such similarities it also appears likely that the reactions catalyzed by 4,6-α-GTases and GS share a similar double-displacement mechanism (Fig. 6). The glutamic acid residue at β-strand 5 donates a proton to the glycosidic oxygen, resulting in bond cleavage. The cleaved glucosyl unit or aglycon segment leaves the active site forming an intermediate. The glucose residue is stabilized by hydrogen bonding involving the aspartic acid that is located after β-strand 7 and the hydroxyl groups of the glucose. Then a covalent β-bond between C1 of the glucose and the nucleophilic aspartate at β-strand 4 forms in order to further stabilize the intermediate. In the second half of the reaction, the glutamate at β-strand 5 is reprotonated by removing a proton from the acceptor substrate. The covalently bonded glucosyl moiety at the donor subsite is transferred to an acceptor substrate with retention of the α-anomeric configuration forming a new α-bond. Besides the similarities, a large number of amino acid residues conserved in regions I, II, III, and IV of GS are different in the 4,6-α-GTase enzymes, especially the residues in region I, and IV that contribute to the -1, +1, and +2 donor/acceptor substrate binding subsites (Fig. 5). These differences in amino acid residues may contribute to the differences in substrate specificity of 4,6-α-GTases and GS.

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Fig. 4. Unrooted phylogenetic tree of characterized GH70 enzymes and some putative 4,6-α-GTases. Alignments and dendrogram construction were carried out using the catalytic core of all enzymes. Each sequence is labeled with the corresponding bacterial strain and enzyme names. The characterized ones are indicated in red frame.

Fig. 5. Sequence alignment of conserved sequences (Motifs II, III, IV, and I) in the catalytic domains of GS and 4,6-α-glucanotransferases in GH70. The catalytic residues (filled triangles) and several conserved amino acid residues (shielded with red color) in subsites -1, +1, and +2 are indicated.

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Fig. 6. Reaction mechanism of GS and 4,6-α-glucanotransferase enzymes. Asp represents the nucleophilic aspartate residue and Glu represents the general acid/base residue.

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Expression and production of 4,6-α-GTases In order to improve the expression and production of 4,6-α-GTases, diverse methods previously reported for DDases, 4-α-GTases, and GS, which are either specificity- or structurally-related to 4,6-α-GTases, may be applied. According to previous reports, these enzymes mainly were produced in two different ways including whole-cell fermentation and heterologous expression.

In 1950s, Hehre et al. reported an enzyme which is capable of synthesizing dextran from maltoheptaose, or the hydrolysates of amylose, amylopectin and glycogen (87). The enzyme cleaved α1→4 bonds and transferred the glucose units to another (acceptor) chain introducing α1→6 linkages, and was named dextran dextrinase (DDase) (25). The distribution of these enzymes thusfar is limited to acetic acid bacteria such as Acetobacter capsulatum, Gluconobacter oxydans (24, 88-93). Unfortunately, to date no corresponding gene and protein sequences have been reported yet. The production of DDases was done in the original strain, i.e. G. oxydans (91, 94). Methods for the improvement of DDase production in G. oxydans by adding polyhydric alcohols such as mannitol, sorbitol and glycerol as substrates have been patented (95). The production of intracellular DDase of G. oxydans ATCC 11894 can also be enhanced by adding various carbon and nitrogen sources, such as glycerol and peptone (91). The yield of DDase in G. oxydans M5 has been improved by optimizing groups of medium constituents using a four-factor five-level central composite design. The optimization resulted in a 17-fold increase of yield of DDase compared to the original conditions (94).

For 4-α-GTases, little research has been done on the production improvement via whole-cell fermentation. Genes encoding 4-α-GTases have been successfully cloned from their original strains; these enzymes were preferably expressed in E.coli or Bacillus subtilis host strains. However, their heterologous expression to some extent is still a challenge. For instance, the Thermococcus litoralis 4-α-GTase tends to aggregate in inclusion bodies when expressed in E. coli as the corresponding gene is rich in AGA and AGG codons encoding arginine. Simultaneous expression of tRNAAGA, tRNAAGG and molecular chaperon GroEL resulted in an increase of both production and solubility of 4-α-GTase (96). Alternatively, codon optimization improved the expression yield of Thermus

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thermophilus 4-α-GTase in E. coli 100-fold (97). In addition, the Thermus scotoductus 4-α-GTase gene has been expressed successfullyin B. subtilis, and is produced extracellularly. To improve the expression, a dual promotor system was used, resulting in an increase of the yield by approx. 12-fold compared to the single promotor system (98).

For GS which are structurally related to 4,6-α-GTases, both whole-cell fermentation in their original host strain and heterologous expression in E. coli have been reported. In the whole cell fermentation, approaches including central composite design, response surface methodology, fed batch fermentation, with pH control, are used to enhance the production of dextransucrase in Leuconostoc strains (99-105). However, over the years more and more GS genes have been cloned and expressed in E. coli (106-109). In recent years, such studies mainly focussed on improving the heterologous expression level of GS. By dropping the induction temperature from 37 °C to 15°C, the yield of dextransucrase LcDS from Leuconostoc citreum HJ-P4 in E. coli sharply increased by 330-fold (108). When changing the E. coli host strain/inducer system for the expression of dextransucrase DexT from Lc. citreum KM20 its activity was enhanced 12 fold under optimized conditions including induction OD600 and concentration of inducer (110). Also, addition of 0.005% (w/v) calcium ions in the fermentation medium increased production of dextransucrase of Lc. mesenteroides PCSIR-4 by 2.5-fold (111). Also deletion of peptide segments from full-length GS has been applied, at either the N- or C-terminus, without significantly influencing its enzymatic activity. For example, the alternansucrase from Leuconostoc mesenteroides NRRL B-1355 (2057 amino acids) in E. coli is poorly soluble, instable, and thus hard to express. By truncation of the C-terminal APY repeats but retaining four CW-like repeats, the protein became more soluble and less degradable than the full-length alternansucrase (112). For the GtfR from Sterptococcus oralis, a deletion of 261 amino acids in N-terminus resulted in a 50-fold increase of yield compared to the original glucanotransferase (113). Moreover, partial gene deletion has been widely used in recombinant expression of other GS, such as Gtf180 from L. reuteri 180, GtfA from L. reuteri 121, and DSRS from Lc. mesenteroides NRRL B-512F (29, 74, 114, 115).

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4,6-α-GTases, such as GtfB, GtfW and GtfML4, have been heterologously expressed in E. coli (26, 28). But the soluble protein expression levels are disappointingly low. High expression levels of GtfB resulted in accumulation (of at least partly active enzyme) in inclusion bodies (Chapter 2) (116). To extract the functional GtfB protein from inclusion bodies, conventional denaturing and refolding steps resulted in a recovery of 65% of the hydrolytic activity of GtfB compared to the soluble enzyme. Also the (non-classical) inclusion bodies (ncIBs) themselves were used as an alternative approach for preparation of active enzyme. The ncIB GtfB only had approximately 10% of the hydrolytic activity of soluble GtfB, but showed a much higher thermostability than soluble GtfB. In view of the high yield and the low preparation cost, as well as the structural stability, ncIB GtfB protein potentially may be used in industrial applications. In the next step the soluble expression of active GtfB enzyme was considerably improved by gene deletion of its N-terminally variable region without influencing the enzymatic activity compared to the full-length version. Truncation of 733 amino acids in the N-terminous resulted in a yield of more than 40 mg pure GtfB-∆N protein from one-liter E. coli culture, which is 75-fold higher than that of full-length GtfB based on molar calculation (Chapter 3) (117).

Assays for measurement of 4,6-α-GTase activity Proper activity assays for 4,6-α-GTase enzymes are essential for their biochemical characterization. In view of the complexity of the 4,6-α-GTase catalyzed hydrolysis and transglycosylation reactions, proper, specific and accurate assays are difficult to establish. However, the available assays for DDases and 4-α-GTases that catalyze somewhat similar reactions as 4,6-α-GTases provided leads, also because they all transfer glucose moieties from the non-reducing end of the donor substrate to the non-reducing end of the acceptor substrate.

The dinitrosalicylic acid (DNS) assay for reducing sugars, conventionally used for glucose measurement, has been adopted for estimating the activity of DDases. However, the poor sensitivity of DNS and interference by unreacted maltodextrins in the reaction mixture hindered its application (118). As an alternative, a viscosity build-up assay was tested based on the change of rheological properties from maltodextrins to dextran. But it was also hampered by

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the complex non-Newtonian and time-dependent flow behavior of the maltodextrin/dextran mixture (118). Another assay, which uses maltose as a standard substrate, based on discrete transglycosylation reactions was successfully applied (119). Generated panose [α-D-glucose-(1,6)-α-glucose-(1,4)-α-glucose] was demonstrated to be a valid indicator for transglycosylation activity. An assay mimicking maltose utilization but based on p-nitrophenyl-α-D-glucopyranoside (NPG) was also reliable (118). Use of NPG is more preferable than the maltose assay because it measures both transglycosylation activity and hydrolysis activity.

Several assays have been developed for characterization of 4-α-GTases. For determining the hydrolysis activity, maltotriose was used as substrate and the released glucose was measured by the glucose oxidase enzyme method. The rate of glucose release was then defined as the hydrolysis activity (120, 121). The disproportionation activity was estimated by measuring the intensity of iodine staining of non-denaturing polyacrylamide electrophoresis gels containing glycogen (122). Lugol’s method was also adapted by monitoring the iodine-staining capacity of products during the conversion of amylose in the presence of maltose as acceptor (123). The iodine-staining capacity can be directly determined by measuring the optical density change (124). However, because 4-α-GTases cleave α1→4 bonds and form new α1→4 bonds, the iodine staining method is less quantitative (120).

The maltose assay has been used for determining the kinetic parameters of 4,6-α-GTase GtfW from L. reuteri DSM20016 (28). However, this assay has significant limitations with other 4,6-α-GTases such as GtfB which has detectable activity on longer maltooligosaccharides but has faint or virtually undetectable activity on maltose. To solve this problem, a combination of assays was developed using amylose as substrate and measuring both iodine staining and D-glucose (GOPOD) release to determine both transglycosylation and hydrolysis activities (Chapter 3) (117). The iodine staining assay determines the total activity of 4,6-α-GTases and is based on the principle that iodine can be trapped by linear (1→4)-α-D-glucans (i.e. amylose) but not by linear (1→6)-α-D-glucans (125, 126). Hydrolysis activity results in release of glucose from the amylose substrate, which also impairs the complex capacity of amylose with iodine atoms. The decrease of the

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complex capacity of amylose thus reflects total activity. Hydrolysis activity of 4,6-α-GTases is measured with the D-glucose assay kit (GOPOD). The transglycosylation activity was subsequently calculated by subtracting the hydrolysis activity from total activity (Fig. 7). These assays with amylose as substrate thus allow direct determination of total and hydrolysis activities, and calculation of the transglycosylation activities of 4,6-α-GTases.

Fig 7. Schematic diagrams of the complexes of iodine with amylose and with the amylose derived products of the 4,6-α-GTase activities. Iodine ions are shown as (short chains of) blue circles.

Biological role of 4,6-α-GTases in LAB As described above, GS usually contribute to homo-EPS formation in their host LAB strains. These secreted GS enzymes are anchored to the cell wall, producing EPS in the presence of sucrose. Depending on the GS, the resulting EPS can be divided into dextran that mainly consists of α1→6 linked glucose units, mutan with mostly α1→3 linkages, alternan with strictly alternating α1→6 and α1→3 linkages, and reuteran with α1→4 and α1→6 linkages. The linkage specificities confer different physiochemical properties to the EPS, e.g. mutan from S. mutans

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contains a relatively high percentage of α1→3 linkages and is water-insoluble, while dextran is rich in α1→6 linkages and is water-soluble. The insoluble mutan may contribute to the formation of dental caries because it adheres to teeth and traps oral bacteria, food debris and salivary components (127-129). EPS is also a major fraction of the extracellular polymeric substances that form the scaffold for the three-dimensional architecture of the biofilm (130). The EPS in extracellular polymeric substances may contribute to other cellular functions (Table 1) in bacterial biofilms.

Table 1. Roles ascribed to exopolysaccharides in biofilms (130, 131).

Biofilm Function Functional Relevance of EPS to Biofilms Adhesion providing the initial steps in the colonization of surfaces

Bacterial cell aggregation

establishing bridges between cells to temporarily immobilize bacterial cell populations and developing high cell densities, and cell–cell recognition

Retention of water maintaining a hydrated microenvironment around biofilms, leading to the survival of cells in water-deficient environments

Cohesion of biofilms forming a hydrated polymer network to mediate the mechanical stability of biofilms and determine biofilm architecture, as well as allowing cell-cell communication

Nutrient source serving as source of carbon containing compounds

Protective barrier conferring resistance to non-specific and specific hosts during infection, confer tolerance to various antimicrobial agents

Sorption of organic compounds and inorganic ions

mediating the accumulation of nutrients from the environment, sorption of xenobiotics and recalcitrant materials, promote polysaccharide gel formation

resulting in ion exchange, mineral formation and the accumulation of toxic metal ions

Enzyme binding stabilizing and accumulating extracellular enzymes Sink for excess energy storing excess carbon under unbalanced carbon to nitrogen ratios

The strains carrying GS encoding genes are typically identified by the appearance of slimy colonies on solid media or viscous solutions in liquid media when grown on sucrose as carbon source (132, 133). Gene inactivation studies also revealed that the slimy colony morphology resulted from GS activity in wild-type LAB strains (132, 133). Besides GS, also the presence of fructansucrases producing β-fructans from sucrose results in the formation of slimy colonies. GS and fructansucrase enzymes can be further distinghuised using raffinose (α-D-galactopyranosyl-(1→6)-α-D-glucopyranosyl-(1→2)-β-D-fructofuranoside) as a carbon source: fructansucrase enzymes are able to utilize raffinose to produce EPS, but GS are not (54, 134). Using this method, many α-glucan producing bacterial strains have been identified from various sources, such as fruit and vegetables, sourdoughs, dairy, mammal intestine, etc (135-137).

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Insights in the biological roles of 4,6-α-GTases in LAB strains were obtained when growing L. reuteri 121 strain 35-5 (that lacks fructan formation and carries both the gtfA and the gtfB genes) on agar plates with maltodextrins (5% w/v, dextrose equivalent 13-17) as carbon source (Chapter 4). Surprisingly, slimy colony morphology (Fig. 8A) was observed (Chapter 6). The structure of the EPS extracted by ethanol precipitation was analyzed by NMR (Fig. 8B), showing that the percentage of α1→6 linkages increased dramatically compared to the substrate maltodextrins. GtfB has been expressed in E. coli and characterized as an enzyme converting maltodextrins into isomalto/malto- oligosaccharides and polysaccharides, with α1→4 and α1→6 linkages (Chapter 4) (26, 27). Moreover, a periodic acid-Schiff (PAS) staining experiment confirmed that the GtfB enzyme contributes to the EPS formation in situ (Chapters 4 and 5). Therefore, GtfB enzyme contributes to the homo-EPS formation in L. reuteri 35-5 using starches or maltodextrins as substrates.

Fig. 8. (A) Colonies of L. reuteri 121 strain 35-5 grown on MRS agar containing 5% glucose or maltodextrins (dextrose equivalent 13-17). Slime formation indicates the production of EPS from maltodextrins but not from glucose. (B) NMR spectra of the substrate maltodextrins and the corresponding EPS from L. reuteri 35-5.

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Application of 4,6-α-GTases 4,6-α-GTases raise the interest of the starch industry because of their capacity of converting starch or maltodextrins into isomalto/malto-polysaccharides (IMMP) with high percentages of α1→6 linkages (138, 139). These modified starch products are novel soluble dietary fibers. Dietary fibers positively contribute to the human health. Based on the physiochemical properties, they are divided into insoluble dietary fibers such as resistant starch, wheat bran, hemicelluloses etc., and soluble dietary fibers such as inulin, pectin, gums, and polydextrose (140). In food applications, the addition of small quantities of dietary fibers may significantly affect the texture, taste and mouth feel of the food products. Based on their physiochemical properties and health effects, these dietary fibers are applied in various food products, e.g. the soluble dietary fibers can be added in juice, viscous liquid and gel products. Some soluble dietary fibers also have many health benefits: reduction of the cholesterol level and thus lowering the risk of heart diseases as the soluble fiber attaches to the cholesterol particles, removing these from the body (141, 142); control of sugar absorption, providing protection against diabetes (143); assisting in keeping the body weight (144); stimulating healthy bowel movements (145). However, the production of soluble dietary fibers is costly because most soluble dietary fibers currently are extracted from foods such as oatmeal, nuts, beans, blueberries and apples (146). Therefore, 4,6-α-GTases producing soluble dietary fiber from cheap starches are of great commercial potential (139).

Of all the 4,6-α-GTases, GtfB from L. reuteri 121 has been studied in most detail using various starches as substrates. The IMMP products consist of linear α1→6-linked glucan chains that are attached to the non-reducing ends of starch fragments (Fig. 9). Comparison of thirty different starches and maltodextrins showed that the substrates with long and linear α1→4-linked glucan chains are most efficient for the production of products with a high percentage of α1→6 linkages (138). The percentage of α1→6 linkages even reached up to 92% when using amylose V as substrate. IMMP is a novel polysaccharide different from dextran although both are rich in α1→6 linkages. Firstly, dextrans are large polymers which are synthesized by dextransucrases using sucrose as substrate, while IMMP is produced by 4,6-α-GTase using starch as substrate. Secondly, the sizes of dextrans produced by glucansucrases (>10 MDa) (53) are much larger

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than of IMMP (10-1000 kDa) (Chapter 4) (28). Thirdly, dextran contains branches that are absent in IMMP. The in vitro experimental results revealed that IMMP, or more precisely the IMMP fraction rich in α1→6 linkage, is indigestible in the small intestine and consumed in the large intestine (138). Moreover, preliminary degradation tests using fecal samples from a single adult showed that the addition of IMMP resulted in an increased formation of short fatty acids acetate and propionate, indicating that IMMP may have a positive effect on human health (138). Conclusively, IMMP are novel soluble dietary fibers that may contribute to the health of the human body.

Fig. 9. Schematic diagram of the 4,6-α-GTase-modified starch (amylopectin) produced by 4,6-α-GTases.

These IMMP polysaccharide- and oligosaccharide products may act as prebiotic substrates (138, 139) defined as “a selectively fermented ingredient that allows specific changes, both in the composition and/or activity in the gastrointestinal microflora that confers benefits upon host well-being and health” (147, 148). In the microflora, Lactobacilli and Bifidobacteria are usual target genera for identifying prebiotics. The known prebiotics including inulin, fructo-oligosaccharides (FOS), galacto-oligosaccharides (GOS), xylo-oligosaccharides (XOS), arabinoxylo-oligosaccharides (AXOS), transgalacto-oligosaccharides (TGOS) are extracted from plants sometimes followed by enzymatic hydrolysis, or produced by chemical or enzymatic synthesis (149). IMMP produced by the GtfB enzyme is a soluble dietary fiber that partially escapes digestion in the upper part of the gastrointestinal tract and small intestine. Thus, it has potential to be a prebiotic that selectively boosts growth of some selected bacteria in the large

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intestine. In this thesis, the EPS produced by L. reuteri 121 were used as carbon sources to stimulate the growth of some typical probiotic Bifidobacteria that secrete amylopullulanases (Chapter 4).

Although the physiochemical properties of oligo-/polysaccharides produced by 4,6-α-GTase from maltodextrins or starches have not been studied, the products synthesized by DDase with similar structures are comparable. The studies of these products may provide us with more thoughts of the application of 4,6-α-GTase. Thus, apart from acting as a dietary fiber, the IMMP may be applied in food as cryostabilizer, fat substitute, or low-calorie bulking agent for sweeteners (25). Moreover, as shown in Fig. 9, because of the changes in the external chains of starch, 4,6-α-GTase treatment may influence the rheological properties of starches, such as the viscous modulus, elastic modulus, and retrogradation properties, expanding the application scope of these enzymatically modified starches.

Scope of this thesis 4,6-α-GTases are newly identified glucanotransferases, producing soluble dietary fibers, isomalto-/malto-polysaccharides (IMMP) from starch and maltodextrins. The in vivo biological roles of 4,6-α-GTase remained to be explored, also in view of their limited distribution in LAB and the fact that the closely related GS clearly contribute to the formation of EPS from sucrose. 4,6-α-GTases potentially are of great commercial value as the starch substrates are cheap and widely available compared to the normal sources of soluble dietary fiber. However, industrial application is still a challenge because 4,6-α-GTases are only available in relatively minor amounts. There is also a general lack of reliable 4,6-α-GTase enzyme activity assays. These topics are addressed in this PhD thesis, also focusing on elucidation of the 4,6-α-GTase reaction mechanism by 3D structural protein analysis and mutagenesis.

Chapter 1 reviews the current knowledge of 4,6-α-GTases in comparison with other specificity- or structurally-related enzymes including members in clan GH-H and DDases.

In chapter 2, active 4,6-α-GTase GtfB protein was prepared from inclusion bodies as it is highly expressed in the heterologous host E. coli, but largely

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accumulates in inclusion bodies. Following conventional denaturing, refolding of GtfB protein resulted in partial recovery of GtfB (hydrolysis) activity, but this required a set of complicated steps. The non-classical inclusion body (ncIB) GtfB enzyme directly extracted from IBs was much more thermostable than the soluble and refolded GtfB enzymes. In view of its high yield, low preparation cost and structural stability, ncIB GtfB protein thus provides a promising option for industrial applications.

In chapter 3, the soluble expression of GtfB was further improved by N-terminal truncation of the full-length protein, resulting in strong enhancement of the soluble expression level of fully active GtfB-∆N (approx. 75 fold compared to full length wild type GtfB) in E. coli. In addition, quantitative assays based on amylose V as substrate are described, allowing accurate determination of both hydrolysis (minor) activity and total activity, and calculation of the transglycosylation (major) activity of these 4,6-α-GTase enzymes, such as GtfB and GtfW. Using these assays, the biochemical properties of GtfB-∆N were characterized in detail, including determination of kinetic parameters and acceptor substrate specificity. Moreover, the GtfB-∆N enzyme displayed high conversion yields at relatively high substrate concentrations, a promising feature for industrial applications.

Chapter 4 reports the in vivo homo-exopolysaccharide (homo-EPS) synthesis by LAB strains with starch as substrate. L. reuteri 121 modifies amylose and the side chain of amylopectin molecules, introducing α1→6 linkages, producing soluble dietary fiber that is structurally similar to the IMMP product synthesized in vitro by the GtfB enzyme. This study provides insights into the in vivo role of 4,6-α-GTase in LAB strains. The IMMP-like EPS produced in vivo can be used as carbon source to stimulate the growth of some typical probiotic Bifidobacterium strains, suggesting that these EPS have potential prebiotic activity and may contribute to the application of probiotic L. reuteri 121 in food and feed (synbiotics).

In chapter 5, a synergistic action of 4,6-α-GTase GtfB and GS GtfA in L. reuteri 35-5 was studied when both starch and sucrose substrates were present. The EPS produced by L. reuteri 35-5 in the presence of sucrose and starch is different from the EPS derived from either of the single substrates in both size and structure. In

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vitro incubations with both GtfA and GtfB and sucrose plus starch further confirmed that both enzymes initially synthesized new oligosaccharides structurally different from those derived from either of the single substrates. This in vivo EPS formation from multiple dietary carbohydrates most likely reflects EPS formation in the oral cavity, potentially also affecting dental plaque formation.

In chapter 6, the GtfB-ΔNΔV protein was successfully crystallized and its high resolution structure solved. It has an overall domain organization resembling GH70 glucansucrases, and an active site architecture reminiscent of GH13 α-amylases. A tunnel-like binding groove allows for a dual mode of action for 4,6-α-GTases involving both exo-specific and endo-specific cleavage/transfer reactions, and explains why GtfB only processes the external (amylose) branches of starch substrates. The GtfB-ΔNΔV crystal structure clearly supports the view that the 4,6-α-GTase subfamily is an evolutionary intermediate between the family GH70 and GH13 enzymes. The structural observations also support the phylogenetic and genomic analyses, revealing a close relation between 4,6-α-GTases and GS, and suggesting that gene duplication as well as horizontal gene transfer events occurred during evolution. We propose a common ancestor for the two enzyme functionalities; loop mutations near the active site likely resulted in either a different substrate specificity (in GS) or in an (α1→6) specific transglycosylation activity (in 4,6-α-GTases). Thus, the evolution of bacterial species like L. reuteri 121, using either their 4,6-α-GTase or GS to synthesize related types of α-glucan with (α1→4) and (α1→6) linkages, may have been driven by the availability of fermentable substrates such as starch and (later) sucrose in the oral cavity.

Chapter 7 summarizes and discusses the results reported in this thesis and proposes topics for future research.

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148. Roberfroid M, Gibson GR, Hoyles L, McCartney AL, Rastall R, Rowland I, Wolvers D, Watzl B, Szajewska H, Stahl B, Guarner F, Respondek F, Whelan K, Coxam V, Davicco MJ, Leotoing L, Wittrant Y, Delzenne NM, Cani PD, Neyrinck AM, Meheust A. 2010. Prebiotic effects: metabolic and health benefits. Brit J Nutr 104:S1-S63. 149. Saad N, Delattre C, Urdaci M, Schmitter JM, Bressollier P. 2013. An overview of the last advances in probiotic and prebiotic field. LWT-Food Sci Technol 50:1-16.

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Chapter 2

Characterization of the 4,6-α-glucanotransferase GtfB enzyme of

Lactobacillus reuteri 121 isolated from inclusion bodies

Yuxiang Bai1,3, Rachel Maria van der Kaaij1, Albert Jan Jacob Woortman2,

Zhengyu Jin3, Lubbert Dijkhuizen1

1Microbial Physiology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, Nijenborgh 7, 9747 AG Groningen, The Netherlands

2Department of Polymer Chemistry, Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 4, 9747 AG Groningen, The Netherlands

3The State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi 214122, China

BMC Biotechnology (2015), 15:49

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Abstract The GtfB enzyme of the probiotic bacterium Lactobacillus reuteri 121 is a 4,6-α-glucanotransferase of glycoside hydrolase family 70 (GH70; http://www.cazy.org). Contrary to the glucansucrases in GH70, GtfB is unable to use sucrose as substrate, but instead converts malto-oligosaccharides and starch into isomalto-/malto- polymers that may find application as prebiotics and dietary fibers. The GtfB enzyme expresses well in Escherichia coli BL21 Star (DE3), but mostly accumulates in inclusion bodies (IBs) which generally contain wrongly folded protein and inactive enzyme. Denaturing and refolding of IB proteins has become a widely accepted method for obtaining active enzymes. Surprisingly, without unfolding and refolding, the GtfB IBs were enzymatically active, and therefore can be considered as non-classical IBs (ncIBs). Expression of GtfB in E. coli yielded > 100 mg/l relatively pure and active but mostly insoluble GtfB protein in IBs, regardless of the expression conditions used. Following denaturing, refolding of GtfB protein was most efficient in double distilled H2O. Also, GtfB ncIBs were active, with approx. 10% of hydrolysis activity compared to the soluble protein. When expressed as units of activity obtained per liter E. coli culture, the total amount of ncIB GtfB expressed possessed around 180% hydrolysis activity and 100% transferase activity compared to the amount of soluble GtfB enzyme obtained from one liter culture. The product profiles obtained for the three GtfB enzyme preparations were similar when analyzed by HPAEC and NMR. SEC investigation also showed that these 3 enzyme preparations yielded products with similar size distributions. FT-IR analysis revealed extended β-sheet formation in ncIB GtfB providing an explanation at the molecular level for reduced GtfB activity in ncIBs. The thermostability of ncIB GtfB was relatively high compared to the soluble and refolded GtfB. In view of their relatively high yield, activity and high thermostability, both refolded and ncIB GtfB derived from IBs in E. coli may find industrial application in the synthesis of modified starches.

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Introduction GtfB is a novel family GH70 enzyme isolated from the probiotic bacterium Lactobacillus reuteri 121 (1). The GtfB protein sequence is similar to that of the glucansucrase GtfA of L. reuteri 121 (61% similarity), but these structurally related proteins have different enzymatic activities (2). Unlike the common GH70 glucansucrases, GtfB is inactive on sucrose, but acts as a 4,6-α-glucanotransferase: It cleaves an α1→4 glycosidic linkage at the non-reducing end of a starch or malto-oligosaccharide (donor) chain, and transfers the glucose to the non-reducing end of another (acceptor) molecule, predominantly forming α1→6 glycosidic linkages, yielding mixtures of linear malto-/isomalto-oligosaccharide chains (1, 3, 4). GtfB also has a relatively low hydrolysis activity. The GtfB final products are very interesting functional carbohydrates, acting as prebiotic oligosaccharides and soluble dietary fiber (5). The enzymatic conversion of the widely available and low cost substrate starch by GtfB yields potentially valuable food ingredients and provides interesting opportunities for industrial applications (4).

Heterologous expression of GtfB in E. coli renders a low amount of soluble GtfB protein and a large amount of GtfB in inclusion bodies (IBs) as described previously for the related GtfML4 enzyme (6). IBs are generated by protein aggregation, resulting in inactive enzymes, and are commonly observed in heterologous expression systems (7). Several studies have shown that active enzymes may be isolated from IBs through proper processing (8). The conventional method involves refolding of the denatured protein. In the first step the misfolded protein is unfolded, followed by a second, gentle, refolding step (9, 10). Several successful refolding methods have been developed, such as dilution, on-column chromatography, dialysis, ultra-filtration and procedures involving multiple steps (11-13). Success in protein refolding, however, varies strongly, and these procedures are often time consuming and relatively expensive (14).

Interestingly, in case of GtfB, we detected its enzyme activity in the IBs, which therefore were renamed into non-classical inclusion bodies (ncIBs) (11, 15). Activity of IB aggregated enzymes may reflect the presence of a percentage of properly folded functional protein (14). Another viewpoint, the “IB-stretch hypothesis”, poses that IBs may include functional enzyme domains, and that

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crucial residues for active protein conformation are not engaged in the inactive β-core of the aggregates (16). These IBs also provide enough pore space for substrate and product molecules (14). Functional proteins in ncIBs thus can be applied directly; one example is the human granulocyte-colony stimulating factor (15). These ncIBs may have additional advantages over solubilized protein, such as high stability and relatively large particle size, useful for enzyme immobilization and preparation of nanomaterials (17, 18).

Heterologous expression attempts with L. reuteri 4,6-α-glucanotransferase enzymes (e.g. GtfB, GtfW and GtfML4) in E. coli yielded low amounts of soluble protein (6). However, expression of GtfB in E. coli BL21 Star (DE3) resulted in an abundant accumulation of GtfB in IBs, as described in this study. Here we report the results of a comparison of soluble GtfB and ncIB GtfB with enzyme preparations obtained by refolding of GtfB protein extracted from IBs, with emphasis on their activity, product specificity and thermostability.

Material and methods

Bacterial strains, plasmids, medium and carbohydrate

Plasmid pET15b was used for GtfB [GenBank: AAU08014.2] (C-terminal His-tag) expression in Escherichia coli strain BL21 Star (DE3) (Invitrogen, Taastrup, Denmark).

E. coli strain BL21 Star (DE3) was grown in Luria-Bertani (LB) medium with 100 mg/l ampicillin and 25 mg/l kanamycin medium. GtfB expression was induced using 0.4 mM Isopropyl β-D-1-thiogalactopyranoside (IPTG). AVEBE MD20 with DE (dextrose equivalent) value 20 was provided by AVEBE (Veendam, The Netherlands). LiBr was purchased from Fisher Scientific and pullulan standards from PSS (Polymer Standard Service, Mainz, Germany). All other materials and chemicals were purchased from Sigma-Aldrich (St. Louis, US).

Growth and inducing conditions

Soluble GtfB expression in E. coli BL21 Star (DE3) was achieved using the procedures described by Kralj et al. (1). For preparation of IBs, the method was modified as follows: Pregrown E. coli BL21 Star (DE3) inoculum was added to 1

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liter medium in a 5 liter shake flask and grown aerobically at 37 °C and 220 rpm until an OD600 of 0.9. The culture was equally divided into four 1 liter flasks and 0.4 mM IPTG was added to induce GtfB expression. The four flasks of 250 ml broth were incubated at induction temperatures of 18, 25, 30 and 37 °C, respectively. Cultures were harvested at OD600 of approx. 1.8. Biomass was collected by centrifugation at 15,000 ×g for 30 min.

Isolation of soluble protein and inclusion bodies

The collected E. coli cell pellets from 250 ml culture were washed with buffer (20 mM Tris-Cl, 50 mM NaCl, pH 8.0) and then resuspended in 8 ml of B-PER protein extraction buffer (Thermo, Rockford, US). To lyse E. coli cells and to remove nucleic acids, 16 µl lysozyme (150,000 U) and 4 µl DNase (20,000 U) were added and stirred for 30 min at room temperature. The proportions of soluble and total GtfB were determined by densitometric analysis of Coomassie-stained SDS-PAGE gels with 8 µl sample, using a Bio-Rad Model Imaging Densitometer (Bio-Rad Laboratories, Hercules, US). The IBs were washed twice with washing buffer (20 mM Tris-Cl, 0.5 mM EDTA, pH 8.0).

Purification of soluble GtfB

The extracted soluble GtfB with 6×His tag was purified by binding to Ni2+ nitrilotriacetic acid (Ni-NTA) as described previously (19). GtfB was eluted with elution buffer (20 mM Tris-HCl, pH 8.0, 200 mM imidazole, 1 mM CaCl2), and further purified by anion-exchange chromatography as described previously (1) using a 1-ml HiTrap Q HP column (GE Healthcare, Uppsala, Sweden). Finally, NaCl was removed using a 5-ml Hitrap desalting column (GE Healthcare) run with wash buffer (20 mM Tris-HCl, pH 8.0, 1 mM CaCl2) (1).

iFOLD Protein Refolding system and preparation of refolded GtfB

The iFOLD Protein Refolding system, containing 94 different refolding buffers, purchased from Novagen (Merck KGaA, Darmstadt, Germany), was used to identify the optimal conditions for GtfB refolding.

A total of 0.5 g prepared IBs was dissolved in 12 ml denaturing buffer (20 mM Tris-HCl with 4.47 % N-lauroylsarcosine sodium salt, 5% glycerol, 50 mM NaCl, 5 mM TCEP, 0.5 mM EDTA, pH 8.0) and stirred till transparent. The solution was centrifuged at 15,000 ×g for 15 min. The supernatant was dialyzed twice

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against 1 liter dialysis buffer (10 mM Tris-HCl, 0.06% N-lauroylsarcosine sodium salt, 0.05 mM EDTA, 0.1 mM TCEP, pH 8.0) at 4 °C for 6 h. At this stage, all GtfB protein was unfolded and no activity could be detected. The protein was refolded by diluting the sample to around 0.1 mg/ml in the different refolding buffers within the iFOLD system, as well as dd H2O, followed by stirring overnight at room temperature. Activity of the refolded GtfB protein was assessed by measuring hydrolysis of maltoheptaose, as described below.

Refolded GtfB with 6×His tag was further purified using Ni-NTA affinity chromatography and anion exchange methods as described previously (1). Protein purity was assessed on 8% SDS-PAGE and protein concentrations measured with Bio-Rad protein assay (Bio-Rad Laboratories, Hercules, US) with bovine serum albumin as standard.

Non-classical inclusion bodies (ncIB) GtfB preparation

The wet IB pellet (0.5 g) was resuspended and homogenized with 10 ml non-denaturing buffer (20 mM Tris-HCl, 0.2% N-lauroylsarcosine, 20% glycerol, pH 8.0) by gently pipetting and short sonication for 10 sec (Soniprep 150, MSE Ltd, London, UK). The suspension was shaken overnight at room temperature (11). The ncIB suspensions were stored at 4 °C, and were homogenized every time before use.

To measure protein content, ncIB suspensions were dissolved in denaturing buffer, followed by dialysis against protein storage buffer (20 mM Tris-HCl, pH 8.0) to remove N-lauroylsarcosine which influences the protein dye reagent (Bio-Rad Co. Ltd, US). Concentrations of these dialyzed proteins were measured by the Bio-Rad Protein assay. Finally, the concentration of ncIB GtfB was calculated from the total concentration of insoluble protein and the percentage of GtfB (Table 1) which was obtained from the densitometric analysis of Coomassie-stained SDS-PAGE gels.

Fourier Transformed Infrared analysis

Soluble and refolded GtfB proteins (0.5 - 3 mg) were precipitated from solution by the addition of hydrated ammonium sulfate (final concentration 30% w/v). ncIBs prepared at various E. coli growth temperatures (18, 25, 30, 37 °C), were washed in dd H2O. All wet pellets were dried in the Savant DNA Speed-Vac

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system for 1-2 h prior to analysis to reduce water interference in the infrared spectra. The infrared spectra of protein samples were recorded on a Bruker IFS66S spectrometer equipped with a liquid-nitrogen cooled MTC detector and Golden-Gate ATR diamond cell. The infrared spectra allowed monitoring of the secondary structure of ncIB GtfB protein prepared at different temperatures in comparison with properly folded and soluble GtfB protein. Typically, 64 scans were collected at a resolution of 4 cm-1 for all samples. Before the examination of the amide I bands, solvent spectra were recorded and subtracted (11). The structure of the amide I region was also analyzed by second derivatives.

Product profiles and enzyme activity assays

The product profiles of the various GtfB preparations were investigated by incubating enzyme in sodium acetate buffer (25 mM, pH 4.7, 1 mM CaCl2) at 37 °C for 24 h, with maltose, maltotriose and maltoheptaose separately as substrates (2). After 24 h or 72 h reactions, all tubes were boiled for 5 min followed by 15,000 × g centrifugation for 5 min. The supernatants were kept for high-pH anion-exchange chromatography (HPAEC) analysis as described below.

GtfB hydrolysis rates were assessed by measuring glucose release. GtfB enzyme solutions (50 µl) with different protein concentrations were added to a final 500 µl reaction system with 10 mM maltoheptaose. At a time interval of 5 min, 50 µl samples were taken and the reaction terminated by addition of 25 µl 0.4 M NaOH, followed by neutralization with 25 µl 0.4 M HCl. The GOPOD kit (Megazyme) was used to detect glucose, as a measure for hydrolysis activity (6). One unit of hydrolysis activity was defined as the amount of enzyme that produces 1 µmol glucose per min. Transferase activity was measured by determining maltose generation when GtfB enzyme was incubated with amylose (0.25%, w/v) as donor and glucose (10 mM) as acceptor substrate. One unit of transferase activity was defined as the amount of enzyme that produces 1 µmol maltose per min.

High-pH anion-exchange chromatography

The reaction products of GtfB were injected onto a 4×250 nm CarboPac PA-1 column connected to a Dionex DX500 workstation (Dionex). Samples were run with a gradient of 30-600 mM NaAc in 100 mM NaOH (1 ml/min), and detected by an ED40 pulsed amperometric detector. A mixture with known concentrations

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of glucose, maltose, panose, maltotriose, maltotetraose, maltopentaose, maltohexaose and maltoheptaose was used as reference.

Nuclear Magnetic Resonance spectroscopy

NMR spectroscopy resolution-enhanced 1D 500-MHz 1H NMR spectra were recorded in D2O on a Varian Inova 500 Spectrometer (NMR Center, University of Groningen) at probe temperatures of 300 K. Samples were exchanged twice with D2O (99.9 atm% D, Cambridge Isotope Laboratories, Inc.) with intermediate lyophilization and then dissolved in 0.6 ml D2O. Chemical shifts (δ) were expressed in parts per million by reference to internal acetone (δ 2.225 for 1H).

Size-exclusion chromatography

DMSO-LiBr (0.05M) was prepared by stirring for 3 h at room temperature followed by degassing for 15 min an ultrasonic cleaner (Branson 1510, Branson, Danbury, CT). Samples were dissolved at a concentration of 4 mg/ml in DMSO-LiBr by overnight rotation at room temperature, followed by 30 min heating in an oven at 80 °C obtaining clear sample solutions. The samples were cooled to room temperature and filtered through a 0.45-µm Millex PTFE membrane (Millipore Corporation, Billerica, MA). The SEC system set-up (Agilent Technologies 1260 Infinity) from PSS (Mainz, Germany) consisted of an isocratic pump, auto sampler without temperature regulation, an online degasser, an inline 0.2 μm filter, a refractive index detector (G1362A 1260 RID Agilent Technologies), viscometer (ETA-2010 PSS, Mainz) and MALLS (SLD 7000 PSS, Mainz). WinGPC Unity software (PSS, Mainz) was used for data processing. The samples (100 µl) were injected with a flow rate of 0.5 ml/min by an autosampler into a PFG guard column with DMSO-LiBr as eluent. The separation was done by three PFG-SEC columns with porosities of 100, 300 and 4000 Å. The columns were held at 80˚C, the refractive index detector at 45 °C and the viscometer was thermostatted at 60 °C. A standard pullulan kit (PSS, Mainz, Germany) with molecular mass from 342 to 805000 Da was used. The specific RI increment value dn/dc was measured by PSS and is 0.072 (private communication with PSS).

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Thermostability determination

The thermostability of the soluble, ncIB and refolded GtfB proteins with the same concentration (75 µg/ml) and the same hydrolysis activity (0.79 U, 25.0 µg/ml soluble GtfB, 38.7 µg/ml refolded GtfB and 249.4 µg/ml 25 °C ncIB GtfB) were determined by incubation in 25 mM sodium acetate buffer (pH 4.7) at 45 °C. Samples were taken at several time intervals up to 30 min, and the residual hydrolysis activity was determined subsequently as described above.

Results and Discussion

GtfB accumulates as soluble protein and in inclusion bodies (IBs)

Heterologous expression of GtfB in E. coli BL21 Star (DE3) resulted in relatively abundant synthesis of this protein but mainly in the form of inclusion bodies. As shown in table 1, this E. coli expression system yielded approximately 150-225 mg of total cell protein per liter of culture, more than 50% of which was the target GtfB protein. However, more than 90% of total GtfB protein accumulated in IBs (Table 1).

Table 1. The yields of total cell protein, and total, soluble, insoluble GtfB, obtained from E. coli. E.coli BL21 Star (DE3) cells grown at different temperatures, and the percentages of insoluble GtfB within the inclusion bodies (IBs) were determined by Bio-Rad protein assay and densitometric analysis of SDS-PAGE separated proteins. All cell samples were harvested at a culture OD600 of 1.8. The experiments were done in duplicate.

Induction temperature (°C)

Total protein (mg/l culture)

Total GtfB (mg/l culture)

Soluble GtfB (mg/l

culture)

Insoluble GtfB (mg/l culture)

% of insoluble

GtfB in total IBs

18 168.5±10.0 92.0±5.5 6.0±1.0 86.0±5.0 82.5±0.5

25 175.5±17.5 95.0±9.5 8.0±1.5 87.0±9.0 85.0±0.5

30 224.5±12.5 132.5±7.5 1.5±0.5 131.0±7.0 83.0±0.5

37 150.5±9.0 86.0±5.0 0.5±0.0 85.5±0.5 73.5±0.5

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Using different induction temperatures, the yields of total protein, soluble and insoluble GtfB protein varied in cultures harvested at the same OD600 value of 1.8. The yield of soluble GtfB increased from 0.5 mg/l at 37 °C to 8.0 mg/l at 25 °C, but no further increase occurred at 18 °C. After His-tag affinity purification, only approx. 1 mg pure GtfB was obtained from 1 liter culture incubated at 18 or 25 °C. The highest amount of insoluble GtfB (131.0 mg/l) was produced at 30 °C, while the purest (85.0%) insoluble GtfB was obtained at 25 °C.

Comparison of hydrolysis and transferase activities of soluble, refolded and

ncIB GtfB

The iFOLD protein refolding matrix with 94 different buffers was used to determine the optimal refolding conditions for GtfB IBs. Refolding efficiency was assessed by measuring hydrolysis activity (glucose release from maltoheptaose). As shown in Fig. 1, refolding was more successful at pH 7.0 and 7.5 compared to pH 8.0 and 8.5. Double distilled (dd) H2O at pH 5.0 without any additives and buffer salts was the optimal refolding environment. Through dd H2O refolding, 64.8% hydrolysis activity was recovered compared to soluble GtfB (see Fig. 1). In addition, among all the refolding conditions tested, conditions with methyl-β-cyclodextrin all resulted in higher efficiency than the ones without methyl-β-cyclodextrin (Fig. 1A). Conceivably, the hydrophobic areas of the GtfB protein are exposed during the folding process and addition of methyl-β-cyclodextrin suppresses the stacking of protein folding intermediates (20).

Non-classical inclusion body (ncIB) research shows that the presence of active protein inside IBs strongly depends on the E. coli growth temperature (11, 15). An example is the expression of human granulocyte-colony stimulating factor in E. coli: compared to 37 °C expression at 25 °C resulted in higher amounts of correctly folded protein inside IBs (15). Using equal amounts of protein, four ncIB GtfB samples obtained at different E. coli growth temperatures displayed relatively low hydrolysis activity in comparison to the soluble and refolded GtfB enzymes (Fig. 1A). The GtfB activity (hydrolysis) of the most active 25 °C ncIBs GtfB was 10.1 % and 15.6 % of that of the soluble and dd H2O refolded GtfB, respectively. To exclude any effect from E. coli proteins, E. coli harboring the

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Fig. 1. Bargraphs of the relative activities of soluble, refolded and different ncIB GtfB enzymes. The refolded GtfB enzymes were obtained from various refolding buffers, and the non-classical inclusion bodies (ncIBs) GtfB were expressed at different temperatures (A). All enzyme concentrations used were 100 µg/ml. The enzyme activity was defined as the release of glucose from maltoheptaose at 37 °C and pH 4.7.The hydrolysis activity of soluble GtfB was set at 100% (0.17 U). The total hydrolysis and transferase activities of soluble, refolded and ncIB GtfB enzymes obtained from one liter E. coli culture expressing GtfB are shown in bargraphs (B) and (C). In (B) 100% activity corresponds to 13.44 U hydrolysis activity and in (C) the 100% value corresponds to a transferase activity of 4.3 U in 1 liter culture.

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pET15b-gtfBD1015N (encoding inactive GtfB protein) was used as a negative control (1).

Based on the yield of 1 liter culture, GtfB ncIBs contained around 180% hydrolysis activity and 100% transferase activity compared to soluble GtfB (Fig. 1B and 1C). Thus ratios of hydrolysis versus transferase activity (Fig. 1 B and C) of ncIB and refolded GtfB were relatively high compared to soluble GtfB. The activity obtained from the total yield of refolded protein in one liter culture is much higher (Fig. 1B and 1C), but the production costs for refolded proteins are estimated to be around 20 times higher than for ncIB proteins (14). The ncIB GtfB proteins show sufficient activity to replace refolded GtfB protein as functional GtfB enzyme.

Comparison of product profiles of soluble, refolded and ncIB GtfB enzymes

incubated with maltose and maltotriose

Soluble (25.0 µg/ml), refolded (38.7 µg/ml) and 25 °C ncIB GtfB (249.4 µg/ml) samples with equivalent GtfB hydrolysis activity (0.04 U) were incubated with 50 mM maltose or maltotriose at pH 4.7 and 37 °C for 72 h, and their product profiles were analyzed by HPAEC (Fig. 2A, 2B). The product profiles from soluble, refolded and ncIB GtfB enzymes were very similar. The generated products were identified based on earlier research by Dobruchowska et al. (2). The structures are depicted in Fig. 2C. In Fig. 2A, two main peaks (1 and 3) identified in all spectra represent glucose and panose, products derived from maltose by hydrolysis and transferase activities. The other reaction products labelled as peak 4a and 5 were panose elongated with one and two (α1→6)-linked glucose residues at the non-reducing site. Also after incubation with maltotriose (Fig. 2B), the product profiles of soluble, refolded and ncIB GtfB were highly similar when comparing peaks 7 to 13. The structures of these products (peaks 7-13) indicate that the larger oligosaccharides resulted from elongation with not only α1→6 but also α1→4 linkages. These data showed that refolded and ncIB GtfB enzymes have the same product and reaction specificity as soluble GtfB, catalyzing hydrolysis and α1→4/α1→6 transglycosylation .

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Fig. 2. Comparison of HPAEC-PAD profiles of the product mixtures of different GtfB preparations. 50 mM maltose (A) and maltotriose (B) were seperately incubated with GtfB preparations with equal hydrolysis activity (25.0 µg/ml soluble GtfB, 38.7 µg/ml refolded GtfB and 249.4 µg/ml 25 °C ncIB GtfB) after 72 h at 37 °C and pH 4.7. The oligosaccharide structures produced were identified (C) according to Dobruchowska et al. (28). G1 to G7 represent glucose, maltose, maltotriose, maltotetriose, maltopentaose, maltohexaose and maltoheptaose, respectively.

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Additionally, NMR analysis (Fig. 3) showed that the α1→6 linkage percentages of AVEBE MD20 (maltodextrins manufactured from potato starch, average degree of polymerization 6) modified by soluble, refolded and ncIB GtfB enzymes during incubation for 3 days were highly similar, increasing in all cases from 0.5% to around 15%.

Fig. 3. One-dimensional 1H NMR spectra of AVEBE MD20 and its products following incubation with GtfB preparations. The spectra were recorded in D2O at 300 K after the incubation of 5% AVEBE MD20 (A) with different GtfB preparations which have equal hydrolysis activity (25.0 µg/ml soluble GtfB, B, 38.7 µg/ml refolded GtfB, C and 249.4 µg/ml 25 °C ncIB GtfB, D) after 72 h at 37 °C and pH 4.7. Rα/β represent the reducing –(1→4)-D-Glcp units and Gα/β represent the D-Glcp units.

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Size analysis of AVEBE MD20 and its products after GtfB enzyme

treatments

The Size Exclusion Chromatograms (SEC) of AVEBE MD20 before and after incubation with soluble, refolded and ncIB GtfB proteins are shown in Fig. 4. The elution volume in SEC is directly related to the hydrodynamic volume of the linear and branched molecules (21). The elution volumes of some pullulan standards are plotted at the x-axis. Since AVEBE MD20 is mainly composed of short oligosaccharides (average degree of polymerization 6), its major peak is seen at a high elution volume (around 32.5 ml). After incubation with GtfB preparations with equivalent hydrolysis activity, the peak at 32.5 ml decreased and shifted to a bi-modal peak at elution volumes of approximately 30 and 32 ml, demonstrating that the short oligosaccharides in MD20 substrate were converted to products with higher molecular mass. The distributions of the products of soluble, ncIB and refolded GtfB were highly similar. The relatively high ratios of hydrolysis versus transferase activity (Fig. 1 B and C) of ncIB and refolded GtfB compared to soluble GtfB may initially result in enhanced synthesis of smaller products. This increased availability of shorter acceptor substrates resulted in reduced average sizes of their product molecules.

Fourier Transform Infrared (FT-IR) studies

The secondary structures of the soluble, refolded and ncIB GtfB (25 °C) proteins were examined by infrared spectroscopy (Fig. 5). The frequency and the shape of the amide I bands in the spectral region between 1690 and 1620 cm-1 provides information about the type of secondary structure present in proteins (22). Because the components in amide I, resulting from different secondary structure elements, are strongly overlapping (Fig. 5A), a second derivative analysis was applied. The main band, at 1653 cm-1 in second derivative spectra (Fig. 5B) indicates the presence of α-helical structures, which may be expected for a GH70 family enzyme containing a TIM barrel fold with 8 α-helices (23). The ncIBs GtfB show distinctive bands at 1627 and 1695 cm-1 (Fig. 5B) with higher intensity than those of soluble and refolded GtfB, indicating that the inclusion bodies have more intermolecular β-sheet structure (24). The FI-IR spectra of the inclusion bodies also show peaks at 1633 and 1653 cm-1, suggesting they also

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contain some native β-sheet and native α-helix structures of the soluble GtfB enzyme, respectively (25).

Fig. 4. SEC chromatograms of AVEBE MD20 and its products following incubation with GtfB preparations. Different GtfB preparations with equal hydrolysis activity (25.0 µg/ml soluble GtfB, 38.7 µg/ml refolded GtfB and 249.4 µg/ml 25 °C ncIB GtfB) were incubated with 5% AVEBE MD20 for 72 h at 37 °C and pH 4.7. Elution volumes of the pullulan standards corresponding to 366, 200, 113, 48.8, 21.7, 10, 6.2, 1.32 and 0.342 kDa, are plotted above the x-axis.

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Fig. 5. Fourier Transform Infrared (FT-IR) spectra in the amide I region of different GtfB preparations. Soluble GtfB, refolded GtfB, and ncIB GtfB proteins isolated from E. coli incubated at different temperatures (25 and 30 °C) were analyzed. (A) FT-IR spectra; (B) second derivatives of the FT-IR spectra.

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Thermostability comparison of soluble, ncIB and refolded GtfB proteins

To compare the thermostability of soluble, ncIB and refolded GtfB proteins, their residual activity was measured after incubation at 45 °C for several time intervals. At a protein concentration of 75 µg/ml, the half-lives of soluble and refolded GtfB were around 3.5 min (Fig. 6A). The half-life of ncIB GtfB protein was 3.0-fold longer. Even after 30 min incubation, 20% of the original activity of ncIB GtfB enzyme remained, while soluble and refolded GtfB enzymes had become completely inactivated. When comparing the three GtfB protein preparations in different concentrations but with the same hydrolysis activity (0.04 U), the half-life of ncIB GtfB was 5.0 times longer than those of soluble and refolded GtfB (Fig. 6B). This is probably due to the extended β-sheet formation in ncIB GtfB protein by amino acid chains which are remote to the active site stabilizing its overall conformation. Thus, with increasing temperatures, domains outside the active site important for stable protein conformation may be more slowly affected in ncIB GtfB protein than in soluble and refolded GtfB proteins.

Fig. 6. Thermostability of soluble GtfB, refolded GtfB, and 25 °C ncIB GtfB proteins. Different GtfB preparactions incubated at the same protein concentration (75 µg/ml, A) and with the same hydrolysis activity (25.0 µg/ml soluble GtfB, 38.7 µg/ml refolded GtfB and 249.4 µg/ml 25 °C ncIB GtfB, B), were assessed separately by measuring their residual activities after incubation at 45 °C up to 30 min. The activity before incubation was set at 100%.

Conclusions This paper reports that the Lactobacillus reuteri 121 GtfB protein is highly expressed in the heterologous host E. coli, but largely accumulates in non-

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classical inclusion bodies (ncIBs), displaying hydrolysis and transferase activity. E. coli growth temperatures of 25 and 30 °C resulted in ncIB preparations with highest GtfB yield, purity and activity. Following denaturing, refolding of GtfB protein in double distilled H2O resulted in highest recovery of GtfB (hydrolysis) activity, but required a set of complicated procedures. The ncIB GtfB enzyme produced at 25 °C displayed 10.1% of the soluble GtfB (hydrolysis) activity and produced similar product profiles from maltose, maltotriose and AVEBE MD20. FT-IR analysis of soluble, refolded and ncIB GtfB proteins confirmed that structural differences exist between these GtfB proteins. The ncIB GtfB enzyme was much more thermostable than the soluble and refolded GtfB enzymes. In view of its high yield, low preparation cost and structural stability, ncIB GtfB protein thus provides a promising option for industrial applications.

Acknowledgements The study was financially supported by the Chinese Scholarship Council (to YB) and by the University of Groningen. LD acknowledges support from the TKI Agri&Food program as coordinated by the Carbohydrate Competence Center (CCC-ABC; www.cccresearch.com)

References 1. Kralj S, Grijpstra P, van Leeuwen SS, Leemhuis H, Dobruchowska JM, van der Kaaij RM, Malik A, Oetari A, Kamerling JP, Dijkhuizen L. 2011. 4,6-α-glucanotransferase, a novel enzyme that structurally and functionally provides an evolutionary link between glycoside hydrolase enzyme families 13 and 70. Appl Environ Microbiol 77:8154-8163.

2. Dobruchowska JM, Gerwig GJ, Kralj S, Grijpstra P, Leemhuis H, Dijkhuizen L, Kamerling JP. 2012. Structural characterization of linear isomalto-/malto-oligomer products synthesized by the novel GTFB 4,6-α-glucanotransferase enzyme from Lactobacillus reuteri 121. Glycobiology 22:517-528.

3. Leemhuis H, Pijning T, Dobruchowska JM, van Leeuwen SS, Kralj S, Dijkstra BW, Dijkhuizen L. 2013. Glucansucrases: Three-dimensional structures, reactions, mechanism, α-glucan analysis and their implications in biotechnology and food applications. J Biotechnol 163:250-272. 4. Dijkhuizen L, van der Maarel MJEC, Kamerling JP, Leemhuis H, Kralj S, JM D. 2010. Gluco-oligosaccharides comprising (α1→4) and (α1→6) glycosidic bonds, use thereof, and methods for providing them. WO/2010/128859.

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5. Leemhuis H, Dobruchowska JM, Ebbelaar M, Faber F, Buwalda PL, van der Maarel MJ, Kamerling JP, Dijkhuizen L. 2014. Isomalto/Malto-polysaccharide, a novel soluble dietary fiber made via enzymatic conversion of starch. J Agric Food Chem 62:12034-12044. 6. Leemhuis H, Dijkman WP, Dobruchowska JM, Pijning T, Grijpstra P, Kralj S, Kamerling JP, Dijkhuizen L. 2013. 4,6-α-Glucanotransferase activity occurs more widespread in Lactobacillus strains and constitutes a separate GH70 subfamily. Appl Microbiol Biotechnol 97:181-193. 7. Singh SM, Panda AK. 2005. Solubilization and refolding of bacterial inclusion body proteins. J Biosci Bioeng 99:303-310. 8. Chow MKM, Amin AA, Fulton KF, Whisstock JC, Buckle AM, Bottomley SP. 2006. REFOLD: An analytical database of protein refolding methods. Protein Expres Purif 46:166-171. 9. Feng YY, Liu L, Wang JP, Liu J, Hu W, Wang XN, Yang Z. 2012. Integrated refolding techniques for Schistosoma japonicum MTH1 overexpressed as inclusion bodies in Escherichia coli. Protein Expres Purif 84:181-187.

10. Freydell EJ, van der Wielen LAM, Eppink MHM, Ottens M. 2011. Techno-economic evaluation of an inclusion body solubilization and recombinant protein refolding process. Biotechnol Progr 27:1315-1328.

11. Peternel S, Grdadolnik J, Gaberc-Porekar V, Komel R. 2008. Engineering inclusion bodies for non denaturing extraction of functional proteins. Microb Cell Fact 7:1-9.

12. Liu XQ, Yang XQ, Xie FH, Song LY, Zhang GQ, Qian SJ. 2007. On-column refolding and purification of transglutaminase from Streptomyces fradiae expressed as inclusion bodies in Escherichia coli. Protein Expres Purif 51:179-186.

13. Bajorunaite E, Sereikaite J, Bumelis VA. 2007. L-Arginine suppresses aggregation of recombinant growth hormones in refolding process from E. coli inclusion bodies. Protein J 26:547-555.

14. García-Fruitós E, Vázquez E, Díez-Gil C, Corchero JL, Seras-Franzoso J, Ratera I, Veciana J, Villaverde A. 2012. Bacterial inclusion bodies: making gold from waste. Trends Biotechnol 30:65-70.

15. Jevsevar S, Gaberc-Porekar V, Fonda I, Podobnik B, Grdadolnik J, Menart V. 2005. Production of nonclassical inclusion bodies from which correctly folded protein can be extracted. Biotechnol Progr 21:632-639.

16. Glover JR, Kowal AS, Schirmer EC, Patino MM, Liu JJ, Lindquist S. 1997. Self-seeded fibers formed by Sup35, the protein determinant of [PSI+], a heritable prion-like factor of S. cerevisiae. Cell 89:811-819.

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17. García-Fruitós E. 2010. Inclusion bodies: a new concept. Microb Cell Fact 9:80. 18. Peternel S, Komel R. 2010. Isolation of biologically active nanomaterial (inclusion bodies) from bacterial cells. Microb Cell Fact 9:66. 19. Kralj S, van Geel-Schutten GH, van der Maarel MJEC, Dijkhuizen L. 2004. Biochemical and molecular characterization of Lactobacillus reuteri 121 reuteransucrase. Microbiol-SGM 150:2099-2112. 20. Aachmann FL, Otzen DE, Larsen KL, Wimmer R. 2003. Structural background of cyclodextrin-protein interactions. Protein Eng 16:905-912.

21. Ahmadi-Abhari S, Woortman AJJ, Hamer RJ, Loos K. 2013. Assessment of the influence of amylose-LPC complexation on the extent of wheat starch digestibility by size-exclusion chromatography. Food Chem 141:4318-4323.

22. Krimm S. 1962. Infrared Spectra and Chain Conformation of Proteins. J Mol Biol 4:528-540.

23. Vujičić-Žagar A, Pijning T, Kralj S, Lopez CA, Eeuwema W, Dijkhuizen L, Dijkstra BW. 2010. Crystal structure of a 117 kDa glucansucrase fragment provides insight into evolution and product specificity of GH70 enzymes. P Natl Acad Sci USA 107:21406-21411.

24. Ami D, Natalello A, Taylor G, Tonon G, Doglia SM. 2006. Structural analysis of protein inclusion bodies by Fourier transform infrared microspectroscopy. BBA-Proteins Proteom 1764:793-799.

25. Wang L. 2009. Towards revealing the structure of bacterial inclusion bodies. Prion 3:139-145.

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Chapter 3

Biochemical characterization of Lactobacillus reuteri Glycoside

Hydrolase family 70 GtfB type of 4,6-α-glucanotransferase

enzymes that synthesize soluble dietary starch fibers

Yuxiang Bai1,2, Rachel Maria van der Kaaij1, Hans Leemhuis1,3, Tjaard Pijning4,

Sander Sebastiaan van Leeuwen1, Zhengyu Jin2, Lubbert Dijkhuizen1

1 Microbial Physiology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, The Netherlands

2 The State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi 214122, China

3 present address: AVEBE, Veendam, The Netherlands

4 Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, The Netherlands

Applied and Environmental Microbiology (2015), 81:7223-7232

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Abstract 4,6-α-Glucanotransferase (4,6-α-GTase) enzymes, such as GtfB and GtfW of Lactobacillus reuteri strains, constitute a new reaction specificity in Glycoside Hydrolase Family 70 (GH70) and are novel enzymes that convert starch or starch hydrolysates into isomalto/malto-polysaccharides (IMMPs). These IMMPs still have linear chains with some α1→4 linkages but mostly (relatively long) linear chains with α1→6 linkages, and are soluble dietary starch fibers. 4,6-α-GTase enzymes and their products have significant potential for industrial applications. Here we report that an N-terminal truncation (1-733 amino acids) strongly enhances the soluble expression level of fully active GtfB-∆N (approx. 75 fold compared to full-length wild type GtfB) in Escherichi coli. In addition, quantitative assays based on amylose V as substrate are described, allowing accurate determination of both hydrolysis (minor) activity (glucose release, reducing power) and total activity (iodine staining), and calculation of the transferase (major) activity of these 4,6-α-GTase enzymes. The data shows that GtfB-∆N is clearly less hydrolytic than GtfW, which is also supported by NMR analysis of their final products. Using these assays, the biochemical properties of GtfB-∆N were characterized in detail, including determination of kinetic parameters and acceptor substrate specificity. The GtfB-∆N enzyme displayed high conversion yields at relatively high substrate concentrations, a promising feature for industrial application.

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Introduction Starch is the second most abundant carbohydrate on earth and a major dietary carbohydrate for humans; as storage carbohydrate it is present in seeds, roots and tubers of plants (1). It consists of α-glucan polymers with (α1→4) linkages and a low percentage of (α1→6) linkages, in the form of amylose and branched amylopectin (2). Starches are applied in various industrial products such as food, paper, and textile, often after processing by physical, chemical or enzymatic treatment (3-6).

Dietary fibers and low glycemic index (GI) food are considered healthy food contributing to our long-term well-being (7, 8). Of all the nutritional types of starch, slowly digestible starch (SDS) with low GI has drawn strongest interest. Annealing/heat-moisture treatment, recrystallization and enzymatic treatment are recognized approaches to obtain SDS (9-11). SDS materials prepared by physical processing suffer losses upon boiling, therefore structural modifications through enzymatic treatment of starch are more desirable. In the human digestive system, the (α1→6) linkages in starch are hydrolyzed at a lower rate than (α1→4) linkages (12, 13). Branching enzymes, alone or in combination with β-amylase, are used to increase the percentage of (α1→4,6) branches in starches (12-14).

The 4,6-α-Glucanotransferase (4,6-α-GTase) enzymes such as GtfB, GtfW and GtfML4 of Lactobacillus reuteri strains constitute a subfamily of Glycoside Hydrolase Family 70; GH70 mainly consists of glucansucrases (GSes) (http://www.cazy.org). Unlike GSes, 4,6-α-GTases are not sucrose-acting enzymes but starch-converting enzymes, capable of converting (1→4)-α-D-gluco-oligosaccharides into dietary fiber isomalto/malto-polysaccharides (IMMPs). Conversion has been proposed to occur mainly by the step-wise addition of single glucose moieties onto the non-reducing end of an α-glucan, introducing linear chains with (α1→6) linkages (15-18). The 4,6-α-GTase enzymes have strong potential for SDS or soluble dietary fiber production in the starch industry (19).

The expression yields of all three studied 4,6-α-GTase enzymes in E. coli are rather low and most of the protein accumulates in inclusion bodies (18, 20); to obtain more active protein, strategies to use denatured refolded GtfB protein, or non-classical inclusion body preparations have been tested. The biochemical and

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catalytic properties of these enzymes, e. g. their hydrolysis and transferase activities, have not been characterized yet because of a lack of suitable quantitative assays. In the present study, the variable N-terminal region of the GtfB enzyme was removed (yielding construct GtfB734-1619) which resulted in increased expression of soluble and active GtfB-∆N enzyme in E. coli. In addition, we developed activity assays to characterize the reactions of 4,6-α-GTase enzymes with amylose. These assays were used to define the optimal reaction conditions of GtfB-∆N and GtfW-∆N, and to quantitatively compare their hydrolysis and transferase activities, as well as to determine key biochemical properties. The developed assays also provide a firm basis for the future characterization of other, natural and engineered, 4,6-α-GTase enzymes.

Materials and Methods

Construction of a truncated GtfB-ΔN mutant

Based on the alignment of the L. reuteri 121 GtfB with glucansucrase sequences, and the crystal structure of L. reuteri 180 Gtf180-ΔN glucansucrase (PDB entry 3KLK), the gtfB gene fragment encoding GtfB (Uniprot entry Q5SBM0) amino acids 734–1619 was amplified by PCR using High Fidelity PCR enzyme mix (Thermo-Scientific, Landsmeer, The Netherlands) with pET15b-GtfB as template and the primers CHisFor-dNgtfB 5′-GATGCATCCATGGGCCAGCTCATGAGAAACTTGGTTGCAAAACCTAATA-3′ and CHisRev-dNgtfB 5′-CCTCCTTTCTAGATCTATTAGTGATGGTGATGGTGATGGTTGTTAAAGTTTAATGAAATTGCAGTTGG-3′. A nucleotide sequence encoding a 6×His-tag was fused in-frame to the 3′ end of the gtfB-ΔN gene, using the reverse primer. The resulting PCR product was digested with NcoI and BglII and was ligated into the corresponding site of pET15b. The construct was confirmed by nucleotide sequencing (GATC, Cologne, Germany). Plasmid pET15b-gtfW-ΔN (with the gtfW gene fragment encoding GtfW-ΔN (Uniprot entry A5VL73) amino acids 458-1363) has been constructed previously (18).

Expression and purification of GtfB, GtfB-∆N and 4,6-αGT-W (GtfW-∆N)

The GtfB protein was produced in E. coli BL21 Star (DE3) carrying the plasmids pRSF-GtfB and pBAD22-GroELS (17). The bacterial inocula were prepared in

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0.4 L Luria-Bertani cultures and grown at 37°C and 220 rpm until the OD600 had reached 0.4-0.5, followed by addition of the inducer 0.4 mM isopropyl β-D-1-thiogalactopyranoside, and L-arabinose (0.02% w/v). The cultures were subsequently incubated at 18 °C and 160 rpm for 16 h in an orbital shaker. Cells were harvested by centrifugation (26000×g, 40 min) and then washed with 20 ml buffer (20 mM Tris/HCl pH 8.0, 250 mM NaCl, 1mM CaCl2). The collected cells were resuspended in 10 ml B-PER protein extraction reagent (Pierce, 100 μg/ml of lysozyme and 5 U/ml of DNase) for 30 min on a rolling device at room temperature, followed by centrifugation (10,000×g, 30 min). The encoded GtfB protein carrying a C-terminal (His)6-tag was purified by His-tag affinity chromatography using a 1-ml Hitrap IMAC HP column (GE Healthcare) and anion exchange chromatography (Resource Q, GE Healthcare). The salt was removed using a 5-ml Hitrap desalting column (GE Healthcare) run with B-PER and 1 mM CaCl2. Purity was checked on 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and protein concentration was measured by Bradford reagent (Bio-Rad, Hercules, CA) and bovine serum albumin as standard.

The L. reuteri DSM 20016 4,6-αGT-W (GtfW-∆N) and GtfB-∆N proteins were expressed and purified according to Leemhuis et al. with minor modification (18). E. coli BL21(DE3)-pET15b_gtfW-∆N/gtfB-∆N was grown in Luria broth containing 100 mg/l ampicillin. Protein expression was induced at an OD600 of 0.4–0.5 by adding isopropyl β-D-1-thiogalactopyranoside to 0.1 mM, and cultivation was continued at 18°C and 160 rpm for 16 h. Cells were harvested by centrifugation, in Tris-HCl buffer (50 mM, pH 8.0), containing NaCl (250 mM), Cell-free extracts were made by sonication followed by centrifugation (10,000×g, 1 h). Purification was performed as described above.

Preparation of amylose V substrate solution

Amylose V was provided by AVEBE (Veendam, The Netherlands). Note that Amylose V is a slightly degraded amylose, as a result of the extraction procedure used to isolate it from regular potato starch. The degree of branching of amylose V was below 0.1 %, as reported earlier (21). Amylose V (1%, w/v) was prepared as stock solution. Amylose V (20 mg) was suspended in 1 ml ddH2O, and dissolved by addition of 1 ml 2 M NaOH. The solution was homogenized until

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clear. Prior to use, the stock solution was neutralized by adding an equal volume of 1 M HCl and the stock was further diluted with reaction buffer.

Enzyme assays

The optimal pH and temperature of 4,6-α-GTases were determined over the pH range of 3.5-7.0 (25 mM sodium acetate (NaAc) buffer, pH 3.5-5.5; 25 mM MES buffer, pH 5.5-6.5; 25 mM MOPS buffer, pH 6.5-7.0) at 40 °C, and temperature range of 30-60 °C at pH 5.0 with 0.25% (w/v) amylose V as substrate. All other reactions were performed in reaction buffer (25 mM NaAc, pH 5.0, 1 mM CaCl2), with 0.25% (w/v) amylose V or 2.5% (w/v) maltodextrins (dextrose equivalent = 13-17, Sigma-Aldrich, US), or 10 mM maltoheptaose (Sigma-Aldrich, Missouri, US) as substrate, at 40 °C. In each reaction, 60 nM of enzyme was added. Every 5 min, 100 µl of reaction mixture was taken and the reaction was stopped by addition of 50 µl of 0.4 M NaOH. Afterwards, 50 µl 0.4 M HCl was added to neutralize the reaction mixture. These samples were stored for GOPOD (Megazyme International, Ireland) determination of glucose released, Nelson-Somogyi measurement of reducing power, and (changes in) iodine staining of amylose.

Paselli MD6 maltodextrins (partial hydrolysis product of potato starch with a dextrose equivalent between 5 and 7 and 2.9% α1→6 glycosidic linkages) was provided by AVEBE and variant concentrations (0.93-55.8% w/v) were incubated with GtfB-∆N [1/1000 (w/w) concentration of substrate] for 72 h at 37°C and pH 5.0.

Amylose-iodine assay

The iodine staining method was used to quantify (remaining) amylose in the reaction mixture (22). Thus, 0.26 g I2 and 2.6 g KI were dissolved in 10 ml distilled water to prepare the 260× concentrated stock. Prior to using the staining buffer, the stock was diluted 260× with distilled water. For each measurement, 150 µl iodine reagent was pipetted into microtiter plate wells, followed by addition of 15 µl enzyme reaction sample solution. The microtiter plate was shaken slowly for 5 min to allow color formation and the optical density was measured at 660 nm. Amylose V solutions (0-0.25%, w/v) were used to prepare a calibration curve.

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Reducing sugar measurement

The Nelson-Somogyi assay adapted to microtiter plates was used to measure the reducing power released through hydrolysis activity of the enzymes with amylose V (23). Thus, 40 µl Somogyi copper reagent, consisting of 4 parts KNa tartrate: Na2CO3:Na2SO4:NaHCO3 (1 : 2: 12 : 1.3) and 1 part CuSO4·5H2O:Na2SO4 (1 : 9) and 40 µl enzyme reaction sample were mixed and incubated at 90 °C for 30 min, after which the mixture was cooled to room temperature. After shaking the microtiter plate for 1 min to remove CO2, 160 µl arsenomolybdate (25 g ammonium molybdate in 450 ml H2O + 21 ml 98% H2SO4 + 3 g Na2HAsO4·7H2O dissolved in 25 ml H2O) was added and mixed by vortexing. Optical densities were read at 500 nm. A calibration curve was made using 0-200 µg/ml glucose solutions as standard.

Glucose measurement

Glucose was measured using the GOPOD kit which was adapted for microtiter plates. Five µl enzyme reaction sample was added to wells containing 195 µl GOPOD reagent, followed by incubation at 40 °C for 30 min. Optical densities were read at 510 nm. A calibration curve was made using 0-200 µg/ml glucose solutions as standard.

Definitions of hydrolysis (H) and total activities, and calculation of

transferase (T) activities

One unit of hydrolysis activity (H) is defined as the release of 1 mg glucose from amylose V per min in the GOPOD assay mixture.

One unit of total activity is defined as the decrease of 1 mg amylose V per min in the assay mixture (as determined by the amylose-iodine staining method).

Amylose V is either converted into free glucose via H or modified by transferase activity (T) introducing linear α1→6 linked glucose chains, losing ability to bind

iodine. T was calculated by subtracting H from Total Activity, 𝑻 �U 𝑚𝑚� � =

Total activity �U 𝑚𝑚� � − 𝑯 �U 𝑚𝑚� �. Also T/H activity ratios can be calculated

subsequently.

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Nuclear Magnetic Resonance (NMR) Spectroscopy

The reaction products were exchanged twice with D2O (99.9 atm% D, Cambridge Isotope Laboratories, Inc.) with intermediate lyophilization and then dissolved in 0.6 ml D2O. Resolution-enhanced 500-MHz 1D 1H NMR spectra were recorded with a spectral width of 4500 Hz in 16k complex data sets and zero filled to 32k in D2O on a Varian Inova Spectrometer (NMR Center, University of Groningen) at a probe temperature of 335 K. Suppression of the HOD signal was achieved by applying a WET1D pulse sequence. All spectra were processed using MestReNova 5.3.

Results

Improving expression of soluble GtfB protein

Heterologous expression of L. reuteri 4,6-α-GTase enzymes in E. coli is relatively poor. Optimization of the growth and induction conditions did not improve the soluble expression level of these enzymes and most of the GtfB and GtfW proteins accumulated in inclusion bodies. Improvement of soluble expression of these enzymes appeared essential, also in view of their potential industrial application. The 4,6-α-GTase enzymes constitute a subfamily of GH70, showing high similarity with glucansucrases in amino acid sequences. As previously reported, deletion of the N-terminal variable region of GH70 glucansucrases, such as GtfA, did not change the glycosidic linkage types present in the products, nor the product sizes but significantly improved the expression levels of the enzymes (24-26). Therefore, we applied a similar truncation approach to GtfB. Alignment of the GtfB and glucansucrase protein sequences (27) resulted in identification of amino acids 1-733 as the GtfB N-terminal variable region (Fig. 1A). This region contains 5 RDV repeats (sequence R(P/N)DV-x12-SGF-x19-22-R(Y/F)S, where x represents a non-conserved amino acid residue) which were also observed previously in Gtf180, GtfA and GtfML1 (28). Its deletion in GtfB resulted in expression of soluble GtfB-∆N at a much higher level compared to the full length GtfB. Moreover, the purity of GtfB-∆N protein was improved significantly compared to that of the full length GtfB enzyme (Fig. 1B). More than 40 mg pure GtfB-∆N protein was obtained from 1 liter E. coli culture, improving expression 75 fold based on molar calculation. The full length GtfB and GtfB-∆N enzymes were compared with regard to their

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reaction specificity and activity using amylose V and maltoheptaose as substrates (data not shown). With maltoheptaose (10 mM) as substrate, the rate of glucose release (GOPOD) and the product profiles of both enzymes were nearly identical. While using amylose V as substrate, the total and hydrolysis activities measured by the GOPOD and iodine staining assay described below were almost the same. The N-terminally truncated GtfB and the full length GtfB thus are virtually identical in activity and products.

Products derived from amylose V and maltodextrins by GtfB-∆N or GtfW-

∆N treatments

Linear (α1→4)-linked compounds are preferred substrates for 4,6-α-GTases, therefore amylose V and short chain (α1→4)-linked maltodextrins were compared as donor substrates for the GtfB-∆N and GtfW-∆N enzymes. 1H NMR analysis of the products of GtfB-∆N and GtfW-∆N incubated with amylose V [(1→4)-α-D-glucan] or maltodextrins [(1→4)-α-D-glucooligosaccharides] for 48 h revealed that both hydrolysis and transferase activities occurred. Besides the presence of (α1→4) linkages (H-1, δ ~5.40), (α1→6) linkages (H-1, δ ~4.97) were newly formed. In the product mixture derived from amylose V, the (α1→6):(α1→4) linkage ratio increased up to 90:10 (Fig. 2a1) for GtfB-∆N and 74:26 for GtfW-∆N (Fig. 2b1), when using identical amounts of purified protein. The spectra also showed the presence of free glucose (Glcα H-1, δ 5.225, Glcβ H-1, δ 4.637), the 4-substituted reducing glucose residues [-(1→4)-α-D-Glcp; Rα H-1, δ 5.225, Rβ H-1, δ 4.652], and a small amount of reducing-end glucose residues which are 6-substitued [-(1→6)-α-D-Glcp; Rα H-1, δ 5.241, Rβ H-1, δ 4.670] (32), suggesting that both GtfB-∆N and GtfW-∆N can use glucose as acceptor substrate, forming maltose, isomaltose and longer products in the presence of donor substrate. Comparison of the spectra Fig. 2a1 and Fig. 2b1 showed that GtfW-∆N produced a much higher percentage of free glucose (23%) compared to GtfB-∆N (8%) from amylose V. Conceivably, GtfW-∆N has a higher ratio of hydrolysis versus

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Fig. 1 (A) Linear schematic representation of the domain organization of full length Gtf180 of L. reuteri 180 and GtfB of L. reuteri 121 (with N-terminal variable region). Structural analysis of L. reuteri 180 Gtf180-ΔN protein has shown that its peptide chain follows a “U”-path and that four of its five domains are built up from discontinuous N- and C-terminal parts of the peptide chain. Only the C-domain is formed from one continuous stretch of amino acids 18). The GtfB protein has a similar organization, except that the C-terminal part of domain V is absent. The amino acid residue numbers indicate the start and end of each domain. (B) SDS-PAGE (8%, Coomassie blue stained) analysis of GtfB and GtfB-ΔN expression. Lane 1, whole cell extract of E. coli with pET15b-gtfB plasmid; lane 2, purified GtfB (predicted Mr: 179 kDa); Lane 3, whole cell extract of E. coli with pET15b-gtfB-ΔN plasmid; lane 4, purified GtfB-ΔN (predicted Mw = 99 kDa); M, marker proteins.

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transferase activity compared to GtfB-∆N, thereby initially generating more free glucose. In the reactions of both GtfB-∆N and GtfW-∆N with maltodextrins that possess branches (Fig. 2a2 and 2b2), lower conversions (36% for GtfB-∆N and 32% for GtfW-∆N) of (α1→4) linkages into (α1→6) linkages were obtained compared to the reactions using amylose V (90% for GtfB-∆N and 74% for GtfW-∆N) as substrate. Long chain amylose thus appears to be a more efficient substrate for conversion of (α1→4) into (α1→6) linkages, and was therefore used as the standard substrate in subsequent activity assay development and further biochemical analysis.

Measurement of the hydrolysis activity of 4,6-α-GTase enzymes

To characterize 4,6-α-GTase enzymes biochemically, assays to determine both their hydrolysis activity (H) and transferase activity (T) are required. All three characterized 4,6-α-GTases synthesize most of all (α1→6) linkages and occasionally an (α1→4) linkage (Fig. 3) (18). Whereas the 4,6-α-GTase transferase activity is dominant, hydrolysis activity also occurs resulting in glucose release (Fig. 3). Reducing power and glucose measurements are widely accepted methods to determine H of starch converting enzymes with either endo- or exo-hydrolyzing activity, such as α-amylase, α-glucosidase, and cyclodextrin glucanotransferase (33-35). In case of 4,6-α-GTases, the measurement of free glucose specifically may lead to an underestimation of H. The reducing power assay representing both free glucose and products of the transferase reaction with glucose as acceptor substrate, may give a more precise estimate of H (Fig. 3). Therefore the Nelson-Somogyi assay measuring reducing power was compared with the GOPOD assay measuring free glucose; the results were the same in the initial stage of the reaction (Fig. S1). Therefore, both assays are equally applicable for measuring the initial rates of hydrolysis activity of 4,6-α-GTase enzymes.

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Fig 2. One-dimensional 1H NMR spectra (D2O, 335 K) of the products derived from amylose V and maltodextrins via L. reuteri 121 GtfB-∆N (a1 and a2, resp.) and L. reuteri DSM 20016 GtfW-∆N (b1 and b2, resp.) treatments.

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Fig. 3. Schematic diagram of the reactions catalyzed by 4,6-α-GTase as proposed by Leemhuis et al. with minor modifications (18). Reducing ends of sugar molecules are shown in light grey color. The molecules with grey frames can only be used as acceptor substrates. The grey frame with light grey fill indicates free glucose.

Measurement of the Total Activity of 4,6-α-GTases

The total activity of 4,6-α-GTases, which includes hydrolysis H and transferase activity T, was estimated with amylose as substrate, using the iodine staining assay. This assay is based on the fact that iodine stains amylose but not linear (1→6)-α-D-glucan, as can be predicted by the theory of steric hindrance; the

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corresponding molecular simulations are shown in Fig. 4. The diameter of the iodine atom is 4.2 Å measured by Raman spectroscopy (36). The linear (1→6)-α-D-glucan model is based on the results of Genin et al (37), who showed that five-glucose-units per rotation in a right-handed helix is the most stable configuration. The inner diameter of such a helix is around 4.1 Å, therefore the iodine anions cannot be trapped into the cavities of the (1→6)-α-D-glucan due to the steric hindrance. We experimentally verified that dextran does not stain with iodine, using 0.5% (w/v) dextran T10 (Mw: 9-11 kDa, Holbaek, Denmark) instead of 0.5% (w/v) amylose V in the iodine staining assay as described in Methods (data not shown). The iodine anions can be entrapped into amylose because of the larger inner diameter (13 Å) and a shorter pitch (8 Å) of each helix (38). H reduces the iodine binding capacity of the amylose substrate since glucose is released gradually from the non-reducing end (Fig. 4) (15). T also impairs the iodine binding capacity of amylose V because it converts the α1→4 linked chains at the non-reducing end into α1→6 linked chains, lacking iodine staining capacity. The iodine binding capacity of the amylose V substrate decreased linearly in time during its reaction with GtfB-∆N (Fig. 5).

Both H and T thus influence the iodine binding capacity of the amylose substrate. Therefore, the iodine assay is applicable for measuring the total activity of 4,6-α-GTases.

Effects of pH, temperature and metal ions on activity of GtfB-∆N and GtfW-

∆N

The pH optimum for H and T of both GtfB-∆N and GtfW-∆N with amylose V was pH 5.0 (Fig. 6). The temperature optimum for H and T of both enzymes at pH 5.0 was 55 °C. However, at temperatures of 50 and 55 °C, the half-life of both enzymes is less than 10 min (18, 20). At 40 °C, both enzymes were stable for more than 2 h. At lower temperature, the activities of both enzymes remained relatively high (Fig. 6a and b). The transferase activity does not change significantly in the range of 40 to 55 °C. Therefore, pH 5.0 and 40 °C are the optimal conditions for both enzymes, resulting in a ratio T/H=4.3:1 for GtfB-∆N and 1.7:1 for GtfW-∆N.

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Fig. 4. (A) Schematic diagrams of amylose and α(1→6)-glucan, and (B) the complexes of iodine with amylose and with the amylose derived products of the 4,6-α-GTase activities, hydrolysis (H) and transglycosylation (T). The round atoms represent iodine ions.

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Fig. 5. Total activity of the L. reuteri 121 GtfB-∆N (60, 90 or 120 nM) enzyme with amylose V (0.25%, w/v) was followed in time by taking samples and measuring the formation of the iodine-amylose V complex. Resulting total activities (U/mg) are indicated.

Aiming to enhance the T/H ratio with the amylose V substrate, most important for industrial application of these enzymes, further variations in incubation temperature and pH were investigated. When the incubation temperature of GtfB-∆N was decreased from 55 to 30 °C, 82% of T and 48% of H remained, resulting in the highest T/H value of 6.0:1 (Fig. 6a). Such higher values of T/H thus result in lower yields of glucose and short-chain oligosaccharides, formed from glucose as acceptor substrate. Rather, more long-chain polysaccharides (IMMPs) are synthesized which are considered as soluble dietary fiber. Using the optimal conditions described above (pH 5.0 and 40 °C), the H, T and total activities of both GtfB-∆N (0.6, 2.2, and 2.8 U/mg) and GtfW-∆N (1.6, 2.3, and 3.9 U/mg) were determined. Compared to GtfB-∆N, GtfW-∆N has relatively high H activity, which is in agreement with the NMR results showing more products with reducing ends formed by GtfW-∆N (Fig. 2).

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Fig. 6. The effects of pH and temperature on L. reuteri 121 GtfB-∆N (a) and L. reuteri DSM20016 GtfW-∆N (b) activities including hydrolysis, transferase and total activities, using amylose V (0.25%, w/v) as substrate in the presence of 1 mM CaCl2. The experiments were done in triplicate. For pH optimization, values of H, T and total activities of both GtfB-∆N (0.6, 2.2, and 2.8 U/mg) and GtfW-∆N (1.6, 2.3, and 3.9 U/mg) determined at pH 5.0 and 40 °C were set as 100%. For temperature optimization, values of H, T and total activities of both GtfB-∆N (0.6, 2.3, and 2.9 U/mg) and GtfW-∆N (1.7, 2.4, and 4.1 U/mg) determined at pH 5.0 and 55 °C were set as 100%.

Both 4,6-α-GTases and glucansucrases (GSes) belong to GH70 and share high protein sequence similarity. GSes are strongly ion-dependent and need Ca2+ to stimulate their transferase activity; for example, in the presence of Ca2+ ions, transferase activity of reuteransucrase GtfA of L. reuteri 121 was enhanced 8 fold (24). In the present study, different metal ions had varying effects on the hydrolysis and transferase activities of 4,6-α-GTase. As shown in Table 1, Cu2+, Fe2+ and Fe3+ ions strongly inhibited both activities of GtfB-∆N and GtfW-∆N. In the presence of Cu2+ ions, no transferase activities were detectable for both enzymes. K+, Na+, and Mg2+ ions had virtual no effect on these activities, especially not on transferase activity, whereas Ca2+ and Mn2+ ions slightly activated both activities of GtfB-∆N and GtfW-∆N.

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Table 1. Effects of various compounds on GtfB-∆N and GtfW-∆N activity. Incubations were performed at 40 °C in a 25 mM NaAc buffer (pH 5.0) with amylose V (0.25%, w/v) as substrate. The hydrolysis activity without additional compound was set at 100% (0.6 U/mg for GtfB-∆N and 1.3 U/mg for GtfW-∆N). ND = not detectable. The experiments were done in duplicate.

Compounds (1 mM)

GtfB-∆N GtfW-∆N Hydrolysis activity (%)

Transferase activity (%)

Hydrolysis activity (%)

Transferase activity (%)

None 100 ± 4 405 ± 35 100 ± 8 180 ± 16 EDTA 75 ± 5 388 ± 51 90 ± 7 143 ± 14 NaCl 118 ± 12 404 ± 22 108 ± 11 171 ± 13 CaCl2 111 ± 19 422 ± 65 121 ± 23 206 ± 33 MnCl2 125± 15 425 ± 71 137 ± 16 223 ± 23 MgCl2 94 ± 12 378 ± 5 108 ± 12 198 ± 18 KCl 86± 10 435 ± 6 109 ± 11 183 ± 10

CuCl2 51 ± 5 ND 35 ± 7 ND FeCl2 50 ± 6 175 ± 42 64 ± 4 74 ± 15 FeCl3 35 ± 5 85 ± 25 46 ± 6 32 ± 10

Kinetic analysis and acceptor specificity of GtfB-∆N

With amylose V as a substrate, GtfB-∆N displayed Michaelis-Menten type kinetics for hydrolysis, transferase, and total enzyme activity. The kinetic parameters of GtfB-∆N were calculated from plots of 1/[V] versus 1/[S]. The turnover rate (kcat) of the transferase reaction is about 4 times higher than that of the hydrolysis reaction (Table 2). The kcat/Km value for the transferase reaction is 3 fold higher than for the hydrolysis reaction, indicating that the transferase activity is predominant, resulting mostly in synthesis of modified amylose (IMMP) and in low glucose release.

Table 2. Kinetic parameters of the L. reuteri 121 GtfB-∆N enzyme with the amylose V substrate. A fixed amount of GtfB-∆N enzyme (60 nM) was used with varying concentrations of amylose V (0.025-0.4%, w/v) at pH 5.0 and 40 °C.

Km (g·L-1) kcat (s-1) kcat / Km (g-1·s-1·L)

Hydrolytic activity 0.50 (±0.04) 9.06±0.18 18.5 (±1.8)

Transferase activity 0.69 (±0.07) 36.22±1.14 53.0 (±7.2)

Total Activity 0.64 (±0.02) 45.04±1.14 70.6 (±4.0)

A molecular mass of 99 kDa (GtfB-∆N) was used in the calculation of the kcat.

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To study the acceptor substrate specificity of GtfB-∆N, the total activities with amylose V in the presence/absence of different acceptor substrates were determined (Table 3). The transglycosylation factor (TF), defined as the ratio of the amylose degradation ratio in the presence/absence of small acceptor substrates was measured. TF reflects the suitability of a certain acceptor substrate for its enzyme (39-41). Addition of these small acceptor substrates increased the reaction rates of GtfB-∆N with amylose V in all cases. The TFs for α1→4 linked maltose and glucose as acceptor substrates were higher than 1 and similar. The TFs for the α1→6 linked acceptor substrates isomalt(tri)ose clearly were much higher, indicating that isomalt(tri)ose are strong acceptor substrates for the GtfB-∆N enzyme (Table 3).

Table 3. Amylose V (0.25%, w/v) degradation activity (total activity, measured using iodine staining assay) and transglycosylation factors of the L. reuteri 121 GtfB-∆N enzyme in the presence of different acceptor substrates (10 mM). Experiments are carried out in duplicate and average data was described.

Amylose degradation ( U/mg)

Transglycosylation factora

Without acceptors 2.8 1.0

Glucose 6.4 2.2

Maltose 7.3 2.6

Maltotriose 7.5 2.7

Isomaltose 13.5 4.7

Isomaltotriose 19.0 6.7

a The transglycosylation factor is defined as the ratio of amylose degradation rates in the presence/absence of an acceptor substrate.

GtfB is functional at higher substrate concentrations

To explore the effects of substrate concentration on the functionality of GtfB, the enzyme was incubated at various Paselli MD6 maltodextrins (partial hydrolysis product of potato starch with a dextrose equivalent between 5 and 7 and 2.9% α1→6 glycosidic linkages) concentrations keeping the substrate to enzyme ratio at 1000:1 on a mass basis. The results show that GtfB is fully functional at high substrate concentrations. GtfB introduced α1→6 glycosidic linkages at all substrate concentrations, though most efficiently at concentrations between 5 and 30% Paselli MD6 with a maximum of 42.2% α1→6 glycosidic linkages at 30%

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concentration (Fig. 7). Moreover, at high substrate concentrations the enzyme has basically no hydrolytic activity, as above 15% (w/v) substrate there is no increase in reducing power compared to the maltodextrins substrate (3.9% reducing power) used.

Fig. 7. The Paselli MD6 substrate (0.93%-55.8%, w/v) was incubated with Lactobacillus reuterii 121 GtfB-∆N enzyme (1-60 µg/ml). The reducing power of the samples and the percentages of α1→6 glycosidic linkages in the products were determined by 1H NMR. Reactions were performed for 72 h, incubated at 37°C and pH 5.0.

Discussion The amino acid sequences of 4,6-α-GTases are highly similar to those of glucansucrases, and they constitute a subfamily of GH70. Most of the glucansucrases characterized from Lactobacilli have relatively large N-terminal variable regions, possibly involved in cell wall binding, but a clear functional role has not been established (28). Previous research showed that truncation of the N-terminal variable region of some glucansucrases had no effect on the size and linkage-type distribution of the products formed, but resulted in a much higher expression level in E. coli (24-26, 42). The purified truncated Gtf180-∆N and GtfA-∆N proteins also yielded crystals suitable for X-ray diffraction (29-31). The subsequent elucidation of high resolution 3D structures of truncated

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glucansucrase proteins revealed that their peptide chains follow a “U”-path with multiple domains (e.g. glucansucrase Gtf180) (29, 31, 43). Multiple sequence alignment of 4,6-α-GTases with glucansucrases showed that they also have a large N-terminal variable domain (data not shown). In GtfB, this N-terminal domain is formed by residues 1-733. Notably, GtfB differs from glucansucrases in lacking the C-terminal polypeptide segment of domain V. Applying a similar strategy as with the glucansucrase, the N-terminal truncation of GtfB (GtfB-∆N) resulted in a significant enhanced expression level in E. coli compared to the full length wild type GtfB without affecting the activity of the enzyme and products formed.

To biochemically characterize this novel group of 4,6-α-GTase enzymes, several assays have been used, such as HPAEC and quantitative TLC measurement of the saccharides formed from maltoheptaose, or the glucose released from maltose as substrate (16, 18). Both assays have limitations; e.g. they do not allow determination of the specific hydrolysis and transferase activities of these 4,6-α-GTases, nor a kinetic analysis. Dextran dextrinases (DDases) from Gluconobacter oxydans also catalyze the cleavage of an α(1→4) linked glucosyl unit from the non-reducing end of a donor substrate and its subsequent transfer to an acceptor substrate forming an α(1→6) linkage (44). The amino acid sequences of these DDase enzymes have not been annotated yet, and further comparison with 4,6-α-GTases are not possible yet. The dinitrosalicylic acid assay based on changes in reducing power, and a viscosity build-up assay based on the changes of rheological properties from maltodextrin to dextran have been used for estimating the activity of DDases (45). However, the poor sensitivity of dinitrosalicylic acid and interference by unreacted maltodextrins, and the complex non-Newtonian and time-dependent flow behavior of the maltodextrin/dextran mixture in the reaction mixture severely limits the use of this method. Later, a more reliable assay based on discrete transglycosylation reactions was successfully applied, using maltose as a standard substrate (44). The generated panose was demonstrated to be a valid indicator for transglycosylation activity. A similar assay using p-nitrophenyl-α-D-glucopyranoside (NPG) instead of maltose also provided a reliable estimate of DDase transglycosylation activity. However, these assays are also not applicable

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for 4,6-α-GTases because of their low or undetectable activity with maltose or NPG. Thus, proper assays for 4,6-α-GTase activities remained to be established.

Starch-converting enzyme activities can be determined with various methods by following amylose degradation/modification (34, 46). Also 4-α-glucanotransferase enzymes that catalyze both intermolecular (disproportionation) and intramolecular (cyclization) reactions can be studied in this way (39, 47). As shown in our study, amylose also is an efficient substrate for the 4,6-α-GTase catalyzed transferase and hydrolysis reactions. The data shows that the iodine-amylose staining assay is a suitable method to estimate the total activity of 4,6-α-GTases, whereas the GOPOD assay serves to determine hydrolysis (H) activity. The data from both assays can be combined calculate transferase (T) activity. Ratios T/H=4.3:1 for GtfB-∆N and 1.7:1 for GtfW-∆N were obtained under the optimal conditions for both enzymes, indicating that GtfB is less hydrolytic than the GtfW enzyme. Under suboptimal temperature conditions the T/H ratio for GtfB even reached 6.0:1. The data clearly shows that GtfB is a proper transglycosylase, with relatively minor hydrolysis activity. T/H ratios have been reported for other transglycosylase enzymes. For glucansucrase Gtf180-∆N from L. reuteri 180 and glucansucrase GtfA-∆N from L. reuteri 121, these ratios are 2.4:1 and 0.6:1 (30, 48); For cyclodextrin glucanotransferase of Thermoanaerobacterium thermosulfurigenes this ratio even is 8.1:1 (49).

Determination of the total activity of GtfB with amylose V in the presence/absence of acceptor substrates clearly showed that isomaltodextrins are preferred over maltodextrins as acceptor substrates. Such information strongly contributes to our understanding of the transglycosylation reaction catalyzed by 4,6-α-GTase enzymes, especially in combination with detailed 3D structural information allowing to identify and characterize acceptor binding sites. To this end we are currently aiming to crystallize constructs of GtfB and its homologues (in preparation).

To conclude, N-terminal truncation of the L. reuteri GtfB enzyme (GtfB-∆N) remarkably enhanced the soluble expression level while retaining full activity. Using amylose V as substrate, the GOPOD assay and the amylose-iodine staining method allowed the characterization of the new 4,6-α-GTase enzymes GtfB and GtfW, including their optimal reaction conditions, hydrolysis and transferase

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activities. The data shows that GtfB is less hydrolytic than GtfW, which is in agreement with the NMR results. Using these assays, various properties of GtfB, the best characterized 4,6-α-GTase enzyme, were determined including its kinetic parameters and acceptor substrate specificity. The GtfB enzyme remained active at high substrate concentrations resulting in high product conversions, a promising feature for industrial applications. Moreover at high substrate concentrations the hydrolytic activity of GtfB is virtually absent. The developed assays thus provide a firm basis for the future characterization of other, natural and engineered, 4,6-α-GTase enzymes.

Acknowledgements The study was financially supported by the Chinese Scholarship Council (to YB), the University of Groningen and by the TKI Agri&Food program as coordinated by the Carbohydrate Competence Center (CCC-ABC; www.cccresearch.nl).

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9. Guraya HS, James C, Champagne ET. 2001. Effect of enzyme concentration and storage temperature on the formation of slowly digestible starch from cooked debranched rice starch. Starch-Stärke 53:131-139.

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19. Dijkhuizen L, van der Maarel MJEC, Kamerling JP, Leemhuis H, Kralj S, JM D. 2010. Gluco-oligosaccharides comprising (α1→4) and (α1→6) glycosidic bonds, use thereof, and methods for providing them. WO/2010/128859.

20. Bai Y, van der Kaaij RM, Woortman AJJ, Jin Z, Dijkhuizen L. 2015. Characterization of the 4,6-α-glucanotransferase GTFB enzyme of Lactobacillus reuteri 121 isolated from inclusion bodies. BMC Biotechnology 15:49.

21. Palomo M, Pijning T, Booiman T, Dobruchowska JM, van der Vlist J, Kralj S, Planas A, Loos K, Kamerling JP, Dijkstra BW, van der Maarel MJEC, Dijkhuizen L, Leemhuis H. 2011. Thermus thermophilus glycoside hydrolase family 57 branching enzyme crystal structure, mechanism of action, and products formed. J Biol Chem 286:3520-3530. 22. Cochran B, Lunday D, Miskevich F. 2008. Kinetic analysis of amylase using quantitative Benedict's and iodine starch reagents. J Chem Educ 85:401-403. 23. Green F, Clausen CA, Highley TL. 1989. Adaptation of the Nelson-Somogyi reducing-sugar assay to a microassay using microtiter plates. Anal Biochem 182:197-199.

24. Kralj S, van Geel-Schutten GH, van der Maarel MJEC, Dijkhuizen L. 2004. Biochemical and molecular characterization of Lactobacillus reuteri 121 reuteransucrase. Microbiol-SGM 150:2099-2112.

25. Monchois V, Arguello-Morales M, Russell RRB. 1999. Isolation of an active catalytic core of Streptococcus downei MFe28 GTF-I glucosyltransferase. J Bacteriol 181:2290-2292.

26. Swistowska AM, Wittrock S, Collisi W, Hofer B. 2008. Heterologous hyper-expression of a glucansucrase-type glycosyltransferase gene. Appl Microbiol Biotechnol 79:255-261.

27. Leemhuis H, Pijning T, Dobruchowska JM, van Leeuwen SS, Kralj S, Dijkstra BW, Dijkhuizen L. 2013. Glucansucrases: Three-dimensional structures, reactions, mechanism, α-glucan analysis and their implications in biotechnology and food applications. J Biotechnol 163:250-272.

28. Kralj S, van Geel-Schutten GH, Dondorff MMG, Kirsanovs S, van der Maarel MJEC, Dijkhuizen L. 2004. Glucan synthesis in the genus Lactobacillus: isolation and characterization of glucansucrase genes, enzymes and glucan products from six different strains. Microbiol-SGM 150:3681-3690. 29. Pijning T, Vujicic-Zagar A, Kralj S, Dijkhuizen L, Dijkstra BW. 2012. Structure of the α -1,6/α -1,4-specific glucansucrase GTFA from Lactobacillus reuteri 121. Acta Crystallogr F 68:1448-1454.

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30. Pijning T, Vujicic-Zagar A, Kralj S, Eeuwema W, Dijkhuizen L, Dijkstra BW. 2008. Biochemical and crystallographic characterization of a glucansucrase from Lactobacillus reuteri 180. Biocatal Biotransfor 26:12-17.

31. Vujičić-Žagar A, Pijning T, Kralj S, Lopez CA, Eeuwema W, Dijkhuizen L, Dijkstra BW. 2010. Crystal structure of a 117 kDa glucansucrase fragment provides insight into evolution and product specificity of GH70 enzymes. Proc Natl Acad Sci USA 107:21406-21411. 32. van Leeuwen SS, Leeflang BR, Gerwig GJ, Kamerling JP. 2008. Development of a H-1 NMR structural-reporter-group concept for the primary structural characterisation of α-D-glucans. Carbohyd Res 343:1114-1119. 33. Aref HL, Mosbah H, Louati H, Said K, Selmi B. 2011. Partial characterization of a novel amylase activity isolated from Tunisian Ficus carica latex. Pharm Biol 49:1158-1166. 34. Xiao Z, Storms R, Tsang A. 2006. A quantitative starch-iodine method for measuring α -amylase and glucoamylase activities. Anal Biochem 351:146-148.

35. Kelly RM, Leemhuis H, Dijkhuizen L. 2007. Conversion of a cyclodextrin glucanotransferase into an α-amylase: assessment of directed evolution strategies. Biochemistry-US 46:11216-11222.

36. Wang DD, Guo WH, Hu JM, Liu F, Chen LS, Du SW, Tang ZK. 2013. Estimating atomic sizes with raman spectroscopy. Sci Rep-UK 3:1-5. 37. Genin AL, Banatskaya MI, Nikitin IV, Skokova IF, Vainshtein EF, Kushnerev MY, Kozlov PV. 1983. Criteria of choice of helix type in crystalline dextrane. Polym Sci U.S.S.R. 25:1717-1722. 38. Yu XC, Houtman C, Atalla RH. 1996. The complex of amylose and iodine. Carbohyd Res 292:129-141. 39. Jung JH, Jung TY, Seo DH, Yoon SM, Choi HC, Park BC, Park CS, Woo EJ. 2011. Structural and functional analysis of substrate recognition by the 250s loop in amylomaltase from Thermus brockianus. Proteins 79:633-644. 40. Tang SY, Yang SJ, Cha HJ, Woo EJ, Park C, Park KH. 2006. Contribution of W229 to the transglycosylation activity of 4-α-glucanotransferase from Pyrococcus furiosus. Biochim Biophys Acta 1764:1633-1638. 41. Tachibana Y, Takaha T, Fujiwara S, Takagi M, Imanaka T. 2000. Acceptor specificity of 4-α-glucanotransferase from Pyrococcus kodakaraensis KOD1, and synthesis of cycloamylose. J Biosci Bioeng 90:406-409. 42. Moulis C, Arcache A, Escalier PC, Rinaudo M, Monsan P, Remaud-Simeon M, Potocki-Veronese G. 2006. High-level production and purification of a fully active

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recombinant dextransucrase from Leuconostoc mesenteroides NRRL B-512F. FEMS Microbiol Lett 261:203-210.

43. Ito K, Ito S, Shimamura T, Weyand S, Kawarasaki Y, Misaka T, Abe K, Kobayashi T, Cameron AD, Iwata S. 2011. Crystal structure of glucansucrase from the dental caries pathogen Streptococcus mutans. J Mol Biol 408:177-186.

44. Naessens M, Cerdobbel A, Soetaert W, Vandamme EJ. 2005. Dextran dextrinase and dextran of Gluconobacter oxydans. J Ind Microbiol Biot 32:323-334. 45. Naessens M, Vandamme E. 2001. Transglucosylation and hydrolysis activity of Glucanobacter oxydans dextran dextrinase with several donor and acceptor substrates, p 195-203, Biorelated polymers: sustainable polymerscience and technology. Kluwer/Plenum, London.

46. Cech J, Friedecky B. 1972. Determination of α amylase by a modification of photometric iodine starch method. Cas Lek Cesk 111:309-311. 47. Terada Y, Fujii K, Takaha T, Okada S. 1999. Thermus aquaticus ATCC 33923 amylomaltase gene cloning and expression and enzyme characterization: Production of cycloamylose. Appl Environ Microbiol 65:910-915. 48. Kralj S, Stripling E, Sanders P, van Geel-Schutten GH, Dijkhuizen L. 2005. Highly hydrolytic reuteransucrase from probiotic Lactobacillus reuteri strain ATCC 55730. Appl Environ Microbiol 71:3942-3950. 49. Kelly RM, Leemhuis H, Rozeboom HJ, van Oosterwijk N, Dijkstra BW, Dijkhuizen L. 2008. Elimination of competing hydrolysis and coupling side reactions of a cyclodextrin glucanotransferase by directed evolution. Biochem J 413:517-525.

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Fig. S1. The glucose and reducing sugar released in 150 min from reactions catalyzed by 60 nM of the L. reuteri 121 GtfB-∆N enzyme with 0.25% (w/v) amylose V as substrate at 40 °C and pH 5.0.

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Chapter 4

Lactobacillus reuteri strains convert starch and maltodextrins

into homo-exopolysaccharides using a cell-associated 4,6-α-

glucanotransferase enzyme

Yuxiang Bai1,2, Markus Böger1, Rachel Maria van der Kaaij1, Albert Jan Jacob

Woortman3, Tjaard Pijning4, Sander Sebastiaan van Leeuwen1, Alicia Lammerts

van Bueren1, Lubbert Dijkhuizen1,*

1 Microbial Physiology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, The Netherlands

2 The State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi 214122, China

3 Department of Polymer Chemistry, Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 4, 9747 AG Groningen, The Netherlands

4Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, The Netherlands

In preparation for submission

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Abstract Exopolysaccharides (EPS) of lactic acid bacteria (LAB) are of interest for food applications. LAB use sucrose as substrate for α-glucan synthesis by extracellular glucansucrase enzymes. Various Lactobacillus reuteri strains also possess 4,6-α-glucanotransferase (4,6-α-GTase) enzymes. Purified 4,6-α-GTases were shown to act on starches (hydrolysates), cleaving α1→4 linkages and synthesizing α1→6 linkages, yielding isomalto-/malto- polysaccharides (IMMP). Here we report that also L. reuteri cells with these extracellular, cell-associated 4,6-α-GTases synthesize EPS (α-glucan) from starches (hydrolysate). NMR, SEC, and enzymatic hydrolysis of EPS synthesized by L. reuteri 121 cells showed that these have similar linkage specificities but generally are much bigger in size than IMMP produced by 4,6-α-GTase GtfB enzyme. The various IMMP-like EPS are efficiently used as growth substrate by probiotic Bifidobacterium strains possessing amylopullulanase activity. These IMMP-like EPS thus have potential prebiotic activity and may contribute to the application of probiotic L. reuteri strains grown on maltodextrins or starches as synbiotics.

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Introduction Lactic acid bacteria (LAB) are a group of Gram-positive, non-sporulating bacteria, of which Lactobacillus and Bifidobacterium strains are regarded as probiotics. Some LAB are “Generally Recognized As Safe” (GRAS) strains and have been used in fermented foods such as cheese and yoghurt since ancient time. LAB produce lactic acid, diacetyl/acetoin, and carbon dioxide that contribute to the texture, flavor and shelf life of fermented foods (1-3). Some LAB also produce exopolysaccharides (EPS) which find application in the improvement of food texture. They also may have interesting health effects for humans, e.g. antitumor activity and immune stimulation (4,5).

Depending on their composition and mechanism of biosynthesis, EPS produced by LAB can be classified into two groups: hetero-EPS and homo-EPS (6). Hetero-EPS consist of more than one carbohydrate moieties, and generally are synthesized intracellularly from activated sugars (7). LAB homo-EPS consist of a single type of monosaccharide, generally D-glucose or D-fructose, and can be divided into α-glucans (e.g. dextran, mutan, alternan and reuteran) and β-fructans (e.g. inulin and levan) (8-10). Most LAB homo-EPS known are produced from sucrose by extracellular glucansucrase and fructansucrase enzymes (6, 11, 12). Lactobacillus reuteri 121 uses the GtfA glucansucrase to convert sucrose into an α-glucan (reuteran) with both α1→4 and α1→6 linkages, mostly alternating (9, 13-15). Glucansucrases are glycoside hydrolase enzymes of family GH70 (www.CAZy.org).

Compared to sucrose, starch is much more abundant and is a cheaper carbon source for LAB industrial cultivation and product formation (16). In the last 30 years, a number of amylolytic LAB strains have been isolated and their starch-acting enzymes investigated (17, 18). Most of these are starch-hydrolyzing enzymes and only few enzymes with starch transglycosylation activity have been characterized (19-24). Recently, various 4,6-α-glucanotransferase (4,6-α-GTase) enzymes have been characterized from L. reuteri strains (25, 26). These transglycosylating enzymes constitute a GH70 subfamily. These 4,6-α-GTases produce isomalto-/malto- polysaccharides (IMMP), which is a new type of soluble dietary fiber, using starch or starch hydrolysates as substrate rather than sucrose (27, 28). The L. reuteri 121 strain GtfB enzyme has been purified and its

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IMMP products have been characterized in detail (27, 28). The gtfA and gtfB genes in L. reuteri 121 are located next to each other in the genome (26, 29).

In this study, 4 Lactobacillus reuteri strains possessing 4,6-α-GTase genes are shown to produce homo-EPS from maltodextrins in vivo. The model strain L. reuteri 121 with the gtfB gene was found to produce homo-EPS from starch or starch hydrolysates. The EPS products of L. reuteri 121 from different types of starch were further investigated with respect to structures and sizes, and compared to the IMMP produced by the GtfB enzyme in vitro. Finally, the L. reuteri 121 EPS derived from different maltodextrins as well as from sucrose (reuteran) were tested as carbon source for growth of various probiotic Bifidobacterium strains in order to evaluate their potential prebiotic activity.

Materials and Methods

Growth conditions

Lactobacillus reuteri strains, including L. reuteri 121 (LMG 18388), L. reuteri DSM20016, L. reuteri TMW1.106 (kindly provided by Prof. Rudi F. Vogel in Technische Universität München) (30), L. reuteri ML1 (LMG 20347), L. reuteri 180 (LMG 18389), and L. reuteri ATCC5573 were subcultured from stocks stored at -80 °C in 10 ml of sugar-free MRS medium (liquid culture or agar plate) supplemented with 1% glucose. The fresh cultures were inoculated anaerobically in modified sugar-free MRS medium with different carbon sources with or without 1% glucose (w/v). One liter medium contained 10 g bactopeptone, 4 g yeast extract, 5 g sodium acetate, 2 g tri-ammonium citrate, 0.2 g MgSO4·7H2O, 0.05 g MnSO4·7H2O; and 1 ml Tween 80, supplemented with 5 g starch or 50 g maltodextrins. For agar medium, 15 g/l agar was added. The native starches, maltodextrins and amylose V were provided by AVEBE (Veendam, The Netherlands) (Table 1).

The Bifidobacterium strains B. breve DSM20091, B. adolescentis DSM 20083 and B. dentium DSM20436 were subcultured from stocks stored at -80 °C in 5 ml of Bifidobacterium medium (BM) supplemented with 1% glucose (31). One liter BM contained 10 g trypticase peptone, 2.5 g yeast extract, 3 g tryptose, 3 g K2HPO4, 3 g KH2PO4, 2 g triammonium citrate, 0.3 g pyruvic acid, 1 ml Tween

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80, 0.574 g MgSO4·7H2O, 0.12 g MnSO4·7H2O in 1 l distilled water), 5 g NaCl. After autoclaving BM was supplemented with 0.05% (w/v) filter-sterilized cysteine-HCl, and strains were grown at 37 °C under anaerobic conditions maintained by GasPak EZ anaerobe container system (BD, New Jersey, US). Aliquots (1%) from overnight cultures were inoculated into 5 ml of BM supplemented with different EPS (0.5%, w/v). BM without an added carbon source and BM with 0.5% (w/v) glucose were used as negative and positive controls, respectively.

Extraction of exopolysaccharides (EPS)

EPS were produced by L. reuteri strains in the sugar-free MRS medium supplemented with various maltodextrins (5.0%, w/v) or starch (0.5%, w/v). Reuteran was produced by L. reuteri 35-5, a mutant derived from L. reuteri 121, using the sugar-free MRS medium supplemented with sucrose (5.0%, w/v) (32). Strains were grown anaerobically at 37 °C for 72 h. EPS were extracted as described below.

For cultures grown with maltodextrins, EPS were extracted from supernatants obtained by centrifugation of liquid MRS cultures (33). Two volumes of cold ethanol (-20 °C) were added to supernatants and held at 4 °C overnight. Precipitates were harvested by centrifugation (12 000 ×g, 30 min, 4 °C) and re-dissolved in 1 volume of ddH2O. EPS were re-precipitated with 2 volumes of ethanol at 4 °C overnight. After centrifugation, the harvested precipitates were dissolved in water, and then dialyzed (10 kDa MWCO, Thermo Scientific) against water for 48 h with changes of ddH2O every 12 h (34).

For cultures grown with starch, the EPS were collected according to a modified method based on the protocol of Lee et al. (35). Total EPS including soluble and insoluble EPS, and cells were precipitated with 2 volumes of cold ethanol (-20 °C) at 4 °C overnight. Samples collected from tubes were centrifuged at 10000 ×g for 10 min. The polysaccharides were solubilized by addition of 10 ml 2 M NaOH. The cells were separated by centrifugation at 10000 ×g for 15 min. EPS present in the supernatant was neutralized by the addition of an appropriate amount of 2 M HCl (36). The ethanol precipitation procedure described above was used to further purify EPS.

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Monosaccharide analysis

EPS samples (0.5 mg) were dissolved in 200 µl Milli-Q water in a glass reaction tube. An equal volume of 4 M trifluoroacetic acid (TFA) was added to the sample. The sample was hydrolyzed for 4 h at 100 °C. Samples were dried under a flow of dry nitrogen. The dry samples were dissolved in 100 µl isopropanol and dried again by evaporation under dry nitrogen. Samples were dissolved in 1 ml dimethyl sulfoxide (DMSO) and used for HPAEC analysis.

Expression of truncated GtfB-∆N, full-length GtfA and full length GtfB

Truncated GtfB-∆N and full-length GtfA were produced as described before (37, 38). The full-length GtfB protein was produced in E. coli BL21 Star (DE3) carrying the plasmids pRSF-GtfB and pBAD22-GroELS as described previously (27). All three enzymes were first purified by His-tag affinity chromatography using a 1-ml Hitrap IMAC HP column (GE Healthcare) followed by anion exchange purification using Hitrap Q FF (GE Healthcare). Protein Purity was checked on 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Protein concentration was measured by Bradford reagent (Bio-Rad, Hercules, CA) using bovine serum albumin as standard.

Activity staining of EPS synthesizing enzymes

After SDS-PAGE, the gels were washed 3 times with dd H2O (30 min each), allowing protein renaturation, and incubated overnight at 37°C in sodium acetate buffer (25 mM, pH 5.0, 1 mM CaCl2) with 5% (w/v) maltodextrins (AVEBE MD20). The activity of EPS synthesizing enzymes in the gel was detected by staining for the polysaccharides by a Periodic Acid-Schiff (PAS) procedure (29). The periodic acid oxidizes the vicinal diols in sugars, creating a pair of aldehydes, which react with the Schiff reagent to give a purple-magenta color. The full-length GtfB expressed in and purified from E.coli was loaded as a reference.

Amylose V conversion by washed L. reuteri 121 cells

The L. reuteri 121 cells were collected by centrifugation (5 000 ×g, 30 min, 4 °C). Cell pellets were washed twice using sodium acetate buffer (25 mM, pH 5.0, 1 mM CaCl2, 0.02% (w/v) NaN3) in order to remove any unbound proteins. The cells were resuspended in the same sodium acetate buffer and were incubated with amylose V (0.25%, w/v) on a shaker at 40 °C to measure enzyme activity

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and to synthesize the enzyme products (37, 39). For amylose V conversion, cells (0.03 U of total GtfB activity per ml reaction volume) with an activity equivalent to that of 60 nM GtfB-∆N enzyme, were incubated with amylose V (0.25%, w/v) on a shaker at 40 °C and pH 5.0 for 24 h.

Preparation of isomalto/malto-polysaccharides (IMMP) with GtfB-∆N

enzyme

IMMP were produced according to Leemhuis et al. (27). Different types of starches and maltodextrins (0.5%, w/v) were incubated with GtfB-∆N enzyme (1.4 U per gram substrate) in 10 mM sodium acetate buffer pH5.0, supplemented with 1 mM CaCl2 and NaN3 (0.02% w/v) at 37 °C. Native starches were pregelatinized by autoclaving (15 min, 121 °C).

Debranching of polysaccharides

Starches, and the IMMP and EPS products derived, were suspended in citrate buffer (50 mM, pH 4.0, 0.02% NaN3) to a final concentration of 10 mg/ml, and slowly heated to 99 °C with a thermomixer (Thermo Fisher Scientific Inc., Waltham, US). The suspension was left at 99 °C for 1 h while shaking. Afterwards, the polymers were debranched at 40 °C for 16 h with 20 U of isoamylase from Pseudomonas sp. (Megazyme, Wicklow, Ireland). The debranched samples were dialyzed (1000 Da MWCO tube, Sigma-Aldrich, US) against Milli-Q and freeze-dried.

NMR analysis

EPS samples produced by the GtfB enzyme in vitro and by L. reuteri 121 cells in vivo were analyzed by NMR according to Leemhuis et al. (27). NMR spectroscopy resolution-enhanced 1D 500-MHz 1H NMR spectra were recorded in D2O on a Varian Inova Spectrometer (NMR Center, University of Groningen) at probe temperatures of 300 K. The samples were exchanged twice with D2O (99.9 atm% D, Cambridge Isotope Laboratories, Inc.) with intermediate lyophilization and then dissolved in 0.6 ml D2O.

Enzymatic hydrolysis

Samples of 1 mg EPS were dissolved in 100 µl citrate buffer (50 mM, pH 3.5-5.0 based on the instructions for these commercial enzymes) and excess amount of

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the following enzymes were added: pullulanase M1 from Klebsiella planticola (Megazyme, Ireland), dextranase from Chaetomium erraticum (Sigma-Aldrich, US), α-amylase from Aspergillus oryzae (Megazyme, Ireland), isoamylase from Pseudomonas sp. (Megazyme, Ireland), and isopullulanase from Aspergillus niger (Megazyme, Ireland), as well as α-amylase plus isoamylase. Samples were incubated for 72 h at 40 °C. The samples were boiled for 10 min to terminate the enzymatic hydrolysis and used for further analysis. The products after α-amylase and isoamylase treatment were fractionated by size-exclusion chromatography on a Bio-Gel P-2 column (100 x 0.9 cm) (Bio-Rad) using ammonium bicarbonate (10 mM) as eluent at a rate of ~10 ml/h and then subjected to NMR analysis.

Size exclusion chromatography (SEC)

DMSO-LiBr (0.05M) was prepared by stirring for 3 h at room temperature followed by degassing for 15 min using an ultrasonic cleaner (Branson 1510, Branson, Danbury, CT). Samples were dissolved at a concentration of 4 mg/ml in DMSO-LiBr by overnight rotation at room temperature, followed by 30 min heating in an oven at 80 °C, obtaining clear sample solutions. The samples were cooled to room temperature and filtered through a 0.45-µm Millex PTFE membrane (Millipore Corporation, Billerica, MA). The SEC system set-up (Agilent Technologies 1260 Infinity) from PSS (Mainz, Germany) consisted of an isocratic pump, auto sampler, an online degasser, an inline 0.2 μm filter, a refractive index detector (G1362A 1260 RID Agilent Technologies), viscometer (ETA-2010 PSS, Mainz) and MALLS (SLD 7000 PSS, Mainz). WinGPC Unity software (PSS, Mainz) was used for data processing. Samples (100 µl) were injected into a PFG guard column using an autosampler at a flow rate of 0.5 ml/min and DMSO-LiBr as eluent. The separation was done by three PFG-SEC columns with porosities of 100, 300 and 4000 Å. The columns were held at 80˚C, the refractive index detector at 45 °C and the viscometer was thermostatted at 60 °C. A standard pullulan kit (PSS, Mainz, Germany) with molecular masses from 342 to 805000 Da was used. The specific RI increment value dn/dc was measured by PSS and is 0.072 (obtained from PSS company).

High-pH anion-exchange chromatography (HPAEC)

The products of GtfB were injected onto a 4×250 nm CarboPac PA-1 column connected to a Dionex DX500 workstation (Dionex). Samples were run with a

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gradient of 30-600 mM NaAc in 100 mM NaOH (1 ml/min), and detected by an ED40 pulsed amperometric detector. A mixture with known concentrations of glucose, isomaltose, isomaltotriose, maltose, panose, maltotriose, maltotetraose, maltopentaose, maltohexaose and maltoheptaose was used as reference.

Thin-layer chromatography (TLC)

The TLC silica gel 60F254 plates (Merck) were run with butanol/acetic acid/water (2:1:1, v/v/v) as solvent. After running for 6 h, TLC plates were developed with 10% (v/v) H2SO4 and 2 g/l orcinol in methanol and then heated by oven at 100 °C for 30 min. Glucose, maltose, maltotriose, maltotetraose, maltopentaose, maltohexaose, and maltoheptaose were used as markers.

Total carbohydrate measurements

Total carbohydrates were measured using the total carbohydrate colorimetric assay kit (Biovision, Milpitas, USA). Of each liquid sample, 5-15 µl was added into a 96-well plate. The total volume was adjusted to 30 µl per well by adding ddH2O. Then, 150 µl of concentrated H2SO4 (98%, v/v) was added per well. The plate was mixed for 1 min on a shaker at room temperature and incubated in an oven at 90 °C for 15 min. Afterwards, 30 µl of developer reagent was added and mixed with sample for 5 min on a shaker at room temperature. The optical density was measured at 490 nm. A calibration curve was made using 0, 4, 8, 12, 16, and 20 µg glucose per well.

Results and Discussion

Lactobacillus reuteri strains produce homo-exopolysaccharides (EPS) from

maltodextrins

Protein blast searches in the NCBI database (http://www.ncbi.nlm.nih.gov/BLAST/) using GtfB of L. reuteri 121 as a query sequence revealed that approx. 28 Lactobacillus strains carry a 4,6-α-GTase encoding gene. Eight out of these 28 are L. reuteri strains. Previously, we have expressed and purified some of these 4,6-α-GTases in E. coli, e.g. GtfB from L. reuteri 121, GtfW from L. reuteri DSM20016, and GtfML4 from L. reuteri ML1, and shown that they produce IMMP from maltodextrins (Fig. 1A)

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(25-27). The question remained whether also in vivo these L. reuteri strains and their genes/enzymes support formation of IMMP-like EPS.

Using maltodextrins 13-17 (dextrose equivalent 13-17, Sigma-Aldrich) as carbon sources for growth, L. reuteri 121 (GtfB), L. reuteri DSM20016 (GtfW), L. reuteri TMW1.106 (Gtf106b), and L. reuteri ML1 (GtfML4), which all possess a 4,6-α-GTase gene, clearly produced ethanol-precipitable EPS, while L. reuteri strains lacking a 4,6-α-GTase gene, such as L. reuteri 180 and L. reuteri ATCC55730, did not. The yields of EPS produced by the L. reuteri DSM20016 (GtfW) and L. reuteri ML1 (GtfML4) strains were relatively low (data not shown), probably because the relatively high hydrolytic activity (39) of their enzymes impaired the formation of polysaccharides that can be precipitated by cold ethanol. These EPS are all composed of glucosyl units (monosaccharide analysis, data not shown) linked by α1→4 and α1→6 linkages (Fig. 1B). Thus, these L. reuteri strains are able to produce α-glucan EPS from maltodextrins, which may be highly relevant for their survival and activities in natural and industrial environments.

4,6-α-GTase enzymes are essential for EPS formation by L. reuteri in vivo

Using the previously established 4,6-α-GTase amylose-iodine assay with amylose V as substrate (39), no activity was detectable in cell-free culture supernatants. However, cell-associated activities were detected in the washed and resuspended cell pellets of all 4 L. reuteri strains with a (putative) 4,6-α-GTase enzyme (Fig. 2A). Amylose V is too large a molecule to pass the L. reuteri cell membrane, therefore intracellular enzymes are not involved here. The combined data indicate that the amylose V modifying enzymes are extracellular and cell-associated and not released into the culture fluid during growth. This is in accordance with the primary structures of 4,6-α-GTases, showing the presence of signal peptides for secretion (40). The 4,6-α-GTases, like glucansucrases, also have variable N-terminal regions. Although the functional role of this N-terminal region has not been established, glucansucrases with this region are always cell wall-bound in lactic acid bacteria (32). The same appears to be true for 4,6-α-GTases.

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Fig. 1. (A) Representation of the IMMP products of 4,6-α-GTase (e.g. GtfB) with maltodextrins (28). (B) NMR spectra of the EPS isolated from L. reuteri 121, L. reuteri DSM20016, L. reuteri ML1, and L. reuteri TMW1.106 cultures grown on maltodextrins AVEBE MD20 (5%, w/v)-MRS medium. To further analyze the functional in vivo role of 4,6-α-GTase, the L. reuteri 121 system was studied in detail as model strain. Periodic Acid-Schiff staining (PAS) is a well-established method that has been used to quickly detect polysaccharides produced by glucansucrase enzymes. Under mild conditions, denatured glucansucrases can be refolded and regain activity in SDS-PAGE gels (29). Previously, we already reported that also the denatured GtfB enzyme can be

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easily renatured in double distilled water (37). PAS staining (Fig. 2B, lane 2) showed that E. coli expressed and purified GtfB protein (~179 kDa) in SDS-PAGE gels produced polysaccharides from maltodextrins. Following SDS-PAGE of L. reuteri 121 total cellular proteins and protein renaturation, incubation with maltodextrins and PAS staining revealed a band, representing an enzyme with a similar molecular mass, (Fig. 2B, lane 1). Previously we reported that L. reuteri 121 GS GtfA (~199 kDa) activity also can be detected by PAS staining of SDS-PAGE gels when using sucrose as substrate (resulting in reuteran formation) (29). GtfA is the only carbohydrate-acting protein in the genome of L. reuteri 121 that has a similar molecular mass as GtfB (Gangoiti et al., in preparation). Unlike the PAS results with sucrose as substrate, no SDS-PAGE band was observed following incubation of E. coli expressed and purified GtfA protein with maltodextrins (Fig. 2B, lane 3). In addition, the resuspended L. reuteri 121 cell pellet was incubated with amylose V (0.25%, w/v) until all substrate was consumed as no color formed when using the iodine-staining assay developed previously (39). NMR analysis revealed that an α-glucan product was formed with 54% of α1→4 linkages and 46% of α1→6 linkages (Fig. 2C). The L. reuteri 121 cells associated 4,6-α-GTase enzyme and the purified GtfB enzyme thus have the same product specificity, cleaving α1→4 linkages and forming α1→6 linkages. The combined data shows that the GtfB enzyme is responsible for EPS formation by L. reuteri 121 cells.

NMR analysis of EPS produced by L. reuteri 121 cells from maltodextins or

starches

Subsequently, we investigated whether L. reuteri 121 cells in liquid cultures also are able to convert starch. Various starches were supplied at a concentration of 5 g/l in order to prevent retrogradation (27). Cultures were supplemented with glucose as carbon and energy source because L. reuteri 121 does not grow on starch alone, while in the case of maltodextrins, no extra glucose (Table 1) was added. The isolated EPS were subjected to NMR analysis. All starches tested were modified during the growth of L. reuteri 121 as is apparent from the increased percentages of α1→6 linkages obtained in all cases (Table 1). Growth of L. reuteri 121 in the presence of starch with a high amylose content and a lower degree of branching resulted in formation of EPS with higher percentages of α1→6 linkages. Highest conversion of α1→4 into α1→6 linkages was

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obtained with wrinkled pea starch, which contains the highest amylose (52.8%, w/w) content. This is in accordance with the in vitro data for starch conversion by GtfB, showing that the amylose content is positively, but the degree of branching is negatively correlated with the percentage of α1→6 linkages in the produced IMMP (27). Compared to these IMMP (27), the L. reuteri 121 EPS derived from the same starch or maltodextrin substrates always have around 0-20% lower percentages of α1→6 linkages (Table 1). The L. reuteri 121 whole cell system provides more complicated reaction conditions, e.g. with pH variations, and the presence of other enzymes and components. Also, in whole cell systems the GtfB enzyme is cell wall-bound. These factors are likely to influence product formation, compared to incubations of the purified free GtfB enzyme with a pure substrate.

Fig. 2. (A) 4,6-α-GTase activity associated with cells of L. reuteri strains using amylose V (0.25%, w/v) as substrate (39). “+” and “-” indicate the presence and absence of activity. (B) Periodic Acid-Schiff (PAS) stained SDS-PAGE gels of whole-cell-extracted proteins of L. reuteri 121 (lane 1), E. coli expressed and purified GtfB (lane 2) and GtfA (lane 3) after incubating with maltodextrins (AVEBE MD20, 5.0%, w/v) overnight at pH 5.0 and 40 °C. (C) 1D 1H NMR analysis of the products (with 54% of α1→4 linkages and 46% of α1→6 linkages) derived from amylose V (0.25%, w/v) incubated with L. reuteri 121 cell pellets. R1α/β represent the reducing –(1→4)-D-Glcp units and D-Glcp units.

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Table 1. Percentages of α1→6 linkages in EPS formed during growth of L. reuteri 121 in MRS-medium with or without 1% of glucose in the presence of different types of starch and maltodextrins, determined by 1H NMR spectroscopy. (n/a, not applicable). The final pH values of the L. reuteri 121 cultures with 1% glucose all were in the range of pH 4.5-4.6.

Substrate

Amylose content

(%) (apparent)

Average DPa

Degree of branching (α1→4,6)

(%)

% (α1→6) in IMMP produced by GtfB enzymed

% (α1→6) in EPS from L. reuteri 121

grown with 1% glucose

% (α1→6) in EPS from L. reuteri

121 grown without 1%

glucose

AVEBE Maltodextrins

Paselli SA2 n/a 50 32 35

Paselli MD6 n/a 20 34 19

AVEBE MD20 n/a 6 25 26

AVEBE SPG30 n/a 4 24 31

AVEBE Starches

Etenia 457 n/a n/a -c 33 22

Arrow root 20.8 n/a - 24 14

Barley 25.5 n/a - 21 15

Corn 29.4 n/a 3.6 21 22

Corn, waxy 0 n/a 4.8 7 11

Mung bean 37.9 n/a - 34 21

Pea, yellow 30.9 n/a - 35 26

Pea, wrinkled 52.8 n/a - 71 52

Potato 36.0 n/a 3.1 28 17

Rice, round grain 25.0 n/a 4.1 13 12

Rice, waxy 0 n/a 4.9 7 11

Sorghum 23.7 n/a - 22 18

Tapioca 23.5 n/a 3.9 20 12

Wheat 28.8 n/a 3.7 22 23

aDegree of polymerization; bn/a, not applicable; cnot available in literature; ddata adapted from Leemhuis et al. (27).

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Size exclusion chromatography analysis of EPS and IMMP derived from

maltodextrins

The sizes of the IMMP products of the GtfB enzyme have not been reported before. Therefore the IMMP-like EPS (Table 1) produced by L. reuteri 121 cells and the IMMP produced by the GtfB enzyme from different maltodextrins were analyzed by SEC in comparison with their original substrate. EPS derived from AVEBE SPG30 (< 1.0 kDa) and AVEBE MD20 (approx. 1.0 kDa) (Table 1) both contain two peaks of much bigger polymers (Fig. 3A and B) of relatively low (Region II, 132 kDa for EPS SPG30, 706 kDa for EPS MD20) and relatively high (Region I, 1.7 MDa for EPS SPG30, 3.3 MDa for EPS MD20) apparent average molar mass (MM) (Fig. 3). The high MM products in all cases represented only a small percentage (2.5%). In general, these EPS thus are much larger than the IMMP produced by the GtfB enzyme from the same maltodextrins (Fig. 3). A possible explanation is that the reaction conditions for the L. reuteri 121 cell-associated GtfB enzyme and the free purified GtfB enzyme differ. Any glucose and maltose produced by the L. reuteri 121 cell-associated GtfB enzyme can be consumed by the cells as carbon source and thus cannot be used as acceptor substrate in further reaction. On the contrary, the glucose and maltose produced in the in vitro GtfB reaction can be used as acceptor substrate (26), resulting in relatively high yield of oligosaccharides, which have no clear increase in chain length compared to maltodextrin substrate.

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Fig. 3. SEC chromatograms of the different maltodextrin substrate (A, SPG30, B, MD20), the EPS produced by L. reuteri 121 cells, and IMMP produced by the free GtfB-∆N enzyme. Elution times of the pullulan standards corresponding to 366, 200, 113, 48.8, 21.7, 10, 6.2, 1.32 and 0.342 kDa, are shown as gray dots.

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SEC analysis of EPS and IMMP derived of Etenia 457 and wheat starch

NMR analysis showed that L. reuteri 121 cells convert various starches into EPS with increased percentages of α1→6 linkages in linear chains (Table 1). To compare starch conversion by the L. reuteri 121 cell-associated GtfB enzyme and by the free GtfB enzyme, Etenia 457 and wheat starch (28.8% amylose) were used as substrates (Table 1). The size of these substrates, and the EPS and IMMP produced were analyzed using SEC.

Etenia 457 is produced from potato starch by amylomaltase (4-α-glucanotransferase) treatment in which the amylose part gradually is transferred to the non-reducing ends of the side chains of amylopectin, resulting in elongated external chains (41). As shown in Fig. 4A, the L. reuteri 121 EPS derived from Etenia 457 (approx. 350-400 kDa) has almost the same SEC profile as Etenia 457 itself (thus no clear change in MM) although the NMR results (Table 1) showed that this EPS has more α1→6 linkages. By contrast, the SEC profile of the IMMP produced by GtfB from Etenia 457 clearly shows that Etenia 457 was converted into two main products with smaller MM (approx. 200 kDa in Region II and approx. 10 kDa in III). The material present in region II most likely represents Etenia 457 derived products with shortened non-reducing end side chains, used as donor substrate or hydrolyzed by GtfB. Region III represents Etenia 457 derived smaller MM products.

As any size difference between Etenia 457 and the IMMP and EPS product derived might be covered by their relatively broad size distribution in the SEC analysis, these three polymers were debranched by isoamylase and subsequently analyzed again by SEC (Fig. 4B) (42). After debranching, the side chains of Etenia 457/isoamylase all eluted in region II’. The debranched products from IMMP and EPS were both smaller than those from the Etenia 457 substrate, indicating that the original amylopectin side chains were modified. However, the EPS and IMMP differed as isoamylase treated EPS formed by L. reuteri 121 cells contained some larger MM material, eluting in Region I′.

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Fig. 4. SEC chromatograms of Etenia 457, the EPS produced from Etenia 457 by L. reuteri 121 and IMMP produced from Etenia 457 by the free GtfB-∆N enzyme, before (A) and after (B) isoamylase treatment. Elution times of the pullulan standards corresponding to 366, 200, 113, 48.8, 21.7, 10, 6.2, 1.32 and 0.342 kDa, are shown as gray dots.

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Starches with high amylose content have been described as preferred substrates for GtfB.27 In L. reuteri 121 cultures, starches with a higher amylose content also yielded higher percentages of α1→6 linkages (Table 1). For instance, the EPS and IMMP derived from wheat starch both have approx. 22% of α1→6 linkages, which is much higher than the original 3.7% (Table 1). As shown in Fig. 5A, the graph representing wheat starch shows a peak in region I representing amylopectin content and a peak in region II representing amylose which are overlapping (42). After fermentation with L. reuteri 121, the amylopectin content decreased with an increase of products in region II. The SEC profile of the IMMP produced by GtfB from wheat starch is largely different from that of the wheat starch substrate. Wheat starch clearly was converted into two main products with smaller MM (approx. 300 kDa and approx. 15 kDa). The data indicates that GtfB also is capable to modify the amylopectin part in native wheat starch.

After debranching, the SEC curve of wheat starch was divided into two parts: one is the amylose part, which cannot be processed by isoamylase (Region I’); the other is the bimodal peak representing the amylopectin branches including side and inner chains (Fig. 5b) (42). For EPS derived from wheat starch, the SEC curve shows significant differences. The peak in region III’ representing the long branches of amylopectin in wheat starch decreased a lot, suggesting that L. reuteri 121 also modifies the side chains of amylopectin in wheat starches. The peak representing amylose in region I’ (> 400 kDa) shifted dramatically and formed a new peak in region II’ (approx. 200 kDa), indicating that the amylose content of wheat starch was converted by L. reuteri 121 and new, smaller products were generated. By contrast, the amylose part in region I’ was completely converted in vitro by the GtfB enzyme as the peak representing amylose disappeared. The amount of the short chains after debranching process increased.

To conclude, L. reuteri 121 cells and the GtfB enzyme are capable of modifying the amylose molecule as well as the side chains of amylopectin molecules in starches, introducing higher percentages of α1→6 linkages. The EPS produced have similar size distribution as the starch substrates although the percentages of α1→6 linkages increased significantly compared to the starch substrates. The produced IMMP are clearly different from the produced EPS and starch

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substrates with regard to their size distribution. This may result from the minor endo-acting activity (Bai et al., to be submitted) of the GtfB enzyme. However, during the L. reuteri 121 fermentations, the interactions of the starch substrate with the cell-associated GtfB enzyme clearly are different from those with the purified GtfB enzyme. Conceivably, the endo-acting activity of the cell-associated GtfB enzyme is impaired compared to the free GtfB enzyme. In addition, any glucose released by the L. reuteri 121 cells may be consumed whereas the free GtfB enzyme is able to use glucose as acceptor substrate (26), resulting in smaller size products. This might influence product synthesis, resulting in the size differences between IMMP and EPS.

Structural analysis of EPS derived from maltodextrins by enzymatic

digestion

IMMP produced by the GtfB enzyme from maltodextrin substrates were characterized as soluble fibers (27). They are structurally different from the known starch and maltodextrin molecules as GtfB builds linear consecutive (1→6)-α-glucan chains onto the non-reducing end of (1→4)-α-glucan chains. Thus IMMP derived from maltodextrins are mainly composed by sequences of -α-D-Glcp-(1→6)-α-D-Glcp-(1→4)-, -α-D-Glcp-(1→6)-α-D-Glcp-(1→6)-, and -α-D-Glcp-(1→4)-α-D-Glcp-(1→4)-, as well as some branching -α-D-Glcp-(1→4,6)-α-D-Glcp-(1→4)- originating from the substrates (Fig. 6B) (27).

To reveal whether the IMMP-like EPS produced by L. reuteri 121 cells and cultures are structurally similar to IMMP produced by the GtfB enzyme, the AVEBE MD6 derived EPS were subjected to hydrolysis by several enzymes that are typical for structural analysis of polysaccharides. α-Amylase (Fig. 6B) is an endo-acting enzyme cleaving α1→4 glucosidic linkages inside starch molecules mainly producing maltose. Dextranase of Chaetomium erraticum is an (1→6)-α-glucan 6-glucanohydrolase that endo-hydrolyzes (α1→6)-α-glucosidic linkages (Fig. 6B) in dextran mainly producing glucose and isomaltose. As shown in Fig. 6A, glucose and maltose were released by α-amylase hydrolysis of EPS, suggesting that this EPS contains -α-D-Glcp-(1→4)-α-D-Glcp-(1→4)- sequences. Likewise, the presence of -α-D-Glcp-(1→6)-α-D-Glcp-(1→6)- sequences were also confirmed because dextranase hydrolysis generated glucose and isomaltose.

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Fig. 5. SEC chromatograms of wheat starch and the EPS produced from wheat starch by L. reuteri 121, and IMMP produced from wheat starch by the free GtfB-∆N enzyme, before (A) and after (B) isoamylase treatment. Elution times of the pullulan standards corresponding to 366, 200, 113, 48.8, 21.7, 10, 6.2, 1.32 and 0.342 kDa, are shown as gray dots.

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Pullulanase type I specifically attacks α1→6 linkages in both -α-D-Glcp-(1→6)-α-D-Glcp-(1→4)- sequences (Fig. 6B) in pullulan and -α-D-Glcp-(1→4,6)-α-D-Glcp-(1→4)- branch points (Fig. 6B) in starch molecules (43). Isoamylase exclusively cleaves the branching α1→4,6 linkages (44). AVEBE MD6 derived EPS were hydrolyzed by both pullulanase and isoamylase, forming similar products (Fig. 6A and Fig. S2). These EPS thus possess -α-D-Glcp-(1→4,6)-α-D-Glcp-(1→4)- branch points which also exist in IMMP (27). Isopullulanase from Aspergillus niger exclusively cleaves the α1→4 linkages in -α-D-Glcp-(1→6)-α-D-Glcp-(1→4)- sequences (Fig. 6B) in pullulan but has no activity on starches (45). After treatment of EPS with isopullulanase, oligomers appeared (Fig. 6A), confirming the presence of linear -α-D-Glcp-(1→6)-α-D-Glcp-(1→4)- sequences. These enzymatic hydrolysis results thus showed that the EPS produced by L. reuteri 121 and IMMP produced by GtfB are structurally similar since both contain the same fragments.

To further investigate the presence of consecutive α1→6 linkages that are typical in IMMP, the EPS derived from AVEBE MD6 was treated with a combination of pullulanase and α-amylase to remove the consecutive α1→4 linkages, and the α1→6 linkages in connecting -α-D-Glcp-(1→6)-α-D-Glcp-(1→4)- sequences and -α-D-Glcp-(1→4,6)-α-D-Glcp-(1→4)- branches. The hydrolyzed products were fractionated on Bio-Gel P2, yielding nine fractions, denoted F1-F9. F8 and F9 were mainly composed of maltose and glucose (TLC, data not shown) that resulted from consecutive α1→4 linkages hydrolyzed by α-amylase. With the increase of DP, the percentage of α1→6 linkages increased (Table S1). In fractions 1 and 2, the polymers were mainly composed of α-glucan with 94% α1→6 linkages. This shows that linear consecutive α1→6 linked fragments inside EPS are present with a broad DP distribution.

Fermentation of L. reuteri EPS by probiotic Bifidobacterium strains

The EPS produced by L. reuteri 121 has a special structure, which is similar to IMMP but different from known α-glucans such as starch, dextran and pullulan. Previously, IMMP were shown to be soluble dietary fibers (27). EPS thus may partially escape digestion in the upper part of the gastrointestinal tract and small intestine, to serve as typical carbon sources for bacterial fermentation in the large intestine. Alternatively, uptake of probiotic L. reuteri 121 most likely results in

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formation of EPS rich in α1→6 linkages in the upper gastrointestinal tract by 4,6-α-GTase enzymatic modification of the available (partially hydrolyzed) starch. The in vivo synthesized EPS may selectively stimulate the growth of (other) probiotic bacteria in the large intestine.

Fig. 6. (A) HPAEC analysis of the hydrolysis products of EPS (from AVEBE MD6) following incubation with an excess of different enzymes, including α-amylase, dextranase, pullulanase, isoamylase, and isopullulanase. A combination of glucose (G1), isomaltose (isoG2), isomaltotriose (isoG3), maltose (G2), panose (P), maltotriose (G3), maltotetraose (G4), maltopentaose (G5), maltohexaose (G6), maltoheptaose (G7), and maltooctaose (G8), were used as standard. EPS without enzymatic treatment was applied as negative control. (B) Representation of the IMMP products of 4,6-α-GTase (e.g. GtfB) with starch and maltodextrins. GtfB builds (1→6)-α-glucan chains with occasionally internal 1→4 linkages onto the non-reducing end of (1→4)-α-glucan chains (28). Schematic diagram of the α-amylase, dextranase, pullulanase, isoamylase, and isopullulanase enzymatic cleavage points in IMMP.

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Bifidobacterium is one of the major genera of bacteria that make up the colon flora. Some of them are used as probiotics in food industry as they may exert a range of beneficial health effects (46, 47). To date, many type II pullulanases (amylopullulanases) with both α-amylase and pullulanase activity have been identified in probiotic Bifidobacterium species from the human gut (31, 48, 49). It was also reported that amylopullulanase is the only extracellular polysaccharide hydrolyzing enzyme in lactobacilli (18). Based on the above data (Fig. 6 and Fig. S2), L. reuteri 121 EPS derived from maltodextrins can be partially hydrolyzed by both α-amylase and pullulanase. Previously, we reported that reuteran derived from sucrose can be hydrolyzed by pullulanase (9). Thus, the Bifidobacterium strains producing extracellular amylopullulanase have strong potential to degrade both L. reuteri 121 EPS derived from maltodextrins, and reuteran, and to use the degradation products as carbon- and energy source. Growth of Bifidobacterium breve DSM 20091 and Bifidobacterium dentium DSM 20436, both containing amylopullulanase activity (48), was studied using EPS of L. reuteri 121 as carbon source. Both B. breve DSM20091 (Fig. 7b) and B. dentium DSM20436 (Fig. 7c) grew well with different EPS. After 12 h of growth, OD600 values of B. breve DSM20091 all reached 80% with EPS derived from MD20, MD6, or SPG30 (Table 1), and 67.5% with EPS derived from MD4-7, compared to a 100% control grown on glucose. Growth curves of B. dentium with EPS followed closely the glucose-control reaching all a final OD600nm value of 2.00 after 8.5 h. The exponential growth phases with the EPS growth substrate did not show any differences to the glucose-control indicating that these EPS are highly suitable as growth substrates. In addition, total carbohydrate measurements and determination of OD600 values and pH values (Table 2), as well as TLC detection (Fig. S1) also showed that EPS was partially or completely used by strains.

As reported, the EPS produced by L. reuteri 121 from maltodextrins is hardly degraded by human gut symbiont Bacteroides thetaiotaomicron (50). In this study, Bifidobacterium adolescentis DSM 20083 that cannot degrade starch (data not shown) was also investigated, showing that it cannot grow with any of the EPS derived from maltodextrins (Fig. 7A). Therefore, EPS of L. reuteri 121 derived from both maltodextrins and sucrose are potential prebiotics which typically stimulate growth of Bifidobacterium strains which have amylopullulanase activity.

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Fig. 7. Growth of B. adolescentis DSM 20083 (a), B. breve DSM 20091 (b) and B. dentium DSM 20436 (c) on different EPS (5 mg/ml) derived from different maltodextrins. Glucose (5 mg/ml) served as positive control.

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Table 2. OD600, pH values and total carbohydrate consumption values were determined after growth of B. breve DSM 20091 and B. dentium DSM 20436 for 24 h with L. reuteri 121 EPS derived from different maltodextrins, and reuteran derived from sucrose. Experiments were done in duplicate and the average data are shown.

Bifidobacterium breve DSM 20091 Bifidobacterium dentium DSM 20436

OD600 pH

Total carbohydrate consumption

(%)

OD600 pH

Total carbohydrate consumption

(%)

Control 0.03 6.8 - 0.03 6.8 -

Glucose 1.68 4.9 100 1.85 5.0 100 EPS Avebe

SPG30 1.54 5.2 74 1.68 5.2 84

EPS Avebe MD20 1.46 5.3 81 1.56 5.3 85

EPS Avebe MD6 1.63 5.2 91 1.68 5.1 88

EPS MD 4-7 1.62 5.2 82 1.70 5.1 86 Reuteran 1.56 5.0 84 1.80 5.0 95

To conclude, in this study, novel homo-EPS produced by probiotic bacterium L. reuteri strains from starch or maltodextrins as substrates were characterized as IMMP-like EPS. The data show that the cell-associated 4,6-α-glucanotransferase enzyme plays an important role in the formation of such IMMP-like EPS in L. reuteri strains. The L. reuteri 121 EPS are structurally similar to the IMMP produced by the free GtfB enzyme, with percentages of α1→6 linkages ranging between 11% to 52% (Table 1). However, the L. reuteri 121 EPS derived from either maltodextrins or starches are generally much larger than the IMMP produced by the in vitro GtfB enzyme. The SEC results also demonstrated that L. reuteri 121 is capable to modify starches by converting both the amylose part and the side chains of amylopectin molecules. Finally, the EPS derived from different maltodextrins were shown to be excellent growth substrates for some probiotic Bifidobacterium strains that have amylopullulanase activity. These EPS thus have potential prebiotic activity and may contribute to the application of probiotic L. reuteri strains grown on maltodextrins or starch as synbiotics.

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Acknowledgements The study was financially supported by the Chinese Scholarship Council (to YB), the University of Groningen, and by the TKI Agri&Food program as coordinated by the Carbohydrate Competence Center (CCC-ABC and CCC-3; www.cccresearch.nl). ALVB is supported by a VENI Grant by the Netherlands Organization for Scientific Research (NWO).

References 1. Siroli, L.; Patrignani, F.; Serrazanetti, D. I.; Tabanelli, G.; Montanari, C.; Gardini, F.; Lanciotti, R., Lactic acid bacteria and natural antimicrobials to improve the safety and shelf-life of minimally processed sliced apples and lamb's lettuce. Food Microbiol. 2015, 47, 74-84. 2. Liu, M. J.; Siezen, R., Comparative genomics of flavour-forming pathways in lactic acid bacteria. Aust. J. Dairy Technol. 2006, 61, 61-68. 3. Yang, T. X.; Wu, K. Y.; Wang, F.; Liang, X. L.; Liu, Q. S.; Li, G. L.; Li, Q. Y., Effect of exopolysaccharides from lactic acid bacteria on the texture and microstructure of buffalo yoghurt. Int. Dairy J. 2014, 34, 252-256. 4. Matsuzaki, C.; Hayakawa, A.; Matsumoto, K.; Katoh, T.; Yamamoto, K.; Hisa, K., Exopolysaccharides Produced by Leuconostoc mesenteroides strain NTM048 as an immunostimulant to enhance the mucosal barrier and influence the systemic immune response. J. Agr. Food. Chem. 2015, 63, 7009-7015. 5. Wang, K.; Li, W.; Rui, X.; Chen, X.; Jiang, M.; Dong, M., Structural characterization and bioactivity of released exopolysaccharides from Lactobacillus plantarum 70810. Int. J. Biol. Macromol. 2014, 67, 71-8. 6. Leemhuis, H.; Pijning, T.; Dobruchowska, J. M.; van Leeuwen, S. S.; Kralj, S.; Dijkstra, B. W.; Dijkhuizen, L., Glucansucrases: three-dimensional structures, reactions, mechanism, α-glucan analysis and their implications in biotechnology and food applications. J. Biotechnol. 2013, 163, 250-272. 7. De Vuyst, L.; De Vin, F.; Vaningelgem, F.; Degeest, B., Recent developments in the biosynthesis and applications of heteropolysaccharides from lactic acid bacteria. Int. Dairy J. 2001, 11, 687-707. 8. Laws, A.; Gu, Y. C.; Marshall, V., Biosynthesis, characterisation, and design of bacterial exopolysaccharides from lactic acid bacteria. Biotechnol. Adv. 2001, 19, 597-625.

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9. van Leeuwen, S. S.; Kralj, S.; van Geel-Schutten, I. H.; Gerwig, G. J.; Dijkhuizen, L.; Kamerling, J. P., Structural analysis of the α-D-glucan (EPS35-5) produced by the Lactobacillus reuteri strain 35-5 glucansucrase GTFA enzyme. Carbohyd. Res. 2008, 343, 1251-1265. 10. Bounaix, M. S.; Gabriel, V.; Morel, S.; Robert, H.; Rabier, P.; Remaud-Simeon, M.; Gabriel, B.; Fontagne-Faucher, C., Biodiversity of exopolysaccharides produced from sucrose by sourdough lactic acid bacteria. J. Agr. Food. Chem. 2009, 57, 10889-10897. 11. Tieking, M.; Ganzle, M. G., Exopolysaccharides from cereal-associated lactobacilli. Trends Food Sci. Tech. 2005, 16, 79-84.

12. van Hijum, S. A. F. T.; Kralj, S.; Ozimek, L. K.; Dijkhuizen, L.; van Geel-Schutten, I. G. H., Structure-function relationships of glucansucrase and fructansucrase enzymes from lactic acid bacteria. Microbiol. Mol. Biol. R 2006, 70, 157-176.

13. Kralj, S.; van Geel-Schutten, G. H.; van der Maarel, M. J. E. C.; Dijkhuizen, L., Biochemical and molecular characterization of Lactobacillus reuteri 121 reuteransucrase. Microbiol-SGM 2004, 150, 2099-2112.

14. Meng, X. F.; Dobruchowska, J. M.; Gerwig, G. J.; Kamerling, J. P.; Dijkhuizen, L., Synthesis of oligo- and polysaccharides by Lactobacillus reuteri 121 reuteransucrase at high concentrations of sucrose. Carbohyd. Res. 2015, 414, 85-92.

15. Dobruchowska, J. M.; Meng, X. F.; Leemhuis, H.; Gerwig, G. J.; Dijkhuizen, L.; Kamerling, J. P., Gluco-oligomers initially formed by the reuteransucrase enzyme of Lactobacillus reuteri 121 incubated with sucrose and malto-oligosaccharides. Glycobiology 2013, 23, 1084-1096. 16. Haydersah, J.; Chevallier, I.; Rochette, I.; Mouquet-Rivier, C.; Picq, C.; Marianne-Pepin, T.; Icard-Verniere, C.; Guyot, J. P., Fermentation by amylolytic lactic acid bacteria and consequences for starch digestibility of plantain, breadfruit, and sweet potato flours. J. Food Sci. 2012, 77, M466-M472. 17. Petrova, P.; Petrov, K.; Stoyancheva, G., Starch-modifying enzymes of lactic acid bacteria - structures, properties, and applications. Starch-Starke 2013, 65, 34-47. 18. Ganzle, M. G.; Follador, R., Metabolism of oligosaccharides and starch in lactobacilli: a review. Front. microbiol. 2012, 3.

19. Pompeyo, C. C.; Gomez, M. S.; Gasparian, S.; Morlonguyot, J., Comparison of Amylolytic properties of Lactobacillus amylovorus and of Lactobacillus amylophilus. Appl. Microbiol. Biot. 1993, 40, 266-269.

20. Champ, M.; Szylit, O.; Raibaud, P.; Ait-Abdelkader, N., Amylase production by three Lactobacillus strains isolated from chicken crop. J. Appl. Bacteriol. 1983, 55, 487-93.

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21. Sen, S.; Chakrabarty, S. L., Amylase from Lactobacillus cellobiosus isolated from vegetable wastes. J. Ferment. Technol. 1984, 62, 407-413.

22. Wasko, A.; Polak-Berecka, M.; Targonski, Z., Purification and characterization of pullulanase from Lactococcus lactis. Prep. Biochem. Biotech. 2011, 41, 252-261. 23. Petrova, P.; Petrov, K., Direct starch conversion into L-(+)-lactic acid by a novel amylolytic strain of Lactobacillus paracasei B41. Starch-Starke 2012, 64, 10-17. 24. van der Maarel, M. J. E. C.; Leemhuis, H., Starch modification with microbial α-glucanotransferase enzymes. Carbohyd. Polym. 2013, 93, 116-121. 25. Leemhuis, H.; Dijkman, W. P.; Dobruchowska, J. M.; Pijning, T.; Grijpstra, P.; Kralj, S.; Kamerling, J. P.; Dijkhuizen, L., 4,6-α-Glucanotransferase activity occurs more widespread in Lactobacillus strains and constitutes a separate GH70 subfamily. Appl. Microbiol. Biot. 2013, 97, 181-193. 26. Kralj, S.; Grijpstra, P.; van Leeuwen, S. S.; Leemhuis, H.; Dobruchowska, J. M.; van der Kaaij, R. M.; Malik, A.; Oetari, A.; Kamerling, J. P.; Dijkhuizen, L., 4,6-α-glucanotransferase, a novel enzyme that structurally and functionally provides an evolutionary link between glycoside hydrolase enzyme families 13 and 70. Appl. Environ. Microb. 2011, 77, 8154-8163.

27. Leemhuis, H.; Dobruchowska, J. M.; Ebbelaar, M.; Faber, F.; Buwalda, P. L.; van der Maarel, M. J.; Kamerling, J. P.; Dijkhuizen, L., Isomalto/Malto-polysaccharide, a novel soluble dietary fiber made via enzymatic conversion of starch. J. Agr. Food. Chem. 2014, 62, 12034-12044.

28. Dobruchowska, J. M.; Gerwig, G. J.; Kralj, S.; Grijpstra, P.; Leemhuis, H.; Dijkhuizen, L.; Kamerling, J. P., Structural characterization of linear isomalto-/malto-oligomer products synthesized by the novel GTFB 4,6-a-glucanotransferase enzyme from Lactobacillus reuteri 121. Glycobiology 2012, 22, 517-528. 29. Kralj, S.; van Geel-schutten, G. H.; van der Maarel, M. J. E. C.; Dijkhuizen, L., Efficient screening methods for glucosyltransferase genes in Lactobacillus strains. Biocatal. Biotransfor. 2003, 21, 181-187. 30. Zhang, Y. H.; Ding, L. L.; Gu, J. Y.; Tan, H. Y.; Zhu, L. B., Preparation and properties of a starch-based wood adhesive with high bonding strength and water resistance. Carbohyd. Polym. 2015, 115, 32-37. 31. Ryan, S. M.; Fitzgerald, G. F.; van Sinderen, D., Screening for and identification of starch-, amylopectin-, and pullulan-degrading activities in Bifidobacterial strains. Appl. Environ. Microb. 2006, 72, 5289-5296. 32. Van Geel-Schutten, G. H.; Faber, E. J.; Smit, E.; Bonting, K.; Smith, M. R.; Ten Brink, B.; Kamerling, J. P.; Vliegenthart, J. F. G.; Dijkhuizen, L., Biochemical and structural characterization of the glucan and fructan exopolysaccharides synthesized by

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the Lactobacillus reuteri wild-type strain and by mutant strains. Appl. Environ. Microb. 1999, 65, 3008-3014.

33. van Geel-Schutten, G. H.; Flesch, F.; ten Brink, B.; Smith, M. R.; Dijkhuizen, L., Screening and characterization of Lactobacillus strains producing large amounts of exopolysaccharides. Appl. Microbiol. Biot. 1998, 50, 697-703.

34. Sims, I. M.; Frese, S. A.; Walter, J.; Loach, D.; Wilson, M.; Appleyard, K.; Eason, J.; Livingston, M.; Baird, M.; Cook, G.; Tannock, G. W., Structure and functions of exopolysaccharide produced by gut commensal Lactobacillus reuteri 100-23. ISME J. 2011, 5, 1115-1124.

35. Lee, I. Y.; Kim, M. K.; Lee, J. H.; Seo, W. T.; Jung, J. K.; Lee, H. W.; Park, Y. H., Influence of agitation speed on production of curdlan by Agrobacterium species. Bioprocess Eng. 1999, 20, 283-287.

36. Cui, J. D.; Qiu, J. Q., Production of extracellular water-insoluble polysaccharide from Pseudomonas sp. J. Agr. Food. Chem. 2012, 60, 4865-4871.

37. Bai, Y.; van der Kaaij, R. M.; Woortman, A. J. J.; Jin, Z.; Dijkhuizen, L., Characterization of the 4,6-α-glucanotransferase GTFB enzyme of Lactobacillus reuteri 121 isolated from inclusion bodies. BMC Biotechnol. 2015, 15, 49.

38. Kralj, S.; van Geel-Schutten, G. H.; Rahaoui, H.; Leer, R. J.; Faber, E. J.; van der Maarel, M. J. E. C.; Dijkhuizen, L., Molecular characterization of a novel glucosyltransferase from Lactobacillus reuteri strain 121 synthesizing a unique, highly branched glucan with α-(1→4) and α-(1→6) glucosidic bonds. Appl. Environ. Microb. 2002, 68, 4283-4291. 39. Bai, Y.; van der Kaaij, R. M.; Leemhuis, H.; Pijning, T.; van Leeuwen, S. S.; Jin, Z.; Dijkhuizen, L., Biochemical characterization of Lactobacillus reuteri Glycoside Hydrolase family 70 GTFB type of 4,6-α-Glucanotransferase enzymes that synthesize soluble dietary starch fibers. Appl. Environ. Microb. 2015, 81, 7223-7232. 40. Kralj, S.; van Geel-Schutten, G. H.; Dondorff, M. M. G.; Kirsanovs, S.; van der Maarel, M. J. E. C.; Dijkhuizen, L., Glucan synthesis in the genus Lactobacillus: isolation and characterization of glucansucrase genes, enzymes and glucan products from six different strains. Microbiol-SGM 2004, 150, 3681-3690. 41. Kaper, T.; van der Maarel, M. J. E. C.; Euverink, G. J. W.; Dijkhuizen, L., Exploring and exploiting starch-modifying amylomaltases from thermophiles. Biochem. Soc. T. 2004, 32, 279-282. 42. Ciric, J.; Woortman, A. J.; Loos, K., Analysis of isoamylase debranched starches with size exclusion chromatography utilizing PFG columns. Carbohyd. Polym. 2014, 112, 458-461.

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43. Doman-Pytka, M.; Bardowski, J., Pullulan degrading enzymes of bacterial origin. Crit. Rev. Microbiol. 2004, 30, 107-121.

44. van der Maarel, M. J.; van der Veen, B.; Uitdehaag, J. C.; Leemhuis, H.; Dijkhuizen, L., Properties and applications of starch-converting enzymes of the α-amylase family. J. Biotechnol. 2002, 94, 137-55.

45. Hii, S. L.; Tan, J. S.; Ling, T. C.; Ariff, A. B., Pullulanase: role in starch hydrolysis and potential industrial applications. Enzyme research 2012, 2012, 921362. 46. Oner, O.; Aslim, B.; Aydas, S. B., Mechanisms of cholesterol-lowering effects of Lactobacilli and Bifidobacteria strains as potential probiotics with their bsh gene analysis. J. Mol. Microb. Biotech. 2014, 24, 12-18. 47. Di Gioia, D.; Aloisio, I.; Mazzola, G.; Biavati, B., Bifidobacteria: their impact on gut microbiota composition and their applications as probiotics in infants. Appl. Microbiol. Biot. 2014, 98, 563-577. 48. Motherway, M. O.; Fitzgerald, G. F.; Neirynck, S.; Ryan, S.; Steidler, L.; van Sinderen, D., Characterization of ApuB, an extracellular type II amylopullulanase from Bifidobacterium breve UCC2003. Appl. Environ. Microb. 2008, 74, 6271-6279. 49. Moller, M. S.; Goh, Y. J.; Viborg, A. H.; Andersen, J. M.; Klaenhammer, T. R.; Svensson, B.; Abou Hachem, M., Recent insight in α-glucan metabolism in probiotic bacteria. Biologia 2014, 69, 713-721. 50. van Bueren, A. L.; Saraf, A.; Martens, E. C.; Dijkhuizen, L., Differential metabolism of exopolysaccharides from probiotic Lactobacilli by the human gut symbiont Bacteroides thetaiotaomicron. Appl. Environ. Microb. 2015. 81, 3973-83.

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Fig. S1. TLC analysis of the saccharides in the cell-free supernatant before (1-5) and after (1’-5’) growth of Bifidobacterium breve DSM 20091 and Bifidobacterium dentium DSM 20436 incubated with L. reuteri 121 EPS derived from different maltodextrins (1, SPG30, 2, MD20, 3, MD6, 4, MD4-7), and reuteran (5) derived from sucrose. BM medium without carbon source and with glucose was spotted as negative control (C) and positive control before (6) and after (6’) fermentation.

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Fig. S2. TLC analysis of the enzymatic hydrolysis of EPS (from AVEBE MD6) by different enzymes, α-amylase, L1; dextranase, L2; pullulanase, L3; isoamylase, L4; and isopullulanase, L5; control (EPS), C.

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Table S1. The amylase-pullulanase treated EPS produced from Paselli MD6 was fractionated by Biogel P2 column into 9 fractions (fraction 1 represents the saccharides with highest DP and fraction 9 represents the saccharides with lowest DP). The linkage specificity of each fraction was recorded by 1H NMR spectroscopy.

Fractions α-1,4 (%) α-1,6 (%)

1 6 94

2 6 94

3 11 88

4 20 80

5 51 49

6 69 31

7 94 6

8 98 2

9 100 0

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Chapter 5

Structural basis of the role of starch and sucrose in homo-

exopolysaccharide formation in Lactobacilli reuteri 35-5

Yuxiang Bai1,2, Justyna M Dobruchowska1, Rachel Maria van der Kaaij1, Gerrit J

Gerwig1,3, Lubbert Dijkhuizen1,*

1Microbial Physiology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, Nijenborgh 7, 9747 AG Groningen, The Netherlands

2The State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi 214122, China

3NMR Spectroscopy, Bijvoet Center for Biomolecular Research, Utrecht University, Padualaan 8, NL-3584 CH Utrecht, The Netherlands

In preparation for submission

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Abstract Lactic acid bacteria (LAB) are abundant producers of exopolysaccharides (EPS) which are important for biofilm formation in the oral cavity and gastrointestinal tract of mammals. Sucrose is a well-known substrate for homo-EPS formation by glucansucrases (GS) of LAB. Starch is the main fermentable carbohydrate in the human diet, and often consumed simultaneously with sucrose. Recently, 4,6-α-glucanotransferases (4,6-α-GTases) of Lactobacillus reuteri strains were characterized that act on starch and maltodextrins, cleaving α1→4 linkages and synthesizing α1→6 linkages, yielding isomalto-/malto-polysaccharides (IMMP) that are soluble dietary fibers. Lactobacillus strains (e.g. L. reuteri 35-5) may possess both a GS (e.g. GtfA) and a 4,6-α-GTases (GtfB), both cell-associated and located extracellularly. In this study, we characterized the EPS formed in vivo by L. reuteri 35-5 cells during growth with either sucrose or maltodextrins, or with both. Size exclusion chromatography and enzymatic hydrolysis revealed variations in EPS size and structure depending on the substrate(s) provided. In vitro incubation of the purified GtfA-∆N plus GtfB-∆N enzymes in the presence of both sucrose and starch resulted in formation of oligosaccharides with terminally and internally consecutive α1→6 linkages, which are structurally different from the oligosaccharides produced from either of the single substrates. In conclusion, the data show that both in vivo and in vitro the GtfA and GtfB enzymes of L. reuteri 35-5, incubated with sucrose plus starch, cross-react and contribute to the synthesis of the final products. This may have strong effects on the functional properties of the EPS formed by lactobacilli, and influence their ability for biofilm formation in the mammalian oral cavity and gastrointestinal tract. Also the relationship between dietary intake of sucrose and starch, dental caries formation and degradability of EPS dietary fibers, are likely to be affected.

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Abstract Figure. Schematic model of the exopolysaccharides produced by GtfA and GtfB enzymes in the presence of sucrose and starch in Lactobacillus reuteri 35-5.

Introduction Exopolysaccharides (EPS) constitute a major fraction of the extracellular polymeric substances that form the scaffold for the three-dimensional architecture of biofilms (1). In many bacteria, EPS are indispensable for biofilm formation: mutants that have lost ability to synthesize EPS are unable or severely compromised in forming mature biofilms (2-5). Lactic acid bacteria (LAB) use glucansucrase (GS) and fructansucrase enzymes to synthesize α-glucan and β-fructan homo-EPS from sucrose (6-10). GS are family 70 glycoside hydrolase (GH70) enzymes that are able to synthesize a diversity of α-glucans varying in linkage type and degree of branching (11). In the oral cavity and gastrointestinal tract of mammals, the α-glucan polysaccharides produced by GS facilitate biofilm formation and significantly enhance the adhesion of bacteria to the tooth enamel or epithelium (12, 13).

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Starch is an important carbohydrate in the human diet and usually consumed simultaneously with sucrose. Simultaneous intake of starch and sucrose resulted in enhanced cariogenicity in human compared to use of either carbohydrate alone (14-17). The EPS matrix produced by Streptococcus mutans from sucrose plus starch contains a larger amount of highly branched α-glucan compared to the EPS derived from either of the single substrates (18). In addition to S. mutans, many other species of LAB that produce EPS from sucrose, using GS, are found in the oral cavity and gastrointestinal tract of mammals (9, 19, 20). No information is available on the EPS synthesized by LAB strains challenged with both sucrose and starch as carbon sources for growth.

In recent years, we have identified a new GH70 subfamily with 4,6-α-glucanotransferase (4,6-α-GTase) enzymes in Lactobacillus strains. These 4,6-α-GTases do not act on sucrose but on α-glucans (starch, maltodextrins), cleaving α1→4 linkages and forming α1→6 linkages, resulting in isomalto-/malto- polysaccharide (IMMP) synthesis that act as soluble dietary fibers (21-23). According to current database searches (http://www.ncbi.nlm.nih.gov/), around 25% of the LAB strains carrying a 4,6-α-GTase gene also possess a GS gene. In our previous study, L. reuteri 121 cultures with the extracellular and cell-associated 4,6-α-GTase GtfB were shown to synthesize IMMP-like EPS from starch (Bai et al., submitted). Here we study whether co-feeding of sucrose and starch influences EPS formation by L. reuteri 121 strain 35-5. This mutant only has GS (GtfA) and 4,6-α-GTase (GtfB) enzymes but no fructansucrase or other homo-EPS producing enzymes (24). GtfA converts sucrose into reuteran with alternating α1→4 and α1→6 glycosidic linkages; GtfB converts starches and maltodextrins into IMMP with consecutive α1→6 linkages (Abstract Figure) (25-28). Here, we analyzed the different L. reuteri 35-5 EPS synthesized during growth with sucrose, maltodextrins, and sucrose plus maltodextrins. We also studied the EPS synthesized by GtfA-∆N and GtfB-∆N enzymes in vitro, by analyzing initial oligosaccharide formation from these substrates. The data shows that homo-EPS formation by L. reuteri 35-5 cells results from the combined activity of GtfA plus GtfB with sucrose plus maltodextrins, in addition to the conventional single-enzyme reactions of GtfA and GtfB with either sucrose or maltodextrins, respectively. Synthesis of such hybrid EPS from sucrose and maltodextrins/starches may widely occur in LAB strains that have similar gene

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combinations. This study provides new insights into homo-EPS formation by LAB, potentially generating new EPS functionalities.

Materials and Methods

Growth conditions

Lactobacillus reuteri 35-5 (29) was subcultured from stocks stored at -80 °C in 10 ml of sugar-free MRS medium (liquid culture or agar plate) supplemented with 1% (w/v) glucose. The fresh cultures were inoculated anaerobically in modified sugar-free MRS medium with sucrose (5%, w/v), or maltodextrins (dextrose equivalent (DE) = 13-17, Sigma-Aldrich, US, 5%, w/v), or sucrose (2.5%, w/v) plus maltodextrins (2.5 %, w/v). One liter medium contained 10 g bactopeptone, 4 g yeast extract, 5 g sodium acetate, 2 g tri-ammonium citrate, 0.2 g MgSO4·7H2O, 0.05 g MnSO4·7H2O; and 1 ml Tween 80.

Extraction of L. reuteri 35-5 whole cell proteins

The cells from different L. reuteri 35-5 cultures (10 ml) grown with various carbon sources were collected by centrifugation (10,000 ×g) and then suspended in 200 µl B-PER protein extraction buffer (Thermo, Rockford, US). The suspensions were frozen in liquid nitrogen and thawed in a 37 °C water bath for three times. Then, 2 µl lysozyme (150,000 U) was added to the suspension. After 1 h incubation at room temperature, 10 µl of each suspension was boiled with 5 µl SDS loading buffer for 10 min. After centrifugation (12,000 ×g, 5 min), the supernatants were analyzed by SDS-PAGE.

Activity staining of EPS synthesizing enzymes

After electrophoresis, the gels with L. reuteri 35-5 proteins were washed 3 times with dd H2O (30 min each), allowing protein renaturation. Then, the gels were incubated overnight at 37°C in sodium acetate buffer (25 mM Tris/HCl, pH 5.0, 1 mM CaCl2) with 5% (w/v) sucrose, 5% (w/v) maltodextrins or 2.5% (w/v) sucrose plus 2.5% (w/v) maltodextrins. The activities of EPS synthesizing enzymes were detected by staining of the polysaccharides by a periodic acid-Schiff (PAS) procedure (50). The full-length GtfB expressed and purified from E. coli was loaded as a reference.

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Extraction of exopolysaccharides (EPS)

EPS were produced by L. reuteri 35-5 in the sugar-free MRS medium supplemented with sucrose (5%, w/v), or maltodextrins (DE = 13-17, 5%, w/v), or sucrose (2.5%, w/v) plus maltodextrins (2.5 %, w/v). Strains were grown anaerobically for 72 h at 37 °C. EPS were extracted from supernatants obtained by centrifugation of liquid MRS cultures (30). Two volumes of cold ethanol (-20 °C) were added to the supernatants and held at 4 °C overnight. The precipitates were harvested by centrifugation (12,000 ×g, 30 min, 4 °C) and re-dissolved in 1 volume of dd H2O. The EPS were re-precipitated with 2 volumes of ethanol at 4 °C overnight. After centrifugation, the harvested precipitates were dissolved in water, and then dialyzed (10 kDa MWCO, Thermo Scientific) against water for 48 h with changes of dd H2O every 12 h followed by freeze drying (31).

Size exclusion chromatography (SEC)

DMSO-LiBr (0.05M) was prepared by stirring for 3 h at room temperature followed by degassing for 15 min using an ultrasonic cleaner (Branson 1510, Branson, Danbury, CT). Samples were dissolved at a concentration of 4 mg/ml in DMSO-LiBr by overnight rotation at room temperature, followed by 30 min heating in an oven at 80 °C, obtaining clear sample solutions. The samples, cooled to room temperature, were filtered through a 0.45-µm Millex PTFE membrane (Millipore Corporation, Billerica, MA). The SEC system set-up (Agilent Technologies 1260 Infinity) from PSS (Mainz, Germany) consisted of an isocratic pump, auto sampler, an online degasser, an inline 0.2 μm filter, a refractive index detector (G1362A 1260 RID Agilent Technologies), viscometer (ETA-2010 PSS, Mainz) and MALLS (SLD 7000 PSS, Mainz). WinGPC Unity software (PSS, Mainz) was used for data processing. Samples (100 µl) were injected into a PFG guard column using an autosampler at a flow rate of 0.5 ml/min and DMSO-LiBr as eluent. Three separate PFG-SEC columns with porosities of 100, 300 and 4000 Å were used. The columns were held at 80˚C, the refractive index detector at 45 °C and the viscometer was thermostatted at 60 °C. A standard pullulan kit (PSS, Mainz, Germany) with molecular masses from 342 to 805000 Da was used. The specific RI increment value dn/dc was measured by PSS and is 0.072 (obtained from PSS company).

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Enzymatic hydrolysis of EPS

The EPS samples (5 mg) were hydrolyzed by pullulanase M1 (Megazyme, Ireland) based on the protocol established previously for the L. reuteri 35-5 EPS derived from sucrose (25). The hydrolyzed samples were subjected to HPAEC analysis.

Monosaccharide analysis

EPS samples (0.5 mg) were dissolved in 200 µl Milli-Q water in a glass reaction tube. An equal volume of 4 M trifluoroacetic acid (TFA) was added to the samples. The samples were hydrolyzed for 4 h at 100 °C. Samples were then dried under a flow of dry nitrogen. The dry samples were dissolved in 100 µl isopropanol and dried again by evaporation under dry nitrogen. Samples were dissolved in 1 ml dimethyl sulfoxide (DMSO) and were subjected to HPAEC analysis.

Expression and purification of GtfA-∆N, GtfB-∆N and full-length GtfB

The N-terminally truncated GtfA-∆N and GtfB-∆N enzymes were produced as described before (32, 33). The full-length GtfB protein was produced in E. coli BL21 Star (DE3) carrying the plasmids pRSF-GtfB and pBAD22-GroELS as described previously (21). All three enzymes were first purified by His-tag affinity chromatography using a 1-ml Hitrap IMAC HP column (GE Healthcare) followed by anion exchange purification using Hitrap Q FF (GE Healthcare). Purity was checked on 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Protein concentration was measured by Bradford reagent (Bio-Rad, Hercules, CA) using bovine serum albumin as standard.

Enzymatic reactions

All reactions were performed in sodium acetate buffer (25 mM, pH 4.7, 1 mM CaCl2) at 37 °C for 4 h. GtfA-∆N (750 nM) and/or GtfB-∆N (150 nM) were used in the reactions with 0.1 M sucrose and/or 0.25% amylose V. After 4 h incubation, the reactions were terminated by heating the samples in boiling water for 10 min. Then the products were subjected to TLC and high-pH anion-exchange chromatography (HPAEC) analyses and further separation.

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Amylose-iodine assay

The iodine staining method was used to quantify (remaining) amylose in the reaction mixture in order to determine the degradation rate of amylose V in the absence or presence of different acceptor substrates (34). It was also used for characterization of the GtfB enzyme (33). I2 (0.26 g) and KI (2.6 g) were dissolved in 10 ml distilled water to prepare the 260× concentrated stock. Prior to use, the stock was diluted 260× with distilled water. For each measurement, 150 µl iodine reagent was pipetted into microtiter plate wells, followed by addition of 15 µl enzyme reaction sample solution. The microtiter plate was shaken slowly for 5 min to allow color formation and the optical density was measured at 660 nm. Amylose V solutions (0-0.25%, w/v) were used to prepare a calibration curve. The acceptor reactions were performed using 0.125 % (w/v) amylose V as donor substrate and the activity was measured as described previously (33).

Separation of oligosaccharides

The individual oligosaccharides produced by GtfA-∆N (and GtfB-∆N) enzymes from sucrose or sucrose plus amylose V were isolated by a combination of Bio-Gel P2 size exclusion chromatography followed by preparative HPAEC. The reaction product mixtures were loaded onto the Bio-Gel P2 size exclusion column (100 x 0.9 cm) (Bio-Rad) using ammonium bicarbonate (10 mM) as eluent at a rate of ~10 ml/h. The oligosaccharide fractions were pooled and each pool was further fractionated by HPAEC-PAD on a Dionex DX500 workstation (Dionex, Amsterdam, The Netherlands), equipped with a CarboPac PA-1 column (250×9mm; Dionex) and an ED40 pulsed amperometric detector, using a linear gradient of 0-500mM sodium acetate in 100 mM NaOH (3ml/min). Collected fractions were immediately neutralized with 4 M acetic acid, desalted on CarboGraph SPE columns (Alltech, Breda, The Netherlands) using acetonitrile:water = 1:3 as eluent and lyophilized (35). The isolated oligosaccharides were subjected to MALDI-TOF-MS and NMR analysis as described below.

Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry

(MALDI-TOF-MS)

The degree of polymerization of isolated oligosaccharides was determined on an AximaTM mass spectrometer (Shimadzu Kratos Inc., Manchester, UK) equipped

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with a nitrogen laser (337 nm, 3 ns pulse width). Positive-ion mode spectra were recorded using the reflector mode at a resolution of 5000 FWHM and delayed extraction (450 ns). The accelerating voltage was 19 kV with a grid voltage of 75.2%; the mirror voltage ratio was 1.12, and the acquisition mass range was 200-7000 Da. Samples (1 µl) were mixed in 1:1 ratio with a solution of 10 mg/ml 2,5-dihydroxylbenzoic acid in 1:1 acetonitrile:H2O.

Nuclear Magnetic Resonance (NMR) spectroscopy

The oligosaccharides produced by GtfA-∆N plus GtfB-∆N were desalted and exchanged twice with D2O (99.9 atm% D, Cambridge Isotope Laboratories, Inc.) with intermediate lyophilization. The dried samples were dissolved in 0.6 ml D2O and were filled in NMR tube for analysis. Resolution-enhanced 1D/2D 500-MHz

1H NMR spectra were recorded with a spectral width of 4500 Hz in 16k complex data sets and zero filled to 32k in D2O on a Varian Inova Spectrometer (NMR Center, University of Groningen) or a Bruker DRX-500 spectrometer (Bijvoet Center, Department of NMR Spectroscopy, Utrecht University) at a probe temperature of 300/310 K. Suppression of the HOD signal was achieved by applying a WET1D pulse sequence. The 2D TOCSY spectra were recorded with spin-lock times of 20–200 ms using an MLEV-17 (composite pulse devised by (36)) mixing sequence. The 2D ROESY spectra were recorded with a mixing time of 200 ms using the standard Bruker XWINNMR software. The carrier frequency was set at the downfield edge of the spectrum to minimize TOCSY transfer during spin-locking. 2D 13C–1H HSQC experiments (1H frequency 500.0821 MHz, 13C frequency 125.7552 MHz) were recorded without decoupling during acquisition of the 1H free induction decay. Resolution enhancement of the spectra was performed by a Lorentzian-to-Gaussian transformation for 1D spectra or by multiplication with a squared-bell function phase shifted by π/(2.3) for 2D spectra. Chemical shifts (δ) are expressed in ppm by reference to internal acetone (δ 2.225 for 1H and δ 31.07 for 13C). MestReNova 5.3 was used for spectra annotation.

High-pH anion-exchange chromatography (HPAEC)

The reaction products of GtfA-∆N, GtfB-∆N and GtfA-∆N plus GtfB-∆N, and the hydrolysis products of EPS, were analyzed on a Dionex DX500 workstation (Dionex) with a 4×250 nm CarboPac PA-1 column. Samples were run with a

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gradient of 30-600 mM NaAc in 100 mM NaOH (1 ml/min) and detected by an ED40 pulsed amperometric detector.

Results and Discussion

4,6-α-Glucanotransferase encoding genes occur concomitantly with

glucansucrase encoding genes in lactic acid bacteria

The purified 4,6-α-GTase GtfB enzyme from L. reuteri 121 (strain 35-5) has been shown to produce IMMP in vitro. L. reuteri 121 (strain 35-5) cells grown on starch in vivo produce EPS that are structurally similar to IMMP; the extracellular and cell-associated GtfB is essential for IMMP-like EPS formation (Bai et al. submitted). Similar 4,6-α-GTase enzymes have been identified and characterized (GtfW and GtfML4) in other L. reuteri strains (22, 33, 37). To date, 28 gene sequences encoding characterized or putative 4,6-α-GTase enzymes (>70% similarity with GtfB enzyme) from Lactobacillus species are present in the NCBI database (http://www.ncbi.nlm.nih.gov/BLAST/). In 8 cases, the same strain also carries a gene encoding a characterized or putative glucansucrase (GS) (Table 1) (38, 39); GS and 4,6-α-GTase genes are located next to each other in 3 L. reuteri strains. These GS are known to convert sucrose into homo-EPS (α-glucan), e.g. the GS GtfA enzyme from L. reuteri 121 (strain 35-5) synthesizes reuteran with alternating α1→4 and α1→6 linkages (25, 40, 41). With larger numbers of 4,6-α-GTase encoding genes rapidly accumulating, more combinations of GS and 4,6-α-GTase enzymes are likely to be discovered in time.

Table 1. Combinations of 4,6-GTase and GS encoding genes occurring in LAB strains. The sequences were retrieved from NCBI database and are labelled by Genebank number.

Strains 4,6-α-GTase GS

Lactobacillus reuteri 121 AAU08014.2 AAU08015.1 Lactobacillus reuteri TMW1.106 ABP88725.1 ABP88726.1

Lactobacillus reuteri ML1 AAU08003.2 AAU08004.1 Lactobacillus reuteri TMW1.656 KOF04763.1 KEK16955.2 Lactobacillus delbrueckii ND02 WP_041812193.1 ADQ61508.1

Lactobacillus plantarum 16 WP_041161886.1 AGO09550.1 Lactobacillus salivarius GJ-24 EGM52218.1 EGM52203.1

Weissella confusa MBF8-1 ACJ54706.1 ACJ64929.1

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Activity staining of GtfA and GtfB enzymes of Lactobacillus reuteri 35-5

Periodic acid-Schiff (PAS) staining was used to visualize enzymes producing polysaccharides present in cells of L. reuteri 35-5. The PAS method has been used successfully to screen for GS positive Lactobacillus strains as denatured GS can be refolded and regain activity in SDS-PAGE gels under mild conditions (42). Following SDS-PAGE of L. reuteri 35-5 whole cell proteins the gels were washed with dd H2O and reaction buffer, and incubated with either maltodextrins (DE = 13.0-17.0), sucrose, or both, to show EPS formation by different enzymatic reactions. Cells of L. reuteri 35-5 grown on different substrates constitutively expressed an active GtfA protein (199 kDa; Fig. 1, upper panel) (24). The other gel band (~120 kDa) most likely is due to (proteolytic) degradation of GtfA (43). It has been shown that removal of the N-terminal region, involved in cell wall anchoring, does neither affect the overall activity of the GtfA enzyme nor the linkage types and size of the polymer synthesized (26, 32, 44). Previously, we already reported that also the denatured GtfB enzyme can be easily renatured in dd H2O (37). After incubation of the gels with maltodextrins, a band at ~180 kDa became visible (Fig. 1, bottom panel), which reflects active GtfB protein (179 kDa). This band was most clearly visible in extracts of L. reuteri 35-5 cells grown on glucose and maltodextrins. As expected, the 199 kDa GtfA band was not visible: maltodextrins do not act as (donor) substrates for EPS formation by GtfA (Fig. 1, bottom panel) (28). The L. reuteri 121 genome sequence (Gangoiti et al., in preparation) analysis showed that GtfA and GtfB are the only two carbohydrate acting enzymes with molecular masses larger than 150 kDa. The combined results thus show that the visible band at ~180 kDa represents the GtfB enzyme (179 kDa). To better distinguish the bands representing GtfA and GtfB activities, gels also were incubated with both sucrose and maltodextrins. This resulted in appearance of two bands at the expected and clearly different positions, demonstrating that these two enzymes are independently active on their own preferred substrate for polysaccharide formation (Fig. 1, middle panel). The combined data thus shows that GtfA and GtfB enzymes are the only two enzymes contributing to homo-EPS formation in the presence of sucrose or maltodextrins/starch.

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Fig. 1. Periodic Acid-Schiff (PAS) staining of α-glucans synthesized in SDS-PAGE gels of whole-cell-extracted proteins of L. reuteri 35-5, visualizing the position of the renatured and active GS GtfA and 4,6-α-GTase GtfB enzymes. L. reuteri 35-5 was grown in MRS medium with glucose, sucrose, maltodextrins, or sucrose plus maltodextrins. Gels were incubated with sucrose (5%, w/v), maltodextrins (5%, w/v) or sucrose (2.5%, w/v) plus maltodextrins (2.5%, w/v) overnight at 37 °C, pH 4.7. Heterologously expressed and purified GtfB from E. coli was loaded as reference protein.

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SEC analysis of the products formed by L. reuteri 35-5 from sucrose,

maltodextrins, and sucrose plus maltodextrins

The GS GtfA mainly acts on sucrose as donor/acceptor substrate, resulting in chain length elongation and oligo-/polysaccharide synthesis. To some extent, GtfA also is able to use maltose/maltodextrins as acceptor substrate (with sucrose as donor substrate) (32). 4,6-α-GTase GtfB mainly acts on maltodextrins/starches, cleaving α1→4 linkages and making α1→6 linkages. Here, we report that, to some extent, GtfB also use sucrose as acceptor substrate (with amylose V as donor substrate) (Fig. 5). It is therefore entirely likely that the final EPS formed by L. reuteri 35-5 cells in the presence of both substrates is not just a mixture, but may have different structures and thus characteristics. The EPS produced by L. reuteri 35-5 from sucrose (5%, w/v), maltodextrins (5%, w/v), and sucrose (2.5%, w/v) plus maltodextrins (2.5%, w/v) were isolated from liquid cultures by ethanol precipitation and subjected to SEC analysis (Fig. 2). The α-glucan EPS derived from sucrose is of high molecular mass (MM) up to 34.7 MDa (24), whereas the majority (97.5%) of the EPS derived from maltodextrins is of low MM (Bai et al., submitted). Growth of L. reuteri 35-5 on sucrose plus maltodextrins resulted in synthesis of an EPS mixture with both high and low MM (Fig. 2) In view of the size distribution obtained, the EPS35-5-sucrose+maltodextrins appears to be composed of a mixture of the EPS35-5-sucrose and EPS35-5-maltodextrins. A slight difference was observed in the low MM region (at approx. 3.5 kDa) between the SEC chromatograms of EPS35-5-maltodextrins and EPS35-5-sucrose+maltodextrins (Fig. 2). The question remained whether the EPS structures at high and low MM changed when L. reuteri 35-5 was grown in the presence of both sucrose and maltodextrins.

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Fig. 2. SEC chromatograms of EPS isolated from MRS liquid cultures of L. reuteri 35-5 grown with sucrose (5%, w/v), maltodextrins (5%, w/v), or sucrose (2.5%, w/v) plus maltodextrin (2.5%, w/v). Elution volume of the pullulan standards corresponding to 366, 200, 113, 48.8, 21.7, 10, 6.2 and 1.32 kDa, are shown as gray dots. The arrow indicates the 3.5 kDa position that differs in the low MM region of the SEC chromatograms of EPS35-5-maltodextrins and EPS35-5-sucrose+maltodextrins.

Enzymatic hydrolysis of the products formed by L. reuteri 35-5 from sucrose,

maltodextrins, and sucrose plus maltodextrins

To explore structural differences among EPS derived from different substrates, these 3 EPS were hydrolyzed by an excess amount of pullulanase type I. This enzyme specifically attacks α1→6 linkages in the backbone chains of pullulan and at branching points of starch molecules (45). As reported, this pullulanase degraded the EPS35-5-sucrose (reuteran) into mainly maltose, panose, maltotriose and maltotetraose (25) (see Fig. 3). This pullulanase also degraded the EPS35-5-maltodextrins, resulting in a low yield of maltooligosaccharides and a trace amount of glucose. Interestingly, when the EPS35-5-sucrose+maltodextrins were hydrolyzed by pullulanase, glucose and some maltooligosaccharides were the main product. Interestingly also a series of new oligosaccharides were formed (eluting between 20 to 30 min) (Fig. 3). Obviously, the EPS35-5-sucrose+maltodextrins is not a simple mixture of EPS35-5-sucrose and EPS35-5-

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maltodextrins. L. reuteri 35-5 cells grown on MRS medium with sucrose plus maltodextrins synthesized EPS that are structurally different from those derived from either sucrose or maltodextrins.

Fig. 3. HPAEC analysis of the hydrolysis products of EPS (5 mg) digested by excess amounts of pullulanase M1 for 72 h. EPS were produced by L. reuteri 35-5 in MRS liquid cultures supplemented with sucrose (5%, w/v), maltodextrins (5%, w/v), and sucrose (2.5%, w/v) plus maltodextrins MD13-17 (2.5%, w/v) and isolated by ethanol precipitation.

In vitro reactions of GtfA-∆N or GtfB-∆N with sucrose and amylose V

As the L. reuteri 35-5 in vivo system for EPS formation is too complex to study directly, the purified GtfA-∆N and GtfB-∆N enzymes (26, 33) were used in vitro to study the initial formation of oligosaccharides from sucrose and/or maltodextrins. GtfA-∆N or GtfB-∆N were separately incubated with sucrose, amylose V and sucrose plus amylose V. Amylose V was used instead of maltodextrins to avoid interference between the substrate and the oligosaccharide products in the HPAEC analysis. As shown in Fig. 4, GtfA-∆N is inactive with amylose V as no degradation products resulting from hydrolysis activity were visible in the HPAEC spectrum. We also failed to detect activity of GtfA-∆N with amylose V when using the amylose-iodine assay (data not shown). Sucrose

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is the preferred substrate for GtfA-∆N and the initial products have been annotated previously (Fig. 4), representing linear oligosaccharides with alternating α1→4 and α1→6 linkages (28). Addition of amylose V to the sucrose/GtfA-∆N incubation mixture also did not result in differences in the oligomer product profile. Unlike maltose, amylose V may be too large to act as acceptor substrate for sucrose/GtfA-∆N reaction. Addition of amylose V, however, may influence the formation of polysaccharides, resulting in formation of new α1→6 and α1→4 linkages at the non-reducing ends of the amylose V substrate.

Fig. 4. HPAEC spectra of the products of GtfA-∆N (750 nM) incubated with 0.1 M sucrose or/and 0.25% (w/v) amylose V for 2 h at 37 °C and pH 4.7. The oligosaccharide structures were annotated as in Dobruchowska et al (28).

GtfB is inactive on sucrose, but acts on amylose V, cleaving α1→4 linkages and making α1→6 linkages, which also results in the formation of glucose and maltose (Fig. 5), as reported previously (23). In the presence of both sucrose and amylose V, GtfB-∆N uses sucrose as acceptor substrate and elongates it with one glucosyl unit, introducing either an α1→4 or α1→6 linkage (predominant), using amylose V as donor substrate. Almost no glucose was generated from both sucrose and amylose V, indicating that the (minor) hydrolysis activity of GtfB-∆N was strongly suppressed.

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Fig. 5. HPAEC analysis of the products of GtfB-∆N (150 nM) incubated with 0.1 M sucrose or/and 0.25% (w/v) amylose V for 2 h at 37 °C and pH 4.7. Peaks are annotated on the basis of retention times of known controls (28).

In vitro reactions of GtfA-∆N plus GtfB-∆N with sucrose and/or amylose V

The purified GtfA-∆N plus GtfB-∆N enzymes were incubated with sucrose, amylose V, or sucrose plus amylose V, to mimic the in vivo synthesis of EPS by L. reuteri 35-5 cells. When incubating both enzymes with the single substrates, similar product profiles were obtained as seen with the single enzymes (Fig. 6). Incubation of both enzymes with both substrates, however, resulted in clear differences. The amount of oligomers with an α1→6 linked non-reducing end (peaks 1, 4, and 5 in Fig. 6) increased compared to those derived from sucrose as sole substrate.

In addition, several new products were formed (e.g. peaks I and II) (Fig. 6). Both fractions were found to contain compounds of a single DP (MALDI-TOF-MS analysis; DP6, data not shown).

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Fig. 6. HPAEC analysis of the products of GtfA-∆N (750 nM) and GtfB-∆N (150 nM) incubated with 0.1 M sucrose or/and 0.25% (w/v) amylose V for 2 h at 37 °C and pH 4.7. The oligosaccharide structures were annotated as in Dobruchowska et al (28). Two novel peaks labelled I and II were annotated based on1D/2D NMR analysis (Fig. 7).

NMR analysis of novel products synthesized by GtfA-∆N plus GtfB-∆N from

sucrose plus amylose V

The new products synthesized in vitro by GtfA-∆N plus GtfB-∆N from sucrose plus amylose V were analyzed by 1D and 2D NMR spectroscopy including TOCSY (150 ms). Starting from the anomeric signals of the residues g, f, A, B, E, J in the 2D TOCSY spectra (Fig. 7) all chemical shifts of the non-anomeric protons of differently substituted Glc residues could be assigned for two isomeric structures I and II (Table 2). Following the earlier developed structural-reporter-group concept rules (46), the set of chemical shifts of residue A (H-1 track, δ 5.380I/5.374II) was in agreement with an internal –(1→4)-α-D-Glcp-(1→4)- unit. Although the anomeric signals of B, E and J strongly overlap, the differences in chemical shifts of their non-anomeric protons could be deduced from the TOCSY built-up series of mixing times (20 ms, 60 ms, 80 ms, 150 ms). The TOCSY B H-1 track (δ 4.961) showed the set of chemical shifts corresponding to the values found for internal –(1→4)-α-D-Glcp-(1→6)- unit; that of residue E (δ 4.955I/4.947II) with a terminal α-D-Glcp-(1→6)- unit; that of residue J (δ 4.967I/II) with an internal –(1→6)-α-D-Glcp-(1→6)- unit (Table 2). Typical chemical shift

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values for sucrose fragment (Glc, g; Fru, f) are δ 5.427, stemming from Glc g H-1, and δ 4.220, stemming from Fru f H-3 (28). The small differences in chemical shift between residue AI H-1 and AII H-1, are due to proximity of A to the sucrose fragment and/or terminal residue E. Additional assignments and confirmation of the assignments were obtained from ROESY cross-peaks and by correlating the 1H resonances to the corresponding 13C resonances in the HSQC spectra (data not shown). These two novel oligosaccharides (DP = 6) with internally or terminally consecutive α1→6 linkages are structurally different from all the reported oligosaccharides produced by the single GtfA or GtfB enzymes from their preferred substrates.

In vitro incubations of GtfA-∆N and GtfB-∆N with sucrose plus amylose V clearly affected initial oligosaccharide synthesis (Figs. 4-7). This reflects the cross-activity of the cell-associated GtfA and GtfB enzymes in EPS formation in vivo, also resulting in modified product structures (Fig. 3).

Fig. 7. 1D 1H NMR and TOCSY analysis of the DP6 compounds in peaks I and II (Fig. 6).

Table 2. 1H chemical shiftsa (D2O, 300K) of linear oligomers (DP6) labelled as I and II in Figs. 6 and Fig. 7.

Residue H-1a H-1b H-2 H-3 H-4 H-5 H-6a H-6b

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g –(1→6)-α-D-Glcp-

(1↔2)-β-D-Fruf 5.427 - 3.57 3.74 3.53 4.05 3.68 -

f -(1↔2)-β-D-Fruf

3.66 3.66 - 4.22 4.05 3.89 3.84 3.77

A –(1→6)-α-D-Glcp-

(1→4)- 5.380I/5.374II - 3.59 3.67 3.49 3.93 3.73 3.97

B –(1→4)-α-D-Glcp-

(1→6)- 4.961I/II - 3.60 3.98 3.65 3.85 3.89 3.82

E α-D-Glcp-(1→6)-

4.955I/4.947II - 3.55 3.73 3.420 3.74 3.84 3.76

J –(1→6)-α-D-Glcp-

(1→6)- 4.967I/II - 3.58 3.70 3.50 3.89 3.75 3.98

aIn ppm relative to the signal of internal acetone (δ 2.225 for 1H)

Specificity of GtfB-∆N using GtfA-∆N products as acceptor substrates

The possible use of GtfA-∆N products from sucrose as acceptor substrates for GtfB-∆N activity with amylose V was studied in more detail. Specific oligosaccharides produced by GtfA-∆N from sucrose were isolated as described by Dobruchowska et al. and incubated with GtfB-∆N in the presence or absence of amylose V as a donor substrate (28). As shown in Fig. 8a, a DP7 oligomer (product 1) with an α1→6 linked non-reducing end can serve as acceptor substrate for GtfB-∆N in the presence of amylose V as donor substrate, resulting in formation of larger products. When compared to another GtfA-∆N product (product 2, DP6) with an α1→4 linked non-reducing end, product 1 with α1→6 linked non-reducing end is clearly preferred as acceptor substrate by GtfB-∆N incubated with amylose V (Fig. 8b). Product 1 is completely converted into a series of elongated products, while most of product 2 remains in the reaction. During polymer synthesis by GtfA-∆N from sucrose, GtfB-∆N thus is able to use the intermediate products with α1→6 linked non-reducing ends as acceptor substrates.

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Fig. 8. HPAEC spectra of products of GtfB-∆N (150 nM) incubated with isolated GtfA-∆N products in the presence or absence of amylose (0.25%, w/v) for 2 h at 37 °C and pH 4.7. The structures of products 1 and 2 were annotated according to Dobruchowska et al (28).

To further characterize the acceptor substrate specificity of GtfB-∆N, the transglycosylation factors (TF) with amylose V in the presence of absence of small acceptor substrates were determined (47-49). The preferred substrate for GtfA-∆N is sucrose, which serves as a strong acceptor substrate for GtfB-∆N, compared to glucose, maltose, and maltotriose. This is in accordance with the observation in Fig. 5. The TF for panose, a trisaccharide with an α1→6 linked non-reducing end, is higher than for maltotriose with an α1→4 linked non-reducing end (see Table 3), proving that acceptor substrates with a α1→6 linked non-reducing ends are stronger acceptors than those with α1→4 linked non-reducing ends.

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Table 3. Amylose V (0.25%, w/v) degradation activity (total activity, measured using iodine staining assay) (33) and transglycosylation factors of the L. reuteri 121 GTFB-∆N (60 nM) enzyme incubated with amylose V in the presence of different acceptor substrates (10 mM). Experiments were carried out in duplicate and the average data is given.

Amylose V degradation rate (U/mg) Transglycosylation factorsa

Without acceptor substrate

2.8 1.0

Maltoseb 7.3 2.6

Sucrose 10.4 3.7

Maltotrioseb 7.5 2.7

Panose 24.7 8.8

a The transglycosylation factor is defined as the ratio of amylose degradation rates in the presence/absence of an acceptor substrate. b The data refers to the paper published previously (33).

Conclusions In this study we have demonstrated that L. reuteri 35-5 with both GS and 4,6-α-GTase enzymes synthesizes a new type of EPS when grown with sucrose plus maltodextrins, compared to the EPS derived from either of the single carbon sources. Subsequent in vitro enzymatic reactions of GtfA-∆N plus GtfB-∆N with sucrose plus amylose V resulted in synthesis of new initial oligosaccharides with terminally and internally consecutive α1→6 linkages. These initially formed GtfA-∆N oligosaccharides are strong acceptor substrates for the GtfB-∆N enzyme when incubated with amylose V. This underpins the above observation that in vivo, GtfB/maltodextrins affect EPS synthesis by GtfA from sucrose. In the presence of starch plus sucrose, EPS synthesized by L. reuteri strain 35-5 cells resulted not only from the action of the GtfA and GtfB enzymes with their preferred substrates sucrose and maltodextrins, but also yielded a new type of EPS reflecting the cross-reactivity of both enzymes. EPS formation in the presence of sucrose plus starch thus may influence biofilm formation by lactobacilli in the oral cavity or gastrointestinal tract of mammals. Also the relationship between dietary intake of sucrose and starch, dental caries formation and degradability of EPS dietary fibers, are likely to be affected.

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Acknowledgements The study was financially supported by the Chinese Scholarship Council (to YB), the University of Groningen, and by the TKI Agri&Food program as coordinated by the Carbohydrate Competence Center (CCC-ABC; www.cccresearch.nl).

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10. van Hijum SAFT, Kralj S, Ozimek LK, Dijkhuizen L, van Geel-Schutten IGH. 2006. Structure-function relationships of glucansucrase and fructansucrase enzymes from lactic acid bacteria. Microbiol Mol Biol R 70:157-176.

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soluble dietary fiber made via enzymatic conversion of starch. J Agric Food Chem 62:12034-12044.

22. Leemhuis H, Dijkman WP, Dobruchowska JM, Pijning T, Grijpstra P, Kralj S, Kamerling JP, Dijkhuizen L. 2013. 4,6-α-Glucanotransferase activity occurs more widespread in Lactobacillus strains and constitutes a separate GH70 subfamily. Appl Microbiol Biot 97:181-193.

23. Kralj S, Grijpstra P, van Leeuwen SS, Leemhuis H, Dobruchowska JM, van der Kaaij RM, Malik A, Oetari A, Kamerling JP, Dijkhuizen L. 2011. 4,6-α-glucanotransferase, a novel enzyme that structurally and functionally provides an evolutionary link between glycoside hydrolase enzyme families 13 and 70. Appl Environ Microb 77:8154-8163. 24. Van Geel-Schutten GH, Faber EJ, Smit E, Bonting K, Smith MR, Ten Brink B, Kamerling JP, Vliegenthart JFG, Dijkhuizen L. 1999. Biochemical and structural characterization of the glucan and fructan exopolysaccharides synthesized by the Lactobacillus reuteri wild-type strain and by mutant strains. Appl Environ Microb 65:3008-3014.

25. van Leeuwen SS, Kralj S, van Geel-Schutten IH, Gerwig GJ, Dijkhuizen L, Kamerling JP. 2008. Structural analysis of the α-D-glucan (EPS35-5) produced by the Lactobacillus reuteri strain 35-5 glucansucrase GTFA enzyme. Carbohyd Res 343:1251-1265.

26. Kralj S, van Geel-Schutten GH, Rahaoui H, Leer RJ, Faber EJ, van der Maarel MJEC, Dijkhuizen L. 2002. Molecular characterization of a novel glucosyltransferase from Lactobacillus reuteri strain 121 synthesizing a unique, highly branched glucan with α-(1→4) and α-(1→6) glucosidic bonds. Appl Environ Microb 68:4283-4291. 27. Dobruchowska JM, Gerwig GJ, Kralj S, Grijpstra P, Leemhuis H, Dijkhuizen L, Kamerling JP. 2012. Structural characterization of linear isomalto-/malto-oligomer products synthesized by the novel GTFB 4,6-α-glucanotransferase enzyme from Lactobacillus reuteri 121. Glycobiology 22:517-528. 28. Dobruchowska JM, Meng XF, Leemhuis H, Gerwig GJ, Dijkhuizen L, Kamerling JP. 2013. Gluco-oligomers initially formed by the reuteransucrase enzyme of Lactobacillus reuteri 121 incubated with sucrose and malto-oligosaccharides. Glycobiology 23:1084-1096. 29. Van Geel-Schutten GH, Faber EJ, Smit E, Bonting K, Smith MR, Ten Brink B, Kamerling JP, Vliegenthart JFG, Dijkhuizen L. 1999. Biochemical and structural characterization of the glucan and fructan exopolysaccharides synthesized by the Lactobacillus reuteri wild-type strain and by mutant strains. Appl Environ Microb 65:3008-3014.

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30. van Geel-Schutten GH, Flesch F, ten Brink B, Smith MR, Dijkhuizen L. 1998. Screening and characterization of Lactobacillus strains producing large amounts of exopolysaccharides. Appl Microbiol Biotechnol 50:697-703.

31. Sims IM, Frese SA, Walter J, Loach D, Wilson M, Appleyard K, Eason J, Livingston M, Baird M, Cook G, Tannock GW. 2011. Structure and functions of exopolysaccharide produced by gut commensal Lactobacillus reuteri 100-23. ISME J 5:1115-1124. 32. Kralj S, van Geel-Schutten GH, van der Maarel MJEC, Dijkhuizen L. 2004. Biochemical and molecular characterization of Lactobacillus reuteri 121 reuteransucrase. Microbiol-SGM 150:2099-2112. 33. Bai Y, van der Kaaij RM, Leemhuis H, Pijning T, van Leeuwen SS, Jin Z, Dijkhuizen L. 2015. Biochemical characterization of Lactobacillus reuteri Glycoside Hydrolase family 70 GTFB type of 4,6-α-Glucanotransferase enzymes that synthesize soluble dietary starch fibers. Appl Environ Microbiol 81:7223-7232. 34. Cochran B, Lunday D, Miskevich F. 2008. Kinetic analysis of amylase using quantitative Benedict's and iodine starch reagents. J Chem Educ 85:401-403. 35. Meng XF, Dobruchowska JM, Gerwig GJ, Kamerling JP, Dijkhuizen L. 2015. Synthesis of oligo- and polysaccharides by Lactobacillus reuteri 121 reuteransucrase at high concentrations of sucrose. Carbohyd Res 414:85-92. 36. Levitt MH, Freeman R, Frenkiel T. 1982. Broad-band heteronuclear decoupling. J Magn Reson 47:328-330.

37. Bai Y, van der Kaaij RM, Woortman AJJ, Jin Z, Dijkhuizen L. 2015. Characterization of the 4,6-α-glucanotransferase GTFB enzyme of Lactobacillus reuteri 121 isolated from inclusion bodies. BMC Biotechnol 15:49.

38. Malik A, Radji M, Kralj S, Dijkhuizen L. 2009. Screening of lactic acid bacteria from Indonesia reveals glucansucrase and fructansucrase genes in two different Weissella confusa strains from soya. FEMS Microbiol Lett 300:131-138.

39. Kaditzky SB, Behr J, Stocker A, Kaden P, Ganzle MG, Vogel RF. 2008. Influence of pH on the formation of glucan by Lactobacillus reuteri TMW 1.106 exerting a protective function against extreme pH values. Food Biotechnol 22:398-418.

40. Arskold E, Svensson M, Grage H, Roos S, Radstrom P, van Niel EW. 2007. Environmental influences on exopolysaccharide formation in Lactobacillus reuteri ATCC 55730. Int J Food Microbiol 116:159-167.

41. Leathers TD, Cote GL. 2008. Biofilm formation by exopolysaccharide mutants of Leuconostoc mesenteroides strain NRRL B-1355. Appl Microbiol Biotechnol 78:1025-1031.

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42. Kralj S, van Geel-schutten GH, van der Maarel MJEC, Dijkhuizen L. 2003. Efficient screening methods for glucosyltransferase genes in Lactobacillus strains. Biocatal Biotransfor 21:181-187.

43. Fabre E, Bozonnet S, Arcache A, Willernot RM, Vignon M, Monsan P, Remaud-Simeon M. 2005. Role of the two catalytic domains of DSR-E dextransucrase and their involvement in the formation of highly α-1,2 branched dextran. J Bacteriol 187:296-303.

44. Van Geel-Schutten GH, Faber EJ, Smit E, Bonting K, Smith MR, Ten Brink B, Kamerling JP, Vliegenthart JF, Dijkhuizen L. 1999. Biochemical and structural characterization of the glucan and fructan exopolysaccharides synthesized by the Lactobacillus reuteri wild-type strain and by mutant strains. Appl Environ Microbiol 65:3008-3014. 45. Doman-Pytka M, Bardowski J. 2004. Pullulan degrading enzymes of bacterial origin. Crit Rev Microbiol 30:107-121. 46. van Leeuwen SS, Leeflang BR, Gerwig GJ, Kamerling JP. 2008. Development of a H-1 NMR structural-reporter-group concept for the primary structural characterisation of α-D-glucans. Carbohyd Res 343:1114-1119.

47. Jung JH, Jung TY, Seo DH, Yoon SM, Choi HC, Park BC, Park CS, Woo EJ. 2011. Structural and functional analysis of substrate recognition by the 250s loop in amylomaltase from Thermus brockianus. Proteins 79:633-644.

48. Tang SY, Yang SJ, Cha HJ, Woo EJ, Park C, Park KH. 2006. Contribution of W229 to the transglycosylation activity of 4-α-glucanotransferase from Pyrococcus furiosus. BBA-Proteins Proteom 1764:1633-1638.

49. Tachibana Y, Takaha T, Fujiwara S, Takagi M, Imanaka T. 2000. Acceptor specificity of 4-α-glucanotransferase from Pyrococcus kodakaraensis KOD1, and synthesis of cycloamylose. J Biosci Bioeng 90:406-409.

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Chapter 6

Crystal structure of the 4,6-α-glucanotransferase GtfB: Evidence

that dietary changes triggered the evolution of glucansucrases

from α-amylases

Yuxiang Bai1,3, Joana Gangoiti-Muñecas1, Bauke W. Dijkstra2, Lubbert Dijkhuizen1, Tjaard Pijning2,*

1 Microbial Physiology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, The Netherlands

2 Laboratory of Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, Nijenborgh 7, 9747 AG Groningen, The Netherlands

3 The State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi 214122, China

In preparation for submission

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Abstract Dental caries is caused by oral bacteria adhering themselves to the tooth enamel, promoted by the formation of biofilms. A major constituent of biofilms are extracellular polysaccharides (EPS) synthesized by glucansucrase (GS) enzymes from sucrose (yielding α-glucans). Since sucrose is a relative recent addition to the human diet, the evolutionary origin of GS is intriguing and may help to understand the origin of caries. GS are classified in glycoside hydrolase family 70 (GH70); it has been proposed that they evolved from GH13 α-amylases. If true, evolutionary events would have had to involve changes in reaction- and substrate specificity; however the details of such changes, and which factors would trigger them, are unclear. Interestingly, a GH70 subfamily with 4,6-α-glucanotransferase (4,6-α-GTase) activity “intermediate” of α-amylases and GS has been discovered. In this study we provide 3D structural, genomic and phylogenetic data supporting the evolution of α-amylases to GS via 4,6-α-GTases. The crystal structure of the 4,6-α-GTase GtfB-ΔNΔV from Lactobacillus reuteri 121 reveals how the α-amylase substrate binding scaffold is retained, but different enough to favor transglycosylation instead of degradation. It also explains how GS could have evolved further towards sucrose utilization instead of starch. In vivo, 4,6-α-GTase and GS gene products within the same Lactobacillus strain complement each other in EPS synthesis, depending on the available carbon source. We propose that the evolution of GS in oral bacteria was triggered by dietary changes, in order to expand the reaction specificity and fermentable substrate specificity.

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Introduction Dental caries, a bacterial infection of the tooth enamel, poses a worldwide health problem (1, 2). It is caused by plaque-forming bacteria that ferment dietary carbohydrates in the oral cavity, and, by doing so, produce acids that dissolve calcium phosphate in the enamel. Adhesion of the oral bacteria to the tooth enamel is thought to be promoted by the formation of biofilms, which is facilitated by the synthesis of extracellular polysaccharides (EPS) (3-6). EPS are synthesized by glucansucrase (GS) enzymes, from sucrose (7-9), a widely accepted but relative recent addition to the human diet (yielding α-glucans). In this light, the evolutionary origin of GS from oral bacteria such as Lactobacillus and Streptococcus species, has raised interest, as it may help to understand the origin of dental caries. Notably, GS, belonging to the glycoside hydrolase family 70 (GH70; www.cazy.org (10), share sequence homology with α-amylases of GH13, which are also widely distributed in oral bacteria. The core domains of GS and α-amylases have similar folds; however the GS have a circularly permuted (β/α)8 barrel in the catalytic domain, and contain auxiliary domains which are absent in α-amylases (11, 12). Moreover, oral α-amylases act on starch-derived oligosaccharides (OS) instead of sucrose. Thus, if GS have evolved from α-amylases, a change in reaction type specificity (from substrate degradation to polymerization) and a change in substrate specificity (from starch to sucrose) would be two key evolutionary achievements. Vujičić-Žagar et al. (12) proposed that intermediate/precursor enzymes attained circular permutation by duplication of a common ancestor α-amylase gene, and that auxiliary domains originate from insertion events; however, little is known about how GS evolved to be able to use sucrose for the synthesis of EPS, and which factors would drive such evolutionary pathways.

Recently we identified 4,6-α-glucanotransferase enzymes (4,6-α-GTases) in Lactobacillus species. These enzymes belong to GH70 and share 45-50% sequence identity with GS, but phylogenetically clearly form a small subfamily, with notable differences occurring near the active site and in loop lengths (13, 14). Biochemical characterization of 4,6-α-GTases showed that in vitro they do not act on sucrose, but instead, like α-amylases, process starch-derived OS; on the other hand their product specificity does resemble the GS in that they synthesize α-glucan oligo-/polysaccharides with a high degree of α1→6 linkages (13-17).

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These 4,6-α-GTases thus have an activity with characteristics from both α-amylases and GS. Whether this activity reflects the role of these enzymes in vivo,remains to be established.

Here we provide 3D structural, genomic and phylogenetic data supporting an evolutionary pathway in which α-amylases evolved to GS via 4,6-α-GTase-like intermediates, with the latter two sharing a common ancestor closely related to subfamily 5 of GH13 α-amylases. The unique active site architecture of the 4,6-α-glucanotransferase GtfB-ΔNΔV from L. reuteri 121 retains an α-amylase-like scaffold, with modifications that explain the preference for transglycosylation and synthesis of polymer products. The 4,6-α-GTase architecture is also different from that of GS and shows which structural changes in the latter are related to shifting towards sucrose utilization. In vivo, depending on the available carbohydrate source (starch-derived OS or sucrose), GS GtfA and 4,6-α-GTase GtfB of Lactobacillus reuteri strain 121 both synthesize α-glucans with α1→4 and α1→6 linkages. We propose that oral bacteria such as Lactobacilli have evolved to expand their reaction specificity and fermentable substrate specificity, driven by changes in carbohydrate uptake in the oral cavity.

Materials and Methods

Cloning, truncation and site-directed mutagenesis, expression, purification

and characterization

The cloning, mutagenesis and truncation stratey of gtfB-ΔN and gtfB-ΔNΔV are described in SI text. Expression and purification of gtfB-ΔN was performed as described by Bai et al. (15); expression and purification of GtfB-ΔNΔV and its mutants followed a similar procedure with a modification as shown below. The single mutant Y1521A and double mutant W790A/W811A were constructed to study the possible role of aromatic stacking interactions of remote sugar binding sites; mutation T920W aimed at blocking donor subsites -2/-3 in the tunnel. Finally, mutants K1128A/N/H/W and Y1055G were constructed to investigate the importance of these residues near subsites -1/+1 and subsite +2, respectively.

After His-tag affinity and anion exchange chromatography, the proteins were purified by gel filtration (Superdex 200 10/300 GL) in sodium acetate buffer (25

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mM, pH 5.0, 150 mM NaCl, 1 mM CaCl2). Dynamic light scattering experiments of wild-type GtfB-ΔNΔV were performed on a Wyatt NanoStar (Wyatt Technology, Santa Barbara, US) at 293 K.

Crystallization and soaking experiments

Attempts to crystallize GtfB and GtfB-ΔN were unsuccessful. For GtfB-ΔNΔV, hanging drop vapour diffusion crystallization experiments were setup at 293 K using drops containing 1.5 μl of protein solution (4-5 mg/ml GtfB-ΔNΔV, 25 mM HAc/NaAc buffer, pH 5.0, 0.15 M NaCl, 1 mM CaCl2) mixed with 1.5 μl precipitant solution (17-20% (w/v) PEG 3350, 0.1 M (NH4)2SO4, 0.1 M NaCl) equilibrated against 0.5 ml of precipitant solution. To avoid excessive nucleation, a microseeding protocol was used in which 0.3 μl of diluted seed stock was introduced to hanging drops containing 1.5 μl of protein solution (as above) mixed with 1.5 μl of a solution containing 31% (w/v) PEG 3350, 0.1 M (NH4)2SO4, 0.1 M NaCl). Crystals of GtfB-ΔNΔV and its mutants tended to grow in clusters, and were cryoprotected in stabilizing solution containing 35% (w/v) PEG 3350, 0.1 M (NH4)2SO4, 25 mM BisTris-HCl buffer pH 5.5, 25 mM HAc/NaAc buffer pH 5.0, 1 mM CaCl2 (supplemented with 10% (v/v) glycerol for the wild-type apo data set). For soaking experiments, wild-type GtfB-ΔNΔV crystals were soaked in stabilizing solution supplemented with 50 mM maltohexaoxe (4-10 minutes); GtfB-ΔNΔV D1015N crystals were soaked in stabilizing solution supplemented with 100 mM maltopentaose (8 min).

Data collection

Data sets for wild-type apo and D1015N/maltopentaose were collected at the European Synchrotron Radiation Facility (ESRF) at beamlines ID29 and ID23-1, respectively, at 100 K; details and processing statistics are given in Table S4. All data sets were processed with XDS (18) and AIMLESS (19). Crystals of GtfB-ΔNΔV and its mutants belong to space group C 2 and tended to show high mosaicity.

Structure determination

The structure of GtfB-ΔNΔV was determined by molecular replacement using PHASER (20), using domains A, B, C and IV of the crystal structure of L. reuteri 180 Gtf180-ΔN (PDB 3KLK) (12) as a search model. The asymmetric unit

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contained two molecules of GtfB-ΔNΔV (Fig. 1A). Rigid-body and restrained refinement was performed using REFMAC5 (21) within the CCP4 package (22). Cycles of restrained refinement were alternated with manual building in COOT (23); non-crystallographic symmetry (NCS) restraints were used throughout refinement. The last 11 or 10 C-terminal residues (including the His6-tag) were not visible in electron density. Carbohydrate ligands were included only in the last stages of refinement; when the ligands had comparable binding modes in both protein molecules, we only described the situation for protein chain A. Final models were validated using MolProbity (24). PDBeFold (25) was used to analyze structural homology to other GH13 and GH70 enzymes. Structure figures were prepared with PyMOL (Schrödinger, LLC).

Enzymatic assays

Enzyme activities of wild-type GtfB-ΔNΔV and mutants derived were measured based on an assay developed by Bai et al. (15). Final products were analyzed by TLC, 1H NMR and HPAEC. Reaction of GtfB-ΔN (and maltase) with a blocked substrate (BPNPG7) was done to investigate the endo-specificity of GtfB-ΔNΔV. Details of the reactions and the respective product analysis are given in SI text.

Modeling and docking

Structural superpositions between GtfB-ΔNΔV and related enzymes were done in PyMOL by fitting the Cα and Cβ atoms of the corresponding three catalytic residues. The following glucansucrase structures were used: Lactobacillus reuteri 180 Gtf180-ΔN complexed with sucrose or with maltose (PDBs: 3HZ3 and 3KLL) (12); Lactobacillus reuteri 121 GtfA-ΔN (PDB: 4AYG) (26); Streptococcus mutans Gtf-SI (PDB: 3AIE) (27). The structure of Bacillus halmapalus α-amylase complexed with a pseudononasaccharide (PDB: 2GJP) (28) and B. subtilis α-amylase E208Q complexed with maltopentaose (PDB 1BAG) (29) served as models for α-amylases.

The structure of the Bacillus subtilis α-amylase maltopentaose complex (PDB 1BAG) (29) served as a template to model donor substrate binding, to extend the donor substrate observed in the GtfB-ΔNΔV - G5 complex towards acceptor subsites. The structure of Bacillus circulans strain 251 CGTase covalent glucosyl-enzyme intermediate (PDB: 1CXL) (36) was used to construct a model

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of the glucosyl-enzyme intermediate in GtfB-ΔNΔV. Subsequently, a model for binding of the acceptor substrate panose (O-α-D-glucopyranosyl-(1,6)-O-α-D-glucopyranosyl-(1,4)-D-glucose) was constructed by placing the trisaccharide in subsites +1 to +3, guided by hydrogen bond interactions and available docking space. Further details of modeling procedures are given in SI text.

Genomic mapping of glucansucrase and 4,6-α-glucanotransferase

The locations of the genes encoding glucansucrase and 4,6-α-glucanotransferase proteins from Lactobacillus reuteri strains were obtained based on the graphics data in NCBI database.

Sequence alignments and phylogenetic analyses

MUSCLE (30) was used to align the amino acid sequences of full-length glucansucrases and 4,6-α-glucanotransferases, and included 4,6-α-glucanotransferases GtfB of Lactobacillus reuteri 121 (Uniprot entry Q5SBM0), GtfML4 of Lactobacillus reuteri ML1 (Q5SBN1), GtfW of L. reuteri DSM 20016 (A5VL73), and glucansucrases Gtf180 L. reuteri 180 (Q5SBN3), GtfA of L. reuteri 121 (Q5SBL9), GtfO of L. reuteri ATCC 55730 (Q4JLC7), GtfML1 of L. reuteri ML1 (Q5SBN0), Gtf-SI of Streptococcus mutans ATCC 700610 (P13470) and DSRE of Leuconostoc mesenteroides NRRL B-1299 (Q8G9Q2). The α-amylases of Bacillus licheniformis (P06278), B. halmapalus (O82839), Geobacillus stearothermophilus (P06279) and B. subtilis (P00691) were aligned separately. Alignment of homology loops A1, A2 and B was done using ESPript (31), adapted by structural superposition of the 4,6-α-glucanotransferase GtfB-ΔNΔV (this study), the glucansucrase Gtf180-ΔN (PDB 3KLK) (12) and the α-amylase from Bacillus licheniformis (PDB 1BLI) (32).

Following alignment of all characterized 4,6-α-GTases and GS, a phylogenetic was performed by the maximum likelihood method using MEGA version 6 (33).

A second phylogenetic analysis included all bacterial GH13 and GH70 enzymes; for the GH70 enzymes, domains A and B were first rearranged to ‘correct’ for the circular permutation of the (β/α)8 barrel (BNAN···ACBC→ACBCBNAN, where the subscript denotes either the N- or C-terminal segment). The rearranged GH70 sequences were then added to the alignment of all characterized GH13 members classified into subfamilies, followed by phylogenetic analysis.

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Homologues of full-length GtfB were searched using Protein-BLAST in the NCBI (http://www.ncbi.nlm.nih.gov) and UniProtKB (http://www.uniprot.org) databases.

In vivo experiments

Growth conditions of plate-grown Lactobacillus reuteri 35-5 are described in SI text. The extracellular polysaccharides (EPS) produced on agar plates were extracted and analyzed by monosaccharide analysis using HPAEC, size determination using SEC, and linkage determination using NMR. Details of the procedures are described in SI text.

Results

Overall structure

The 4,6-α-glucanotransferase GtfB from L. reuteri 121 could only be successfully crystallized after deletion of the N-terminal domain and domain V. The 1.80 Å resolution crystal structure of the resulting construct (GtfB-ΔNΔV, residues Met-762-1619-His6) shows the presence of four domains (A, B, C and IV; Fig. 1A). In the asymmetric unit, the two molecules can be superimposed almost perfectly with an r.m.s.d. of 0.27 Å; most of the crystal contacts between them are made by domains C, A and B lying adjacent to each other in opposite directions. In solution, the estimated molecular mass of 85 kDa observed in dynamic light scattering experiments suggests that GtfB-ΔNΔV is monomeric. The core domains A (catalytic), B and C of GtfB-ΔNΔV are also found in GH13 α-amylases, while the auxiliary domain IV is only found in GH70 glucansucrases. The other auxiliary domains V and N of L. reuteri 121 GtfB precede domain IV and likely extend away from the rest of the protein. Like in glucansucrases, the polypeptide chain follows a “U-course” forming first the N-terminal halves of domains IV, B and A, then domain C, and then the C-terminal halves of domains A, B and IV (Fig. 1A). In the catalytic domain A, the (β/α)8 barrel is circularly permuted as in other GH70 enzymes; this permutation leads to a different order of the four homology regions (II-III-IV-I) than in GH13 α-amylases but keeps their relative spatial positioning. At the interface of domains A and B, a deep pocket is observed which is surrounded by residues strictly conserved in GH13 and GH70 enzymes, including the proposed catalytic residues (see below). By homology,

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this pocket would harbor subsites -1 and +1 (nomenclature according to Davies_1997) (34) where catalysis takes place.

Fig. 1. (A) Overall structure of GtfB-ΔNΔV showing the two molecules in the asymmetric unit. From the N-terminal residue (Nt) the polypeptide forms the N-terminal segments of domains IV (yellow), B (green) and A (blue), then domain C (purple) and the C-terminal segments of domains A, B and IV towards the C-terminal residue (Ct). The location of the catalytic site at the interface of domains A and B is indicated. (B) Loop arrangement around the active site of GH13 α-amylases (left), 4,6-α-glucanotransferases (middle) and GH70 glucansucrases (right), at the border of the catalytic domain A (blue) and domain B (green), looking down the (β/α)8 barrel. In this view, acceptor substrate subsites are positioned left of the catalytic residues (shown in stick representation), and donor substrate subsites to their right. Loop A2 (red) is conserved in all three enzyme classes, but only in glucansucrases it forms a wall adjacent to the active site. Loop A1 (purple) and loop B (brown) are present only in 4,6-α-glucanotransferases and glucansucrases, but they are longer in the former, where they form a flap covering the space adjacent to the active site, forming a tunnel. In glucansucrases these two loops are shorter and leave the active site surrounding much more open. Structures used are Bacillus licheniformis α-amylase (PDB 1BLI) (32), Lactobacillus reuteri 121 GtfB-ΔNΔV (this work) and Lactobacillus reuteri 180 Gtf180-ΔN (PDB 3KLK) (12).

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Active site architecture – comparison with α-amylases and glucansucrases

GtfB-ΔNΔV shows structural homology with both GH13 α-amylases and GH70 glucansucrases, with differences in the arrangement of loops around the active site (Fig. 1B). Regarding α-amylases, the structurally closest homologue of GtfB-ΔNΔV is the α-amylase from B. licheniformis (PDB 1BLI (32)); superposition of domains A+B results in an r.m.s.d. of 2.10 Å. Although in GtfB-ΔNΔV the (β/α)8 barrel is circularly permuted and domain IV (together with other auxiliary domains) is inserted into domain B, a long groove similar to α-amylases resides at the interface of domains A and B. This groove bends at the position of the deep pocket; differences are mainly found in the loops surrounding this groove. First, GtfB-ΔNΔV has an extra helix-loop-helix subdomain between strand β4 and helix α5 of the domain A (β/α)8 barrel; the loop connecting the helices (residues 1139-1151 in GtfB-ΔNΔV, hereafer referred to as “loop A1”) forms a large protrusion towards domain B, which is absent in α-amylases. Second, a long loop connecting the N-terminal segments of domains IV and B (residues 905-924, “loop B”) folds over the groove to contact the first helix of the helix-loop-helix subdomain with its tip (residues 917-918); notably, this loop has no equivalent in α-amylases. Third, the loop connecting strand β7 and helix α8 of the (β/α)8 barrel (“loop A2”, residues 1430-1440), is slightly longer than in α-amylases and takes a different course, creating a slight protrusion adjacent to loop A1.

Comparison with the structurally closest glucansucrase homologue of GtfB-ΔNΔV, Gtf180-ΔN (from L. reuteri 180, PDB 3KLK) (12), shows that domains A+B can be superimposed with an r.m.s.d. of 0.84 Å; thus their overall structure is very similar (more so than for α-amylases). Nevertheless, differences between GtfB-ΔNΔV and Gtf180-ΔN are again observed in the three loops described before. Notably, loops A1 and loop B are longer in GtfB-ΔNΔV than in Gtf180-ΔN; together with a 20-30o rotation of the helix-loop-helix subdomain, this results in a much less open arrangement near the active site with respect to GS. In addition, loop A2 is 5 residues shorter in GtfB-ΔNΔV than in GS and therefore protrudes less from the protein surface.

Together, the above mentioned structural features of GtfB-ΔNΔV result in a unique active site architecture. While the “base” of the long α-amylase like groove is maintained, its different loop arrangement creates a tunnel which is

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covered by loops A1 and B, and lined by loop A2. Superposition with α-amylases indicates that the tunnel would harbor donor subsites -2 and -3 (see also below). In contrast to the situation in GS, access to these donor subsites is not restricted, because of a shorter loop A2. Sequence alignment of the regions containing loops A1, A2 and B (Fig. S2B) shows that other 4,6-α-GTases likely have the same unique loop arrangement.

At the amino acid residue level, six of the seven strictly conserved GH13 residues surrounding subsites -1 and +1 also found in GtfB-ΔNΔV (the three proposed catalytic residues D1015, E1053 and D1125 plus residues R1013, H1124 and D1479); the seventh one is replaced by a glutamine residue (Q1484) like in glucansucrases (Fig. 2A and S1A). The arrangement of residues around subsites -1 and +1 thus is very similar to both that of GH13 α-amylases and GH70 GS, but also reveals a few differences, mainly near subsite +1. First, a tyrosine residue (Y1055) replaces the tryptophan residue found in almost all GS (and α-amylases); furthermore, the region Q1126-R1127-K1128-N1129 following the transition state stabilizing residue (D1125) is different from bothcorresponding regions in GS and in α-amylases (Figs. S1 and S2).

Soaking experiments and modeling

We performed several soaking experiments using either crystals of the wild-type enzyme or of the inactive mutant D1015N, and a range of glucooligosaccharides (DP2-DP7) with different α1→4 and/or α1→6 linkage patterns, as well as acarbose. Ligand electron density was only observed in donor subsites, with the reducing end glucosyl unit always occupying subsite -1 or -2. The donor subsites are bordered by several loops (from domain A and B) which, with respect to the apo structure, did not show large movements upon ligand binding (positional differences <0.5 Å). In contrast to donor subsites, acceptor subsites (+1, +2 etc.) never revealed enough electron density to model a bound carbohydrate. Although it seems that there is enough space for a maltooligosaccharide chain to occupy acceptor subsites, we noticed that access to these sites may be hindered by symmetry-related neighbouring protein molecules. Below, the soaking experiments with maltopentaose and maltohexaose are described.

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(A)

(B) (C)

Fig. 2. Active site of 4,6-α-glucanotransferase L. reuteri 121 GtfB-ΔNΔV; molecule A of the asymmetric unit is shown with domains A, B and IV in blue, green and yellow, resepctively. (A) Apo GtfB-ΔNΔV showing residues surrounding the cleavage site; catalytic residues are labeled in bold. A glycerol molecule bound in the active site is shown with yellow carbon atoms. Dashed black lines indicate the tunnel structure. (B) Structure of the GtfB-ΔNΔV D1015N - maltopentaose complex with the pentasaccharide bound in subsites -1 to -5, and a maltose near remote subsite I (W790/W811 in domain IV) in the extension of the donor binding groove. The catalytic residues are indicated. (C) Structure of GtfB-ΔNΔV soaked with maltohexaose; the transglycosylation pentasaccharide product (64-α-D-glucosyl-maltotetraose) bound in subsites -2 to -6 has an α1→6 linked glucosyl unit at the non-reducing end (indicated by the asterisk).

Soaking of maltopentaose (G5) in crystals of the inactive mutant GtfB-ΔNΔV D1015N revealed the pentasaccharide bound in subsites -1 to -5 in both molecules of the asymmetric unit (Figs. 2B and S1B). Comparison with ligand-bound α-amylase structures such as those from B. halmapalus (PDB: 2GJP) (28)

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and B. licheniformis (PDB: 1BAG) (32) shows that G5 has a similar helical binding mode in GtfB-ΔNΔV. Notably, subsites -2 and -3 are shielded from the solvent by loops A1 and B. Most of the ligand-protein interactions are observed in subsites -1, -2 and -3; in particular, the reducing end glucosyl unit bound in subsite -1 has numerous hydrogen bond interactions with the conserved GH13/GH70 residues. In other donor subsites, residues from loops A1 and A2 (subsites -2, -3) and from loops in domain B (subsites -4, -5) provide most of the direct or water-mediated hydrogen bond interactions to the bound substrate.

In wild-type GtfB-ΔNΔV crystals soaked with maltohexaose (G6) we observed electron density for an isomalto/maltooligosaccharide occupying subsites -2 to -6 (Figs. 2C and S1C), adopting a similar conformation as G5. Interestingly, the non-reducing end glucosyl unit of the oligosaccharide bound in subsite -6 is α1→6-linked; it has a stacking interaction with Y1521, a residue which is conserved in all putative 4,6-α-GTases. In subsite -1, remaining positive electron density was visible, but not enough to convincingly model a complete glucosyl unit.

The soaking experiment with G5 revealed three more binding sites for carbohydrates. The first remote site is formed by two tryptophan residues in domain IV (W790 and W811) whose aromatic side chains form a hydrophobic stacking platform for the placed disaccharide maltose (Fig. 2B); remaining positive electron density suggested that the actual bound ligand is longer but too flexible to be fully observable. The bound maltose mostly stacks with W811 and lies more or less in the extension of the donor tunnel at a distance of about 16 Å from subsite -6, corresponding to 2 or 3 glucosyl units. Notably, the tryptophan pair is unique to and conserved in 4,6-α-GTases; in most GS only one of the two tryptophan residues is conserved, while α-amylases lack both due to the absence of domain IV. Two more remote sites reside in domain A, near the loop connecting β6 to α7 and the loop connecting β7 to α8, and near helices α2 and α3; the maltose and a maltotriose ligand placed in electron density respectively only make few interactions with protein residues. Since the mentioned secondary structure elements are either non-conserved or semi-conserved in α-amylases and glucansucrases, these remote sites were not considered further.

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Modeling

We modeled donor substrate binding across the cleavage site by superposition with the maltopentaose bound in subsites -3 to +2 in B. subtilis α-amylase (PDB 1BAG) (35). Only minor adjustments of the glycosidic torsion angles were needed to model the glucosyl moieties in positive subsites (+1 and +2) without clashes in GtfB-ΔNΔV (Fig. 3A). Thus, maltooligosaccharides likely can bind in GtfB-ΔNΔV in a similar way as observed in α-amylases. Residues that may interact with sugar units at subsites +1 and +2 are N1019, E1053 (the catalytic acid/base), H1056, and possibly K1128. Hydrophobic interactions may be provided by Y1055 and L971. We also modeled acceptor binding by first constructing a model for a covalent glucosyl-enzyme intermediate at residue D1015, based on the experimentally observed structure of a CGTase covalent intermediate (36). In agreement with their high conservation of residues surrounding subsite -1, the GtfB-ΔNΔV covalent intermediate model is very similar. The model for panose (O-α-D-glucopyranosyl-(1,6)-O-α-D-glucopyranosyl-(1,4)-D-glucose) bound in this covalent glucosyl-enzyme intermediate model of GtfB shows the trisaccharide in subsites +1 to +3, with the non-reducing end glucosyl unit at subsite +1 in a productive orientation that is different from the donor substrate model. Residues possibly involved in hydrogen bond interactions at subsites +1 and +2 include N1019, E1053 (the catalytic acid/base), H1056, N1126 and Y1079 while residues Y1055 and L971 may have hydrophobic interactions. The glucosyl moiety at subsite +3 would be only loosely bound.

Reaction specificity

To test for endolytic activity, GtfB-ΔN was incubated with a blocked 4-nitrophenyl-α-maltoheptaoside substrate (BPNPG7) in the presence of a maltase. This resulted in the release of p-nitrophenol as indicated by an increase in 405 nm absorption. In contrast, incubation with α-glucosidase alone did not result in such an increase (Fig. S3).

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(A) (B)

(C)

Fig. 3. (A) Model for donor substrate binding across the cleavage site: maltoheptaose (yellow carbon atoms) bound in subsites -5 to +2, based on the maltopentaose-bound crystal structure. Putative hydrogen bond interactions with the sugar units in subsites -1, +1 and +2 are indicated as red dashed lines; residue Y1055 may provide a hydrophobic stacking interactions at subsite +2. At the reducing end (indicated by an asterisk), there is space for longer donor substrates. (B) View (from a different angle) of the model for acceptor substrate binding; panose (yellow carbon atoms) is bound in subsites +1 to +3 of the covalent glucosyl-enzyme intermediate model of GtfB-ΔNΔV (cyan carbon atoms). In this orientation, the O6 atom of the non-reducing end glucosyl unit of panose can attack the C1 atom of the covalent intermediate (indicated by the arrow) to form an α1→6 glycosidic linkage. (C) Proposed reaction pathways of 4,6-α-GTases. R = reducing end, NR = non-reducing end; four of the six donor subsites are indicated, of which subsites -2 and -3 are in the tunnel. Newly formed bonds are indicated in red. In (a), donor binding involves a single subsite, or multiple ones (grey units); after the first transglycosylation step, the product can either diffuse away to give products of type 1 or 2, or shift into the tunnel to undergo a next reaction cycle (b), eventually giving products of type 3 or 4. Products of type 2 and 4 can become acceptors for subsequent reaction cycles. Eventually, most of the products will have isomaltooligosaccharide segments at their non-reducing end are obtained (5).

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Genomic analysis

Fig. S8 shows the genomic location mapping of fully or partially sequenced 4,6-α-glucanotransferases in their host bacteria. Using the L. reuteri 121 GtfB sequence as query to search the NCBI database, 28 putative 4,6-α-GTases (identity >50%) were identified almost exclusively in Lactobacillus strains.

In several Lactobacillus strains, the 4,6-α-GTase gene coexists with a GS gene in the same genome. Moreover, downstream or upstream of the 4,6-α-GTase or GS gene, transposase encoding genes were observed. Especially for 4,6-α-GTase genes, almost all are accompanied by a transposase encoding gene on either or both sides.

Phylogenetic analysis

A phylogenetic tree using the full-length sequences of characterized GH70 enzymes (Fig. S9), including 4,6-α-GTases as well as GS, reveals that GS from Lactobacilli are homologous to those from Streptococcus, Leuconostoc and Weisella species. Notably, the characterized 4,6-α-GTases from L. reuteri strains are more homologous to their GS counterparts (in the same organism) than to the GS from other species. This is supported by the results from a Protein-BLAST search of the NCBI database using the L. reuteri 121 GtfB sequence; e.g. the highest homology scores are obtained for the glucansucrase GtfA from the same strain rather than for GS from other strains (data not shown; similar results were obtained in a vice versa analysis using the L. reuteri 121 GtfA sequence). The phylogenetic tree also suggests that L. reuteri 4,6-α-GTase and GS genes originated from a common ancestor gene.

A phylogenetic analysis using the catalytic cores (domains A and B) of bacterial GH13 enzymes, shows the clustering of homologues sequences into different subfamilies of GH13, in accordance with their different reaction specificities (Fig. S10). Including the rearranged catalytic cores (B-A-(C)-A-B→A-B-A) of GH70 GS and 4,6-α-GTase enzymes in this analysis reveals that these two functionalities have a common ancestor, which is closely related to maltooligosaccharide-processing α-amylases from GH13 subfamily 5, but phylogenetically much more distant from members of GH13 subfamilies 4 and 18 active on sucrose.

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DISCUSSION

Unique structural features of GtfB-ΔNΔV

The crystal structure of GtfB-ΔNΔV is the first representative of the 4,6-α-glucanotransferase (4,6-α-GTase) subfamily within GH70. As expected from sequence alignments, the overall domain order and arrangement of GtfB-ΔNΔV resembles that of the GH70 glucansucrases (GS), with a circular permutation of the (βα)8 barrel in the catalytic domain, and a similar spatial arrangement of the catalytic triad (residues D1015, E1053 and D1125). Surprisingly however, while in GS sugar binding subsites beyond -1 are blocked by the loop connecting β7 and α8 (12), in GtfB-ΔNΔV this loop is shorter, uncovering multiple donor subsites much like in GH13 α-amylases (37), while subsites -2 and -3 are occluded from the solvent by two long loops from domain A and B. Together the three loops, which seem to be specific for 4,6-α-GTase, create a tunnel-like structure capable of holding maltooligosaccharides, as exemplified by the structures of the complexes with maltopentaose and with 64-α-D-glucosyl-maltotetraose. Thus, 4,6-α-GTases structurally have characteristics both from GH70 glucansucrases as well as from GH13 α-amylases.

Mechanism and mode of action of 4,6-α-GTases

Knowledge of the GtfB-ΔNΔV crystal structure and its complexes combined with the results of blocked substrate and acceptor reactions prompt us to propose an extended reaction scheme with a second mode of action (Fig. 3C). Notably, while the reaction scheme proposed by Leemhuis et al. (14) described an exo-α1→4 glycosidase activity and a concomitant transfer of single glucosyl units, we propose an additional endo-α1→4 glycosidase activity with transfer of multiple glucosyl units, supported by the finding of the ‘donor tunnel’ with subsites and by the observed release of 4-nitrophenol from BPNPG7. The extended scheme better explains the observed substrate and product specificity of GtfB and other 4,6-α-GTases (17).

The first step in the 4,6-α-GTase reaction is donor substrate binding across the cleavage site; modeling a maltooligosaccharide in GtfB by superposition with an α-amylase showed that the binding modes likely are very similar (Fig. 3A) such that maltooligosaccharides can enter the tunnel and occupy multiple donor

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subsites, thus contributing to the shared substrate specificity between 4,6-α-GTases and α-amylases. On the other hand, differences in some of the residues surrounding subsite +1 between 4,6-α-GTases and GS (Fig. S1A) may explain why 4,6-α-GTases do not use sucrose as donor substrate like glucansucrases. For example, in the Gtf180-ΔN - sucrose complex, a conserved tryptophan (W1065) and a conserved glutamine (Q1140) residue provide direct hydrogen bonds to the fructosyl moiety of the substrate in subsite +1. Instead, at the equivalent positions, GtfB-ΔNΔV and other 4,6-α-GTases have a tyrosine (Y1055), unable to provide an equivalent hydrogen bond, and a lysine (K1128), exchanging a positively charged side chain by a neutral one. Moreover, differences in residues of loops from domain B that line the active site result in a somewhat wider pocket in GtfB-ΔNΔV. Together, although there is space for sucrose to bind (not shown), the shape and properties of subsite +1 likely result in a too low affinity for sucrose to be a substrate. Instead, maltooligosaccharides are the preferred donor substrates, and the fact that our soaking experiments did not reveal occupied acceptor subsites but only occupied donor subsites suggests that the latter have a higher affinity. Combined with the observation of endoglycolytic activity of GtfB-ΔN with BPNPG7 (Fig. S3), we conclude that donor substrate binding can involve multiple donor subsites (Fig. 3C, grey squares). After donor substrate binding, cleavage of the α1→4 glycosidic bond and formation of the covalent intermediate occurs in the same way as in α-amylases, yielding covalent glucosyl- but also oligoglucosyl-enzyme intermediates. Thus, endolytic cleavage is a significant activity of GtfB. Regarding the transglycosylation step, linkage type specificity must originate from residues forming acceptor subsites. For example, in the glucansucrase Gtf180-ΔN - maltose complex (12), resembling α1→6 specificity, residues providing hydrogen bonds to the acceptor are mainly from homology regions II, III and IV (domain A); the importance of these residues in linkage specificity has been shown in several mutation studies (38-40). To investigate acceptor binding in GtfB-ΔNΔV (Fig. 3B), we used panose because it is a better acceptor than maltotriose or isomaltotriose. The modeling results show that this trisaccharide can indeed bind in a productive orientation in subsites +1 to +3 for the formation of an α1→6 glycosidic linkage. Residues E1053 (homology region III, the catalytic base) and N1019 (homology region II, known to affect transglycosylation) seem critical for the orientation of the glucosyl unit in subsite +1, while other residues (N1129 from homology region IV, and Y1079) may help

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orient the acceptor in subsites +2 and +3. Residue Y1055 (conserved in 4,6-α-GTases) may provide a non-specific hydrophobic stacking platform for acceptors at subsite +2, similar to the corresponding tryptophan in glucansucrases; the loss of activity (especially transglycosylation) in mutant Y1055G supports this hypothesis. The partially occluded active site of 4,6-α-GTases explains why these enzymes are less hydrolytic than α-amylases with a fully accessible binding groove. Finally, the presence of the tunnel-forming loops A1 and B near the cleavage/transglycosylation site prevents the 4,6-α-GTases to cleave or form branched oligosaccharide products, as opposed to glucansucrases.

After transglycosylation, in the initially proposed mechanism, products having a (single) α1→6 linked glucosyl unit bond at the non-reducing end would diffuse out of the active site via the acceptor side (Fig. 4, panel a). Based on the obtained crystal structures and observed reaction specificity of GtfB-ΔN(ΔV), we propose that such products alternatively can shift into the tunnel by one or more positions, and become donor substrates for the next reaction cycle (Fig. 4, panel B). Evidence for this comes from the pentasaccharide bound in subsites -2 to -6 observed in the G6 soaking experiment with active GtfB-ΔNΔV, representing a transglycosylation product with an α1→6 linked glucosyl unit at the non-reducing end, shifted by 5 positions, showing that movement of transglycosylation products in the donor direction is possible. Accordingly, mixed isomalto-/maltooligosaccharides, also representing transglycosylation products, are processed by GtfB-ΔN (Fig. S4); this suggests they occupy multiple donor subsites and can be cleaved in an endolytic fashion. The observation of bound saccharides at remote site I (W790/W811) in almost all our soaking experiments with other malto- and isomaltooligosaccharides (data not shown) leads us to speculate that hydrophobic stacking interactions at about 16 Å distance from donor subsite -6 provide extra affinity in the extension of the donor groove/tunnel. At subsite -6, residue Y1521 plays a similar role. The fact that mutants W790A/W811A and Y1521 have only a partly reduced enzyme activity (Table S2) suggests that affinity at remote donor subsites is functional but not essential (for transglycosylation). The importance of the tunnel for transglycosylation is further supported by the observation that mutant T920W has a heavily impaired transglycosylation activity, likely because the large aromatic side chain blocks its access. Finally, the tunnel feature explains why GtfB-ΔN selectively modifies

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unbranched segments of starch while leaving the core structure intact. This specificity provides interesting opportunities for the industrial application of 4,6-α-GTase enzymes in starch-processing.

Genomic and phylogenetic analysis of 4,6-α-GTase genes

The occurrence of homologous genes in a species is believed to be the result of either horizontal gene transfer (orthologs) or gene duplication (paralogs). During evolution, such events are proposed to be driven by changes in habitat such as the availability of fermentable enzyme substrates (41). Upon orthologous or paralogous acquisition, the genes may be partly modified to adapt to the new environment, and afterwards may further evolve independently (42, 43). Earlier studies showed evidence that both horizontal gene transfer and gene duplication events have occurred for GH70 glucansucrases (GS) of oral Streptococcus and Leuconostoc species (29, 44, 45). The results of our genomic and phylogenetic analysis suggest that the same phenomena are involved in the evolution of 4,6-α-GTase genes in Lactobacilli. First, in most of the Lactobacillus species, the 4,6-α-GTase gene is flanked by a transposase gene on either or both sides (Fig. S8), as was observed earlier for several of their GS genes (8). Transposases are involved in horizontal gene transfer, and have been associated with intra- and interspecies genomic variation of oral bacteria such as Streptococci (46, 47). Interestingly, Hoshino et al. (41) stated that GS genes from Streptococci were acquired from Lactobacilli via transposase-mediated gene transfer, when these species encountered each other in fermented food. Thus, the presence of 4,6-α-GTase-adjacent transposase genes suggests that also 4,6-α-GTase genes may have undergone horizontal gene transfer during their evolution from one species to another. Second, we observed tandem-arranged GH70 genes encoding a 4,6-α-GTase and a GS in several of the Lactobacillus strains (Fig. S8); their high sequence and length similarity clearly suggest they are the result of gene duplications. This is further supported by the fact that nearby the 4,6-α-GTase / GS gene pairs, transposases genes are found which mediate gene duplication. Indeed, tandem GH70 genes are found in many bacteria, suggesting they readily duplicate. Together, we conclude that the evolution of 4,6-α-GTase and GS genes in lactic acid bacteria involved both horizontal gene transfer and gene duplication, leading to two closely related subfamilies with different reaction specificities.

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The high sequence similarity between 4,6-α-GTase and GS gene pairs within a genome also is reflected in the results of a phylogenetic analysis of GH70 enzymes in lactic acid bacteria (Fig. S9). Earlier phylogenetic studies only included the conserved catalytic regions (13, 41) and suggested that the 4,6-α-GTase cluster constitutes a separate subfamily independent of glucansucrases. In contrast, our analysis of the full-length genes shows that 4,6-α-GTase genes are closely related to their GS tandem counterpart, more than to GS genes from other species. Moreover, in accordance with the evidence for gene duplication events described above, the GH70 4,6-α-GTase and GS genes clearly share a common ancestor. In a second phylogenetic analysis using rearranged catalytic cores of GH13 and GH70 enzymes (Fig. S10), we found that this common ancestor is closely related to α-amylases from GH13 subfamily 5 (GH13_5) rather than to sucrose-acting enzymes from subfamilies 4 and 18. Interestingly, enzymes from this subfamily are mainly found in Streptococcus, Bacillus and Exigobacterium species, correlating with the occurrence of GH70 enzymes (see CAZy database) (10). Enzymes from GH13_5 process maltooligosaccharides; combining this observation with the present-day substrate specificities of GH70 4,6-α-GTases and GS (maltooligosaccharides and sucrose, respectively), we propose that the common ancestor of GH70 4,6-α-GTases and GS processed maltooligosaccharides. The ability to utilize sucrose would then have been acquired later (in glucansucrases). Our conclusions from the genomic and phylogenetic analyses are supported by the crystal structure of GtfB-ΔNΔV. The domain organization of GtfB-ΔNΔV is shared with glucansucrase GtfA from the same organism, supporting that tandem 4,6-α-GTase and GS pairs within a single genome arose from gene duplication, while the active site architecture of GtfB-ΔNΔV resembles more the GH13 α-amylases. Thus, the 4,6-α-GTase subfamily can be regarded as an evolutionary intermediate between GH13 maltooligosaccharide-processing α-amylases and GH70 glucansucrases.

Functional role of 4,6-α-GTases

From the above described bioinformatic analyses we speculated how GH70 genes may have evolved. To further underpin this, we investigated EPS production in vivo in a Lactobacillus strain constitutively expressing both a 4,6-α-glucanotransferase and a glucansucrase. Provided with maltodextrins or sucrose as carbon source, L. reuteri 35-5 utilizes either its 4,6-α-glucanotransferase GtfB

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or its glucansucrase GtfA enzyme for the synthesis of α-glucans with (α1→4 and α1→6) linkages (chapter 5). The extracellular polymers, present in the slimy halos around the colonies, also possess α1→4 and α1→6 linkages and have a similar size distribution. In short, both L. reuteri enzymes synthesize a related type of EPS, from different substrates.

Although the in vivo functional roles of glucansucrases in lactic acid bacteria remain to be understood, many glucansucrases have been proven to contribute to biofilm formation in vivo by producing α-glucans. Importantly, for Streptococcus mutans, which is considered the principle pathogen of dental caries, it was shown that the α-glucan produced by its glucansucrase facilitates the adherence of the bacterium to the tooth surface (4). In Lactobacillus reuteri, gene knockout results demonstrated that, without glucansucrase, biofilm formation is less effective (5). Notably, these studies were based on the supply of sucrose. Recently, we also show that in Lactobacillus reuteri 121 (strain 35-5) the uptake of starch, the most abundant dietary carbohydrate, also results in α-glucan production, and that the responsible enzyme is its 4,6-α-glucanotransferase GtfB (Bai et al., submitted). The α-glucans produced by L. reuteri on agar plates with sucrose or starch are structurally similar to that produced by its glucansucrase (GtfA). Therefore, also 4,6-α-GTases have great potential to contribute to biofilm formation by utilizing starch instead of sucrose. This finding is also in accordance with the above proposed ideas about gene evolution; both the 4,6-α-GTase and GS produce related α-glucans, but from different substrates, due to adaptive evolution driven by substrate variation.

Evolutionary relationships between α-amylases, 4,6-α-glucanotransferases

and glucansucrases

Based on the structural observations in GtfB-ΔNΔV, and the genomic and phylogenetic analyses of 4,6-α-glucanotransferases we propose the following hypothesis for the evolutionary relation of this GH70 subfamily to GH13 α-amylases and GH70 glucansucrases. α-Amylases from GH13 subfamily 5 mainly degrade starch and starch-derived malto-oligosaccharides. They possess a long and fully solvent-accessible substrate binding groove with multiple donor subsites, due to the relatively short loops adjacent to this groove. As a consequence, these enzymes are mainly hydrolytic and produce glucose and/or short oligosaccharides.

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Evolution of GH13_5 enzymes, involving domain insertion and circular permutation, leads to a common ancestor of family GH70 that acquired transglycosylation activity with maltooligosaccharides to produce polysaccharides instead. In order to achieve such different reaction specificity (from hydrolysis to polymerization), some of the loops (A1 and B) surrounding the donor half of the binding groove elongated, and partially shielded donor subsites from the solvent. This prevented diffusion of oligosaccharide reaction intermediates out of the binding groove, while keeping them available for the subsequent acceptor reaction (transglycosylation) and favoring a more processive mode of action. Whether they still had the 4-transglygosylation specificity observed in α-amylases, or already possessed the 6-transglycosylation specificity observed in 4,6-α-glucanotransferases is unknown. Evolution towards the glucansucrase subfamily, possessing a different substrate specificity (sucrose instead of starch), resulted from the elongation of another loop (A2) that blocked donor subsites beyond -1. Concurrently, loops A1 and B were no longer needed to shield the donor half of the binding groove, and could become shorter. Also, slight changes around subsite +1 likely were needed to prefer interaction with the fructosyl moiety in the donor reaction.

Fig. 4. Evolutionary relationships between α-amylases, 4,6-α-glucanotransferases and glucansucrases and the role of loops A1, A2 and B. The triangle indicates the catalytic site for cleavage and transglycosylation. The α-amylases that process maltooligosaccharides (white circles represent glucose units) have an open binding groove with multiple donor (-1 - -4) and acceptor (+1 - +3) subsites, lacking loops A1 and B, and are unable to synthesize polymers. The 4,6-α-glucanotransferases acquired this capability via elongation of loops B and A1 forming a tunnel at the donor half of the groove, allowing them to better capture the intermediate for subsequent transglycosylation and thus shift their product specificity towards synthesis of α-glucan polymers. Subsequently, a shift in substrate specificity was obtained by elongation of loop A2 which blocks donor subsites beyond -1; loops A1 and B became shorter; in this way, glucansucrases acquired the capability to use sucrose (grey circle represents fructose unit) for the synthesis of α-glucan polymers.

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Conclusions The crystal structure of GtfB-ΔNΔV is the first representative of the GH70 4,6-α-glucanotransferase subfamily, with an overall domain organization resembling GH70 glucansucrases, and an active site architecture reminiscent of GH13 α-amylases. A tunnel-like binding groove allows for a dual mode of action for 4,6-α-GTases involving both exo-specific and endo-specific cleavage/transfer reactions, and explains why GtfB only processes the outer (amylose) branches of starch substrates. The GtfB-ΔNΔV structure provides a starting point for (rational) mutation of 4,6-α-GTases to alter enzyme specificity, e.g. in starch-processing applications aiming at the production of prebiotic dietary fibers from a cheap substrate, using a GRAS bacterial strain. The GtfB-ΔNΔV crystal structure clearly supports our previous proposal that the 4,6-α-GTase subfamily is an evolutionary intermediate between GH70 and GH13 enzymes. The structural observations support our phylogenetic and genomic analyses, revealing the close relation between 4,6-α-GTases and GS, and suggesting that gene duplication as well as horizontal gene transfer events occurred during evolution. Loop mutations near the active site of a common ancestor likely lead to either an α1→6 specific transglycosylation activity (in 4,6-α-GTase) or to a different substrate specificity (in GS). We propose that the evolution of bacterial species such as Lactobacillus, using 4,6-α-glucanotransferases and glucansucrases to produce α-glucan exopolysaccharides with α1→4 and α1→6 linkages, was driven by changes in the availability of fermentable sugars in the human diet.

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SI Materials and Methods

Construction of truncated GtfB-ΔNΔV

Truncation of GtfB lacking both the N-terminal variable domain and domain V was based on the alignment of the Lactobacillus reuteri 121 GtfB with glucansucrase sequences and on the crystal structures of Gtf180-ΔN glucansucrase (PDB entry 3KLK) from Lactobacillus reuteri 180 and DSR-E-ΔN (PDB entry 3TTO) from Leuconostoc mesenteroides NRRL B-1299. The gtfB gene fragment encoding GtfB (Uniprot entry Q5SBM0) amino acids 761–1619 was amplified by PCR using High Fidelity PCR enzyme mix (Thermo-Scientific, Landsmeer, The Netherlands) with pET15b-gtfB as template and the primers CHisFor-dNdVgtfB 5′- GATGCATCCATGGGACCAGGTACTTGGGAAAATATGGCTTTCGCTCAA-3′ and CHisRev-dNdVgtfB 5′- CCTCCTTTCTAGATCTATTAGTGATGGTGATGGTGATGGTTGTTAAAGTTTAATGAAATTGCAGTTGG-3′. A nucleotide sequence encoding a 6×His-tag was fused in-frame to the 3′ end of the gtfB-ΔN gene, using the reverse primer. The resulting PCR product was digested with NcoI and BglII, and was ligated into the corresponding site of pET15b. The constructed plasmid was transformed into Escherichia coli DH5α. The construct extracted from E. coli DH5α was confirmed by nucleotide sequencing (GATC Biotech, Glasgow, UK) followed by transforming into Escherichia coli BL21 star (DE3).

Site-directed mutagenesis of GtfB-∆N∆V

The plasmid constructed above was used as the template for site-directed mutagenesis mutagenesis. Mutations were introduced by PCR with primers (see Table S1) using the Quik-Change site-directed mutagenesis kit (Stratagene. La Jolla, CA). The primers are purchased from Sigma-Aldrich. The PCR product was cleaned up with the PCR cleaning up kit (Sigma-Aldrich) followed by transformation into Escherichia coli BL21 star (DE3). Mutation of the plasmid was confirmed by gene sequencing in GATC Biotech (Glasgow, UK).

Substrate endo specificity

GtfB-∆N∆V (1 μM) was incubated with Blocked 4-Nitrophenyl-α-Maltoheptaoside (BPNPG7, 10 mM) and α-glucosidase (yeast maltase,

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Megazyme, 20 U/ml), at pH 6.0 (25 mM sodium citrate buffer) and 40 °C for 5 h; release of the 4-nitrophenyl group was determined by following the absorbance change at 405 nm.

GtfB∆N (1 μM) was incubated with 63-α-D-Glucosyl-maltotriosyl-maltriose (10 mM) in sodium acetate buffer (25 mM, pH 5.0, 1 mM CaCl2) at 40 °C for 24 h. Reaction products were analyzed by HPAEC (see below). Maltooligosaccharides (G1-G8) and 63-α-D-glucosyl-maltriose were applied as references.

Acceptor specificity

GtfB-∆N∆V (60 nM) was incubated with amylose (0.25%, w/v) in sodium acetate buffer (25 mM, pH 5.0, 1 mM CaCl2) at 40 °C, in the presence or absence of acceptor substrates maltotriose, isomaltotriose and panose (10 mM). Reaction rates were determined using an amylose-iodine assay; transglycosylation factors were calculated as described by Bai et al. (2015).

Characterization of mutants

Enzymatic characterization of the mutants and the analysis of their products are shown in Table S2 and Fig. S6. Mutation of the catalytic nucleophile D1015 and of the general acid/base E1053 resulted in complete inactivation of the enzyme; activity was below the detection level. Enzyme activities of the single mutants T920W and Y1521A (tunnel), K1128A/N/H/W (subsite -1/+1), Y1055G (subsite +2) and of the double mutant W790A/W811A were determined using amylose V as a substrate; products were analyzed by TLC, 1H NMR and HPAEC.

Compared to wild-type GtfB-ΔNΔV (Fig. S6A), mutation of the aromatic residues at positions 790, 811 and 1521 to an alanine resulted in a 30-40% loss of total activity; however linkage specificity and product spectra were virtually unaffected.

Mutant T920W retained only 17% of total activity. The almost complete absence of α1→6 linkages in products indicates that this mutant has impaired transglycosylation activity and is virtually hydrolytic, accumulating glucose as the main product over time (Fig. S6B).

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Removing the aromatic side chain in the Y1055G mutant results in a 60% loss of total activity and a 10-fold decrease in the relative amount of α1→6 linkages in its products. Compared to wild-type GtfB-ΔNΔV, this mutant released less free glucose over time, but hydrolyzed amylose to form maltooligosaccharides (Fig. S6C).

Mutation of K1128 to A, N, H or W resulted in elevated relative amounts of α1→4 linkages in the products and free glucose; accompanied by a slight decrease (K1128H) or marked increase (K1128A/N/W) of the total activity. All mutants produced similar products, acting endolytically and mainly converting amylose to maltooligosaccharides without forming transglycosylation products. The K1128W mutant produced lower amounts of glucose, and released maltooligosaccharides of different DP (Fig. S6D).

Modeling

Donor substrate modeling. The structure of the GtfB-ΔNΔV with maltopentaose (G5) bound in subsites -1 to -5 was superposed on that of Bacillus subtilis α-amylase complexed with G5 in subsites -3 to +2 (PDB 1BAG) (35), using their catalytic triads to improve the superposition; as a result, the glucosyl moiety at subsite -1 superimposed almost perfectly. The G5 in GtfB-ΔNΔV was then extended in acceptor subsites +1 and +2 by connecting two glucosyl moieties, and adjusting the glycosidic torsion angles such as to avoid clashes with protein residues. The result is shown in Fig. 3A.

Acceptor substrate modeling. The apo GtfB-ΔNΔV crystal structure was superposed on that of the Bacillus circulans strain 251 CGTase covalent maltotriosyl-enzyme intermediate (PDB: 1CXL) (36); a covalently linked glucosyl moiety was then ‘attached’ to the OD2 atom of the catalytic nucleophile D1015. A slight rotation of the OD2-C1 bond was applied to optimize hydrogen bond possibilities of the glucosyl unit with surrounding protein residues. Subsequentely, the trisaccharide panose was modeled in acceptor subsites +1 to +3 guided by the following requirements. First, the O6 of the non-reducing end glucosyl unit should point towards the anomeric C1 atom of the covalent glucosyl-enzyme intermediate, at a reasonable distance for attack. Second, a hydrogen bond interaction to the same O6 should be provided by the catalytic base E1053; this interaction is required for the abstraction of the O6 proton in

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order to activate the acceptor for the transglycosylation to take place. Furthermore, hydrogen bond interactions likely are provided to the non-reducing end glucosyl unit by residue N1019, as mutation of this residue is known to decrease the relative α1→6 transglycosylation activity (data not shown). The other two glucosyl units of panose were then placed such that, by adjusting glycosidic torsion angles, clashes with protein residues are avoided and enough space for longer acceptor substrates is available (at the reducing end). The result is shown in Fig. 3B.

In vivo growth conditions

Lactobacillus reuteri 35-5 was subcultured from stocks stored at -80 °C in 10 ml of sugar-free MRS medium (liquid culture or agar plate) supplemented with 1% (w/v) glucose. The fresh cultures were inoculated anaerobically in modified sugar-free MRS-agar plate with sucrose (5%, w/v), or maltodextrins (dextrose equivalent (DE) = 13-17, Sigma-Aldrich, US, 5%, w/v). One liter MRS-agar medium contained 10 g bactopeptone, 4 g yeast extract, 5 g sodium acetate, 2 g tri-ammonium citrate, 0.2 g MgSO4·7H2O, 0.05 g MnSO4·7H2O; and 1 ml Tween 80 and 15 g/l agar.

Growth of L. reuteri 35-5 on either sucrose or maltodextrins results in the formation of transparent colonies surrounded by a slimy halo (Fig. S7A). Previously, it has been shown that the EPS formed by L. reuteri 35-5 from sucrose is a reuteran containing both α1→6 and α1→4 linkages (49). Monosaccharide analysis (data not shown) and NMR analysis (Fig. S7B) showed that the EPS derived from the maltodextrin-grown cultures also has with α1→6 and α1→4 linkages (61 versus 39%, respectively).

In-gel PAS staining of enzymes (Fig. S7D) (Bai et al., submitted) revealed that, when incubated with sucrose substrate, the constitutively expressed glucansucrase GtfA (199 kDa) is associated with reuteran synthesis, in agreement with earlier studies (48). The minor band at ~120 kDa may represent GtfA-ΔN, which is known to synthesize a reuteran very similar in size distribution and linkage type to that produced by the full-length enzyme (49). In case of incubation of the gel with maltodextrins, the band corresponding to GtfA is absent in the PAS stained gel, while a band corresponding to GtfB (179 kDa) is stained, indicating that not the glucansucrase GtfA, but the 4,6-α-glucanotransferase GtfB is associated with

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α-glucan synthesis when L. reuteri 35-5 is supplied with maltodextrins. SEC analysis of the EPS isolated from the plated cultures (Fig. S7C) reveals that the synthesized reuterans are approximately of the same size (35-40 MDa). When incubated with a mixture of sucrose and maltodextrins, both bands corresponding to GtfA and GtfB are associated with α-glucansynthesis. Notably, genomic data suggest that the only carbohydrate acting enzymes larger than 150 kDa are GtfA and GtfB (Gangoiti et al., in preparation).

Extraction of extracellular polysaccharides (EPS)

The extracellular polysaccharides (EPS) were washed out by 10 ml water from MRS-agar plates that had been incubated for 72 h at 37 °C with different carbon sources. Two volumes of cold ethanol (-20 °C) were added to the dissolved samples and kept at 4 °C overnight. The precipitate was harvested by centrifugation (12 000 ×g, 30 min, 4 °C) and re-dissolved in 1 volume of dd H2O. The EPS were re-precipitated with 2 volumes of ethanol at 4 °C overnight. After centrifugation, the harvested precipitate was dissolved in water, and then dialyzed (10 kDa MWCO, Thermo Scientific) against water for 48 h with changes of dd H2O in each 12 h (50). The EPS from followed by ethanol precipitation as described above, and analyzed by 1D 1H NMR (see below).

Monosaccharide analysis

EPS samples (0.5 mg) were dissolved in 200 µl Milli-Q water in a glass reaction tube. An equal volume of 4 M trifluoroacetic acid (TFA) was added, followed by hydrolysis for 4 h at 100 °C. Samples were dried under a flow of dry nitrogen and subsequently dissolved in 100 µl isopropanol and dried again by evaporation under dry nitrogen. Samples were dissolved in 1 ml dimethyl sulfoxide (DMSO) and used for HPAEC analysis (see below).

Size exclusion chromatography (SEC)

DMSO-LiBr (0.05M) was prepared by stirring for 3 h at room temperature followed by degassing for 15 min using an ultrasonic cleaner (Branson 1510, Branson, Danbury, CT). Samples were dissolved at a concentration of 4 mg/ml in DMSO-LiBr by overnight rotation at room temperature, followed by 30 min heating in an oven at 80 °C, obtaining clear sample solutions. The samples were cooled to room temperature and filtered through a 0.45-µm Millex PTFE

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membrane (Millipore Corporation, Billerica, MA). The SEC system set-up (Agilent Technologies 1260 Infinity) from PSS (Mainz, Germany) consisted of an isocratic pump, auto sampler, an online degasser, an inline 0.2 μm filter, a refractive index detector (G1362A 1260 RID Agilent Technologies), viscometer (ETA-2010 PSS, Mainz) and MALLS (SLD 7000 PSS, Mainz). WinGPC Unity software (PSS, Mainz) was used for data processing. Samples (100 µl) were injected into a PFG guard column using an autosampler at a flow rate of 0.5 ml/min and DMSO-LiBr as eluent. The separation was done by three PFG-SEC columns with porosities of 100, 300 and 4000 Å. The columns were held at 80˚C, the refractive index detector at 45 °C and the viscometer was thermostatted at 60 °C. A standard pullulan kit (PSS, Mainz, Germany) with molecular masses from 342 to 805000 Da was used. The specific RI increment value dn/dc was measured by PSS and is 0.072 (obtained from PSS company).

Nuclear magnetic resonance (NMR) spectroscopy

EPS samples isolated from L. reuteri 35-5 agar plate were analyzed by NMR. Resolution-enhanced 1D 500-MHz 1H NMR spectra were recorded in D2O (99.9 atm% D, Cambridge Isotope Laboratories, Inc.) on a Varian Inova Spectrometer (NMR Center, University of Groningen) at probe temperatures of 300 K. Spectra were recorded with 16k complex points, using a 5000 Hz spectral width. Data are expressed relative to internal acetone (δ1H 2.225) Prior to analysis, samples were exchanged twice with D2O with intermediate lyophilization and then dissolved in 0.6 ml D2O. Spectra were processed using MestReNova 5.3 software (Mestrelabs Research SL, Santiago de Compostella, Spain), using Whittaker Smoother baseline correction and zero filling to 32k complex points.

High-performance anion-exchange chromatography (HPAEC)

Samples were injected onto a 4×250 nm CarboPac PA-1 column connected to a Dionex DX500 workstation (Dionex). Samples were run with a gradient of 30-600 mM NaAc in 100 mM NaOH (1 ml/min), and detected by an ED40 pulsed amperometric detector. A mixture with known concentrations of glucose, isomaltose, isomaltotriose, maltose, panose, maltotriose, maltotetraose, maltopentaose, maltohexaose and maltoheptaose was used as reference.

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Thin-layer chromatography (TLC)

The TLC silica gel 60F254 plates (Merck) were run with butanol/acetic acid/water (2:1:1, v/v/v) as solvent. After running for 6 h, TLC plates were developed with 10% (v/v) H2SO4 and 2 g/l orcinol in methanol and then heated by oven at 100 °C for 30 min. Glucose, maltose, maltotriose, maltotetraose, maltopentaose, maltohexaose, and maltoheptaose were used as markers.

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Table S1. Oligonucleotides used for site-directed mutagenesis of gtfB-ΔNΔV

Mutation in GtfB-ΔNΔV

protein Sequence (5´→3´)*

W790A GGTTACTTAAGTTATACTGATGCGTATCGTCCTTATGGCACAAG CTTGTGCCATAAGGACGATACGCATCAGTATAACTTAAGTAACC

W811 GGTACAAAACAACTGCAATGGATGCGCGTCCATTACTGATGTATATTTGCAAATATACATCAGTAATGGACGCGCATCCATTGCAGTTGTTTTGTACC

T920W CGACCAGATAAAAGTGGATGGATTGATAGTGATCAAGTC GACTTGATCACTATCAATCCATCCACTTTTATCTGGTCG

D1015N GGTTTCCGAGTTAATGCTGCTGATA TATCAGCAGCATTAACTCGGAAACC

Y1055G GTTATAATGAGGGCGGTCATTCTGGTGCTGCTC GAGCAGCACCAGAATGACCGCCCTCATTATAAC

K1128A CTAACCATGATCAACGAGCGAATTTGATTAATAG CTATTAATCAAATTCGCTCGTTGATCATGGTTAG

K1128N CTAACCATGATCAACGAAACAATTTGATTAATAG CTATTAATCAAATTGTTTCGTTGATCATGGTTAG

K1128H CTAACCATGATCAACGACACAATTTGATTAATAG CTATTAATCAAATTGTGTCGTTGATCATGGTTAG

K1128W CTAACCATGATCAACGATGGAATTTGATTAATAG CTATTAATCAAATTCCATCGTTGATCATGGTTAG

Y1521A GCAAATCAAATTTATTTTGCAGCTACAACTGGTGGCGGAAATGG CCATTTCCGCCACCAGTTGTAGCTGCAAAATAAATTTGATTTGC

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Table S2. Effects of mutations on the reaction and product specificity of L. reuteri 121 GtfB-ΔNΔV.

a Total and hydrolytic activities were determined using 0.125% w/vol amylose V as substrate. Assays were performed at 37 ᵒC in 25 mM sodium acetate buffer, pH 4.7 containing 1 mM CaCl2. b Total activities were measured by the iodine-staining assay. c Hydrolytic activities were determined by following the release of glucose in time using the GOPOD kit (15). d The percentages of glucose, α1→4 and α1→6 glycosidic linkages were determined by 1H NMR analysis of the product mixtures synthesized by incubations of amylose V (0.6% w/v) with the different enzymes (95 mU ml-1).

Enzyme Enzyme activity (U mg-1) a Product mixture linkages (%) d

total b hydrolytic c α1→6 α1→4 free glucose units

wild-type 2.9 ± 0.17 0.37 ± 0.02 82 10 8

W790A/W811A 1.7 ± 0.02 0.27 ± 0.06 80 11 9

Y1521A 2.1 ± 0.17 0.36 ± 0.01 82 9 9

T920W 0.5 ± 0.01 0.27 ± 0.01 4 34 62

Y1055G 1.2 ± 0.15 0.08 ± 0.03 8 75 17

K1128A 5.4 ± 0.47 0.48 ± 0.01 2 87 11

K1128N 6.5 ± 0.09 0.68 ± 0.01 3 86 11

K1128H 2.1 ± 0.11 0.26 ± 0.01 4 79 16

K1128W 3.3 ± 0.08 0.11 ± 0.08 1 94 5

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Fig. S1. (A) Comparison of L. reuteri 121 GtfB-ΔNΔV (this work; molecule A of the asymmetric unit is shown, left) with the L. reuteri 180 Gtf180-ΔN - sucrose complex (right) (12). Blue represents domain A and green represents domain B; the tunnel in GtfB-ΔNΔV is indicated with dashed lines. Catalytic residues are labeled bold. In Gtf180-ΔN, the fructosyl moiety in subsite +1 has hydrogen bond interactions with residues Q1140 and W1065 and, via a water-mediated hydrogen bond network, with residues N1411 and D1458. In GtfB-ΔNΔV, the corresponding residues are K1128, Y1055, L1390, but the fourth residue is absent; moreover, the side chains of Y1055 and L1390 are unable to make hydrogen bonds. Finally, differences in residues of domain B cause the active site pocket to be wider in GtfB-ΔNΔV. (B) GtfB-ΔNΔV D1015N - maltopentaose complex (stereo view) showing direct and water-mediated hydrogen bond interactions of the pentasaccharide with protein residues in donor subsites -1 to -5. Catalytic residues are labeled bold; residues from loops A1, A2 and B are labeled purple, red and brown, respectively. (C) Stereo view of the pentasaccharide observed in the GtfB-ΔNΔV - G6 soak showing hydrogen bond interactions, in donor subsites -2 to -6.

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Fig. S2. Sequence alignment of 4,6-α-glucanotransferase (4,6-α-GTase), glucansucrases (GS) and α-amylases (α-Amy); the latter were aligned separately. Alignments were made using ESPript v3.0 (31). Numbering corresponds to the sequences of LrGtfB and BlAMY. Top: conserved motifs I-IV. The seven conserved α-amylase superfamily residues are indicated by triangles (▲), including the catalytic residues (NU = nucleophile, A/B = general acid/base, TS stab = transition state stabilizing residue). Other residues near subsites -1 and +1 are indicated by stars (*). Bottom: loop regions around the active site: loops A1, A2 and B (grey shading); α-amylases were aligned only for loop A2 since these enzymes do not have an equivalent to loops A1 and B. LrGtfB = Lactobacillus reuteri 121 GtfB, LrGtfML4 = L. reuteri ML1 GtfML4, LrGtfW = L. reuteri DSM 20016 GtfW, LrGtf180 = L. reuteri 180 Gtf180, LrGtfA = L. reuteri 121 GtfA, LrGtfO = L. reuteri ATCC 55730 GtfO, LrGtfML1 = L. reuteri ML1 GtfML1, SmGtf-SI = Streptococcus mutans Gtf-SI, LmDSRE = Leuconostoc mesenteroides NRRL B-1299 DSRE, BlAMY = Bacillus licheniformis α-amylase, BhAMY = Bacillus halmapalus α-amylase, GsAMY = Geobacillus stearothermophilus α-amylase, BsAMY = Bacillus subtilis α-amylase.

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Fig. S3. Release of p-nitrophenol from BPNPG7 (structure shown in inset) incubated with α-glucosidase (red) or α-glucosidase + GtfB-ΔN (blue) as measured by the absorbance change at 405 nm. The exo-acting yeast maltase cannot cleave the substrate; in contrast, with the addition of GtfB-ΔNΔV, p-nitrophenol is released. This indicates that GtfB-ΔN cleaves α1→4 linkages in an endo fashion, releasing the blocked end of the substrate; subsequently the maltase hydrolyzes the intermediate products until it reaches the non-reducing end to release p-nitrophenol.

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Fig. S4. HPAEC analysis of 63-α-D-glucosyl-maltotriosyl-maltriose (10 mM) processed by GtfB-∆N (1 μM) in sodium acetate buffer (25 mM, pH 5.0, 1 mM CaCl2) at 40 °C for 24 h (black line). Glucose (G1), fructose (F), isomaltose (G2), sucrose (S), maltose (G2), panose (P), maltose (G2), maltotriose (G3), maltotetraose (G4), maltopentaose (G5), maltohexaose (G6), maltoheptaose (G7), maltooctaose (G8), and 63-α-D-glucosyl-maltriose were applied as references (colored lines).

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Fig. S5. TLC analysis of the product mixtures obtained from the incubations of amylose V (0.6% w/vol) with 95 mU ml-1 of wild-type GtfB-ΔNΔV, and its mutants: Lane 1, wild-type GtfB-ΔNΔV; lane 2, W790A-W811A; lane 3, Y1521A; lane 4, T920W; lane 5, Y1055G; lane 6, K1128A; lane 7, K1128N; lane 8, K1128H; and lane 9, K1128W. The reaction mixtures were incubated at 37 ᵒC and pH 4.7 during 48 h. Std , standards; G1, glucose; G2, maltose; G3, maltotriose; G4, maltotetraose; G5, maltopentaose; G6, maltohexaose; G7, maltoheptaose; Pol, polymer.

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Fig. S6. HPAEC-PAD profiles of the oligosaccharide mixtures formed upon the incubation of amylose V with (A) GtfB-ΔNΔV, (B) T920W, (C) Y1055G, and (D) K1128W for t = 10 min, 30 min, 24 h (pH 4.7, 37 ᵒC). The identity of peaks was assigned using commercial oligosaccharide standards (G1, glucose; G2-G7, maltose to maltoheptaose; iso-G2, isomaltose; iso-G3, isomaltotriose; Pa, panose).

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Fig. S7. In vivo EPS formation by L. reuteri 35-5. (A) Growth of L. reuteri 35-5 on MRS agar supplemented with different carbon sources, forming slimy or non-slimy colonies. (B) 1D 1H NMR analysis of the EPS extracted from the slimy colonies on agar plates shows that their composition is similar with respect to the glycosidic linkage types present (α1→4 and α1→6). (C) SEC analysis of the EPS extracted from the slimy colonies on agar plates. (D) Periodic Acid-Schiff (PAS) stained SDS-PAGE gels (adapted from Bai et al., submitted) of whole cell proteins of L. reuteri 35-5 grown in the presence of either sucrose, sucrose plus maltodextrins or maltodextrins, revealing the association of the glucansucrase GtfA with sucrose-derived EPS and of the 4,6-α-glucanotransferase GtfB with maltoedextrins-derived EPS.

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Fig. S8. Genomic mapping of 4,6-α-glucanotransferase (4,6-α-GTase) and glucansucrase genes. Included are gtf180 from Lactobacillus reuteri 180, gtfB and gtfA from L. reuteri 121, gtf106B and gtf106A from L. reuteri TMW1.106, gtfML4 and gtfML1 from L. reuteri ML1, 4,6-α-GTase and GS genes from L. reuteri TMW1.656, and 4,6-α-GTase and GS genes from L. salivarius GJ-24, other putative 4,6-α-GTase genes from L. delbrueckii, L. plantarum, L. sanfranciscensis, L. mucosae, and Pediococcus pentosaceus.

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Fig. S9. Unrooted phylogenetic tree of characterized 4,6-α-glucanotransferases (green box) and glucansucrases from Lactobacillus reuteri strains (pink box) and other lactic acid bacteria. Alignments and dendrogram construction were carried out using full length protein sequences (including N-terminus, domain V and IV) with MEGA version 6, using the maximum likelihood method. The bar corresponds to a genetic distance of 0.1 substitution per position (10% amino acid sequence difference).

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Fig. S10. Phylogenetic tree of family GH13 and GH70. The bacterial protein sequences were all characterized based on CAZY database (www.cazy.org) (10). Domains A and B of GH70 members were extracted and rearranged according to the (non-circularly permuted) GH13 domain arrangement. The sequences classified into different subfamilies as represented by different colors.

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Acknowledgements The study was financially supported by the Chinese Scholarship Council (to YB), the University of Groningen and by the TKI Agri&Food program as coordinated by the Carbohydrate Competence Center (CCC-ABC; www.cccresearch.nl).

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4. Koo H, Xiao J, Klein MI, Jeon JG. 2010. Exopolysaccharides produced by Streptococcus mutans glucosyltransferases modulate the establishment of microcolonies within multispecies biofilms. J Bacteriol 192:3024-3032.

5. Walter J, Schwab C, Loach DM, Ganzle MG, Tannock GW. 2008. Glucosyltransferase A (GtfA) and inulosucrase (Inu) of Lactobacillus reuteri TMW1.106 contribute to cell aggregation, in vitro biofilm formation, and colonization of the mouse gastrointestinal tract. Microbiol-SGM 154:72-80. 6. Xiao J, Klein MI, Falsetta ML, Lu BW, Delahunty CM, Yates JR, Heydorn A, Koo H. 2012. The exopolysaccharide matrix modulates the interaction between 3D architecture and virulence of a mixed-species oral biofilm. Plos Pathog 8:4. 7. Carlsson J, Grahnen H, Jonsson G. 1975. Lactobacilli and Streptococci in the mouth of children. Caries Res 9:333-339.

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9. van Hijum SAFT, Kralj S, Ozimek LK, Dijkhuizen L, van Geel-Schutten IGH. 2006. Structure-function relationships of glucansucrase and fructansucrase enzymes from lactic acid bacteria. Microbiol Mol Biol R 70:157-176.

10. Lombard V, Ramulu HG, Drula E, Coutinho PM, Henrissat B. 2014. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res 42:D490-D495.

11. Leemhuis H, Pijning T, Dobruchowska JM, van Leeuwen SS, Kralj S, Dijkstra BW, Dijkhuizen L. 2013. Glucansucrases: Three-dimensional structures, reactions,

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mechanism, α-glucan analysis and their implications in biotechnology and food applications. J Biotechnol 163:250-272.

12. Vujičić-Žagar A, Pijning T, Kralj S, Lopez CA, Eeuwema W, Dijkhuizen L, Dijkstra BW. 2010. Crystal structure of a 117 kDa glucansucrase fragment provides insight into evolution and product specificity of GH70 enzymes. P Natl Acad Sci USA 107:21406-21411.

13. Kralj S, Grijpstra P, van Leeuwen SS, Leemhuis H, Dobruchowska JM, van der Kaaij RM, Malik A, Oetari A, Kamerling JP, Dijkhuizen L. 2011. 4,6-α-Glucanotransferase, a novel enzyme that structurally and functionally provides an evolutionary link between Glycoside Hydrolase enzyme families 13 and 70. Appl Environ Microb 77:8154-8163. 14. Leemhuis H, Dijkman WP, Dobruchowska JM, Pijning T, Grijpstra P, Kralj S, Kamerling JP, Dijkhuizen L. 2013. 4,6-α-Glucanotransferase activity occurs more widespread in Lactobacillus strains and constitutes a separate GH70 subfamily. Appl Microbiol Biot 97:181-193. 15. Bai YX, van der Kaaij RM, Leemhuis H, Pijning T, van Leeuwen SS, Jin ZY, Dijkhuizen L. 2015. Biochemical characterization of the Lactobacillus reuteri Glycoside Hydrolase family 70 GTFB type of 4,6-α-glucanotransferase enzymes that synthesize soluble dietary starch fibers. Appl Environ Microb 81:7223-7232.

16. Dijkhuizen L, van der Maarel MJEC, Kamerling JP, Leemhuis H, Kralj S, JM D. 2010. Gluco-oligosaccharides comprising (α1→4) and (α1→6) glycosidic bonds, use thereof, and methods for providing them. WO/2010/128859.

17. Dobruchowska JM, Gerwig GJ, Kralj S, Grijpstra P, Leemhuis H, Dijkhuizen L, Kamerling JP. 2012. Structural characterization of linear isomalto-/malto-oligomer products synthesized by the novel GTFB 4,6-α-glucanotransferase enzyme from Lactobacillus reuteri 121. Glycobiology 22:517-528.

18. Kabsch W. 2010. Xds. Acta Crystallogr D 66:125-132. 19. Evans PR, Murshudov GN. 2013. How good are my data and what is the resolution? Acta Crystallogr D 69:1204-1214. 20. Mccoy AJ, Grosse-Kunstleve RW, Adams PD, Winn MD, Storoni LC, Read RJ. 2007. Phaser crystallographic software. J Appl Crystallogr 40:658-674.

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23. Emsley P, Lohkamp B, Scott WG, Cowtan K. 2010. Features and development of Coot. Acta Crystallogr D 66:486-501.

24. Chen VB, Arendall WB, Headd JJ, Keedy DA, Immormino RM, Kapral GJ, Murray LW, Richardson JS, Richardson DC. 2010. MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr D 66:12-21.

25. Krissinel E, Henrick K. 2004. Secondary-structure matching (SSM), a new tool for fast protein structure alignment in three dimensions. Acta Crystallogr D 60:2256-2268. 26. Pijning T, Vujicic-Zagar A, Kralj S, Dijkhuizen L, Dijkstra BW. 2012. Structure of the α-1,6/α-1,4-specific glucansucrase GTFA from Lactobacillus reuteri 121. Acta Crystallogr F 68:1448-1454. 27. Ito K, Ito S, Shimamura T, Weyand S, Kawarasaki Y, Misaka T, Abe K, Kobayashi T, Cameron AD, Iwata S. 2011. Crystal structure of glucansucrase from the dental caries pathogen Streptococcus mutans. J Mol Biol 408:177-186. 28. Lyhne-Iversen L, Hobley TJ, Kaasgaard SG, Harris P. 2006. Structure of Bacillus halmapalus α-amylase crystallized with and without the substrate analogue acarbose and maltose. Acta Crystallogr F 62:849-854. 29. Fujiwara T, Terao Y, Hoshino T, Kawabata S, Ooshima T, Sobue S, Kimura S, Hamada S. 1998. Molecular analyses of glucosyltransferase genes among strains of Streptococcus mutans. FEMS Microbiol Lett 161:331-336. 30. Edgar RC. 2004. MUSCLE: a multiple sequence alignment method with reduced time and space complexity. BMC Bioinformatics 5:1-19. 31. Robert X, Gouet P. 2014. Deciphering key features in protein structures with the new ENDscript server. Nucleic Acids Res 42:W320-W324.

32. Machius M, Declerck N, Huber R, Wiegand G. 1998. Activation of Bacillus licheniformis α-amylase through a disorder → order transition of the substrate-binding site mediated by a calcium-sodium-calcium metal triad. Structure 6:281-292.

33. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. 2013. MEGA6: molecular evolutionary genetics analysis version 6.0. Mol Biol Evol 30:2725-2729.

34. Davies GJ, Wilson KS, Henrissat B. 1997. Nomenclature for sugar-binding subsites in glycosyl hydrolases. Biochem J 321:557-559. 35. Fujimoto Z, Takase K, Doui N, Momma M, Matsumoto T, Mizuno H. 1998. Crystal structure of a catalytic-site mutant α-amylase from Bacillus subtilis complexed with maltopentaose. J Mol Biol 277:393-407. 36. Uitdehaag JCM, Mosi R, Kalk KH, van der Veen BA, Dijkhuizen L, Withers SG, Dijkstra BW. 1999. X-ray structures along the reaction pathway of cyclodextrin glycosyltransferase elucidate catalysis in the α-amylase family. Nat Struct Biol 6:432-436.

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37. Davies GJ, Brzozowski AM, Dauter Z, Rasmussen MD, Borchert TV, Wilson KS. 2005. Structure of a Bacillus halmapalus family 13 α-amylase, BHA, in complex with an acarbose-derived nonasaccharide at 2.1 angstrom resolution. Acta Crystallogr D 61:190-193. 38. Meng XF, Dobruchowska JM, Pijning T, Lopez CA, Kamerling JP, Dijkhuizen L. 2014. Residue Leu(940) Has a crucial role in the linkage and reaction specificity of the glucansucrase GTF180 of the probiotic Bacterium Lactobacillus reuteri 180. J Biol Chem 289:32773-32782. 39. Kralj S, van Geel-Schutten IGH, Faber EJ, van der Maarel MJEC, Dijkhuizen L. 2005. Rational transformation of Lactobacillus reuteri 121 reuteransucrase into a dextransucrase. Biochemistry 44:9206-9216. 40. Kralj S, Eeuwema W, Eckhardt TH, Dijkhuizen L. 2006. Role of asparagine 1134 in glucosidic bond and transglycosylation specificity of reuteransucrase from Lactobacillus reuteri 121. FEBS J 273:3735-3742. 41. Hoshino T, Fujiwara T, Kawabata S. 2012. Evolution of cariogenic character in Streptococcus mutans: horizontal transmission of Glycosyl Hydrolase family 70 genes. Sci Rep-UK 2:518. 42. Qian WF, Zhang JZ. 2014. Genomic evidence for adaptation by gene duplication. Genome Res 24:1356-1362.

43. Sanchez-Perez G, Mira A, Nyiro G, Pasic L, Rodriguez-Valera F. 2008. Adapting to environmental changes using specialized paralogs (vol 24, pg 154, 2008). Trends Genet 24:327-327.

44. Passerini D, Vuillemin M, Ufarte L, Morel S, Loux V, Fontagne-Faucher C, Monsan P, Remaud-Simeon M, Moulis C. 2015. Inventory of the GH70 enzymes encoded by Leuconostoc citreum NRRL B-1299 - identification of three novel α-transglucosylases. FEBS J 282:2115-2130. 45. Simpson CL, Cheetham NWH, Giffard PM, Jacques NA. 1995. 4 glucosyltransferases, GtfJ, GtfK, GtfL and GtfM, from Streptococcus salivarius ATCC-25975. Microbiology-UK 141:1451-1460. 46. Fujiwara T, Hoshino T, Ooshima T, Sobue S, Hamada S. 2000. Purification, characterization, and molecular analysis of the gene encoding glucosyltransferase from Streptococcus oralis. Infect Immun 68:2475-2483. 47. Hakenbeck R, Balmelle N, Weber B, Gardes C, Keck W, de Saizieu A. 2001. Mosaic genes and mosaic chromosomes: Intra- and interspecies genomic variation of Streptococcus pneumoniae. Infect Immun 69:2477-2486. 48. Van Geel-Schutten GH, Faber EJ, Smit E, Bonting K, Smith MR, Ten Brink B, Kamerling JP, Vliegenthart JFG, Dijkhuizen L. 1999. Biochemical and structural

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characterization of the glucan and fructan exopolysaccharides synthesized by the Lactobacillus reuteri wild-type strain and by mutant strains. Appl Environ Microb 65:3008-3014.

49. Kralj S, van Geel-Schutten GH, Rahaoui H, Leer RJ, Faber EJ, van der Maarel MJEC, Dijkhuizen L. 2002. Molecular characterization of a novel glucosyltransferase from Lactobacillus reuteri strain 121 synthesizing a unique, highly branched glucan with α-(1→4) and α-(1→6) glucosidic bonds. Appl Environ Microb 68:4283-4291.

50. Sims IM, Frese SA, Walter J, Loach D, Wilson M, Appleyard K, Eason J, Livingston M, Baird M, Cook G, Tannock GW. 2011. Structure and functions of exopolysaccharide produced by gut commensal Lactobacillus reuteri 100-23. ISME J 5:1115-1124.

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Summary and thesis discussion

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Starch is one of the major carbohydrates on earth and is the main dietary carbohydrate in our life. Apart from the raw starches, many starch derivatives produced via physical, chemical, and enzymatic treatments are abundantly applied in food, paper, and textile industries because of their distinctive properties. Among these approaches, the enzymatic treatment has advantages such as low energy consumption, low cost, high controllability, high productivity. Therefore, more and more starch-acting enzymes including hydrolases and transglycosidases are introduced in food and pharmaceutical manufacturing processes to produce glucose and fructose syrup, cyclodextrins, thermoreversible gels or glycogen, or to improve the flour and dough for baking, etc.

4,6-α-Glucanotransferases (4,6-α-GTases) are recently reported enzymes that predominately cleave an α1→4 bond from the non-reducing ends of starch or maltodextrins and transfer the cleaved glucose unit to the non-reducing ends of the acceptor substrate, mostly introducing an α1→6 bond and occasionally an α1→4 bond, forming isomalto-/malto-oligosaccharides (IMMO) and polysaccharides (IMMP). Unlike other starch hydrolyzing enzymes, which are general members of Glycoside Hydrolase family (GH) 13, 57, or 77, 4,6-α-GTases are classified in a subfamily of GH70; family GH70 is mainly composed of glucansucrases (GS). The GS including dextransucrase, mutansucrase, reuteransucrase, and alternansucrase, are active on sucrose, producing α-glucans with various linkage types such as α1→2, α1→3, α1→4, and α1→6 linkages. Like GS, 4,6-α-GTases are exclusively found in lactic acid bacteria (LAB). However, 4,6-α-GTases are completely inactive on sucrose but active on starch. Based on the primary structure analysis, the 4,6-α-GTases are homologous to GS. Both types of enzymes possess a circularly permutated (α/β)8-barrel acting as catalytic core, and the same domain organization consisting of a variable N-terminal region, domains A, B, C, and extra domains IV and V (Chapter 1).

Production and characterization of 4,6-α-GTases The IMMP produced by 4,6-α-GTases is of commercial potential as it can serve as soluble dietary fiber and potentially as a prebiotic. The 4,6-α-GTases-modified starches also may have special rheological properties, and thus may find applications in the preparation of novel and valuable food products. However, the introduction of 4,6-α-GTases in industrial processes is still a challenge.

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GtfB of Lactobacillus reuteri 121 is the first described representative of 4,6-α-GTases. The IMMP produced by GtfB has been well investigated. The yield of soluble full-length GtfB enzyme expressed in Escherichia coli is poor and cannot meet the demand of downstream research and industrial production. A large quantity of GtfB protein accumulates in inclusion bodies (Chapter 2). The traditional denaturing-refolding procedures resulted in a high recovery of activity even in double distilled H2O. Moreover, the inclusion bodies themselves also display some enzymatic activity without any need for further processing. They therefore are regarded as non-classical inclusion bodies (ncIBs), which are known to retain activity due to partial preservation of the native protein structure. The native, refolded, and ncIB GtfB enzymes all produce IMMP with similar size distribution. In addition, ncIB GtfB protein clearly has a higher thermostability. Active GtfB protein from inclusion bodies in the form of refolded or ncIB GtfB enzyme may thus find industrial application in synthesis of IMMP modified starch. However, the expression level of soluble enzymes, which is preferable for industrial production, has remained unsatisfactorily low. The LAB GS are homologous to 4,6-α-GTase enzymes, and deletion of (parts of) their N-terminal and/or C-terminal domains successfully enhanced the yield of soluble active GS enzyme. GtfB differs from GS in lacking the C-terminal polypeptide segment of domain V. Notably, truncation of the variable N-terminal region (1-733 amino acids) resulted in a dramatic increase of the yield of soluble GtfB∆N enzyme (approx. 75 fold compared to full length GtfB) without influencing the activity of the enzyme and products formed (Chapter 3).

Development of proper activity assays for the characterization of 4,6-α-GTases is vital for their further biochemical characterization and development of industrial applications. Existing assays do not allow determination of the specific hydrolysis and transglycosylation activities nor a kinetic analysis. Therefore, we developed a new assay derived from the traditional amylose-iodine staining assay (Chapter 3). Based on the theory of steric hindrance, the assay allows accurate determination of both hydrolysis and total activity, and calculation of the transglycosylation activity of 4,6-α-GTases. With this assay, the biochemical properties of 4,6-α-GTases have been characterized in detail, including determination of hydrolysis (H) and transglycosylation (T) activities, kinetic parameters, and acceptor

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substrate specificity. Based on the developed assay, a ratio of T/H=4.3:1 for GtfB-∆N was obtained under the optimal conditions.

Crystal structure of GtfB Although the structure of IMMP produced by GtfB has been characterized, the catalytic mechanism and structure-function relationships of 4,6-α-GTases remained largely unclear. Chapter 6 reports the high resolution crystal structure of a truncated but fully active fragment of GtfB (GtfB-ΔNΔV) from L. reuteri 121. GtfB-ΔNΔV displays a GS-like domain arrangement; however unlike GS, the active site surprisingly features a long α-amylase-like binding groove with multiple donor subsites in a tunnel. The presence of this tunnel, formed by three loops unique to 4,6-α-GTases, explains why GtfB processes only the linear external chains of starch, thus preferring high-amylose starches as substrate. By combining structural information from an enzyme-donor substrate complex with analysis of acceptor specificity we can now fully explain the observed product spectrum of 4,6-α-GTases with a new mechanism, in which the enzyme transfers not only single, but also multiple glucosyl units to acceptor substrates. In addition, the GtfB-ΔNΔV crystal structure clearly supports the idea that the 4,6-α-GTase GH70 subfamily is an evolutionary intermediate between α-amylase and glucansucrase enzymes.

Biological role of 4,6-α-GTases in Lactobacillus reuteri GS more widely exist in LAB strains and are known to contribute to the formation of exopolysaccharides (EPS), which play a role in the biofilm formation. In the oral cavity, the produced biofilm containing insoluble EPS enhances the adhesion of bacteria to tooth surfaces and consequently stimulates dental caries formation. EPS may also have other functionalities for the strain such as retention of water, nutrient source, protective barrier, sorption of organic compounds, immobilization of enzymes, and so on. As 4,6-α-GTase is able to produce polysaccharide from maltodextrins or starch instead of sucrose in vitro, the question was raised whether 4,6-α-GTase also contributes to the in vivo EPS formation in their host LAB (Chapter 4). In the model strain L. reuteri 121, the gtfB gene was found to be essential for EPS formation when the strain was grown on maltodextrins or starch as carbon source. L. reuteri 121 can modify both amylose and external chains of amylopectin in starch by converting α1→4

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linkages into α1→6 linkages. The formed EPS have the same assembling segments as IMMP produced in vitro by GtfB. In addition, the EPS can be used as carbon and energy source by some probiotic Bifidobacterium strains, revealing that the EPS are potentially prebiotic. Therefore, the L. reuteri strains that can produce dietary fiber and potential prebiotics may find applications in the fermented food industry (as synbiotics).

In the oral cavity, Lactobacillus and Streptococcus strains are ubiquitous bacteria and have been proven to produce EPS from sucrose through their glucansucrase. Sucrose and starch are major carbohydrates in the modern diet and their cariogenicity has been widely investigated. Although the cariogenicity of starch alone is a matter of debate, uptake of a combination of starch and sucrose is known to enhance cariogenicity by influencing the polysaccharide formation in biofilm. The mechanism is not fully understood. The discovery of the tandem glucansucrase GtfA encoding gene gtfA and 4,6-α-GTase GtfB encoding gene gtfB in L. reuteri strains may provide new insights with regard to cariogenicity through biofilm formation. Both in vivo and in vitro experimental results indicated a synergistic action of 4,6-α-GTase GtfB and GS GtfA in the presence of both starch and sucrose (Chapter 5). Therefore, a new pathway of EPS formation from multiple dietary carbohydrates in LAB strains was found and may contribute to the research on dental caries.

Opportunities and Challenges 4,6-α-GTases producing soluble dietary fiber and potential prebiotics from starch or maltodextrins are likely to draw strong attention from the food industry including bakery, beverage, sausage, and so on. The produced IMMP with certain percentages of α1→4 and α1→6 linkages may also function to stabilize protein during lyophilization, to enhance the solubility of proteins and drug molecules, to substitute fat, to replace starch as low-calorie bulking agent. In addition, 4,6-α-GTase-modified starches may have typical rheological properties and thus be used in many aspects. The elucidation of the three-dimensional structure of GtfB-∆N∆V will support the further engineering of this protein, e.g. aiming to change its substrate specificity or to synthesize products with different linkage specificity. Moreover, the discovery of Lactobacillus strains carrying 4,6-α-GTase encoding genes opens up new prospectives for the application of these strains in food

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fermentations. In addition to the regularly produced nutrients, such as short chain fatty acids, these Lactobacillus strains synthesize soluble dietary fiber and potential prebiotics by converting starch that is abundantly present in fermented foods, such as sourdough, soy bean sauce.

However, many questions still challenge the application of 4,6-α-GTase. Firstly, how 4,6-α-GTase exactly modifies starch especially amylopectin is unclear. One possibility is that the α1→4-linked side chains of amylopectin are coverted into α1→6-linked chains, which are still located at the outside of amylopectin molecules. The other possibility is that the α1→4-linked side chains of amylopectin are cleaved and formed separate α1→6-linked chains.Secondly, the mechanism and location for IMMP degradation in the human intestinal tract remains to be investigated. Thirdly, the thermostability and activity of 4,6-α-GTase remain to be improved by enzyme engineering. For this purpose, computational design is preferable. Moreover, for application of Lactobacillus strains that possess 4,6-α-GTases, improvement of their expression levels and enhancement of their activities under fermentation conditions are challenges. But such challenges constitute the charm of research on 4,6-α-GTases.

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Samenvatting en discussie

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Zetmeel is een van de belangrijkste koolhydraten op aarde en een belangrijke component van het humane dieet. Voor de talloze toepassingen van zetmeel in voedsel, papier- en textielindustrie en farmacie worden zetmeelderivaten geproduceerd via fysische, chemische en enzymatische behandelingen. Enzymatische omzetting van zetmeel kent verschillende voordelen zoals een laag energieverbruik, lage kosten, hoge bestuurbaarheid en hoge productiviteit. Daarom worden steeds meer enzymen die op zetmeel werken, zoals hydrolases en transglycosidases, geïntroduceerd in de levensmiddelen- en farmaceutische industrie voor de productie van ingrediënten zoals glucose- en fructosestroop, cyclodextrines en thermoreversibele gelen, en voor de structuurverbetering van deeg en brood, etc.

4,6-α-Glucanotransferases (4,6-α-GTases) zijn recent ontdekte enzymen die een α1→4 binding van het niet-reducerende uiteinde van zetmeel of maltodextrines hydrolyseren en het afgesplitste (glucose)molecuul overbrengen naar het niet-reducerende uiteinde van een acceptor substraat. Meestal wordt hierbij een α1→6 glycosidische binding gevormd, en af en toe een α1→4 glycosidische binding. Op deze wijze worden isomalto-/malto- oligosacchariden (IMMO) en polysacchariden (IMMP) gevormd. In tegenstelling tot andere zetmeel hydrolyserende enzymen, die meestal behoren tot de glycoside hydrolase familie (GH) 13, 57, of 77, zijn 4,6-α-GTases ingedeeld in een onderfamilie van GH70; GH70 bestaat voornamelijk uit glucansucrases (GS). De GS enzymen, dextransucrase, mutansucrase, reuteransucrase en alternansucrase, gebruiken sucrose voor de productie van α-glucanen met verschillende soorten bindingstypen zoals α1→2, α1→3, α1→4 en α1→6 glycosidische bindingen. Net als GS worden 4,6-α-GTases uitsluitend gevonden in melkzuurbacteriën (Lactic Acid Bacteria, LAB). In tegenstelling tot GS zijn 4,6-α-GTases inactief op sucrose, maar actief op zetmeel. Beide typen enzymen bezitten een circulair gepermuteerde (α/β)8-cilinder als katalytische kern en ze hebben dezelfde domein opbouw, die bestaat uit een variabel N-terminaal gedeelte, domeinen A, B, C, en extra domeinen IV en V (Hoofdstuk 1).

Productie en karakterisatie van 4,6-α-GTases

De IMMP geproduceerd door 4,6-α-GTases kunnen op termijn mogelijk worden toegepast in voedingsmiddelen, zoals oplosbare voedingsvezels met een

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prebiotische werking. Zetmeel dat enzymatisch is gemodificeerd met deze enzymen kan ook speciale rheologische eigenschappen hebben die interessant zijn voor toepassing in de voedingsmiddelen industrie. Echter, de industriële toepassing van 4,6-α-GTases vormt voorlopig nog een uitdaging.

GtfB van Lactobacillus reuteri 121 is het eerst beschreven 4,6-α-GTase. De productie van IMMP door GtfB is goed onderzocht. Wanneer het GtfB enzym tot expressie wordt gebracht in Escherichia coli is de opbrengst laag, en onvoldoende voor toepassingen op industriële schaal. Echter, een grote hoeveelheid GtfB eiwit hoopt zich in onoplosbare vorm op in zogenaamde inclusionbodies (Hoofdstuk 2) in de cellen van E. coli. De traditionele manier om hieruit actief enzym te halen, via het denatureren en hervouwen van het eiwit, resulteerde in een aanzienlijk herstel van de enzymatische activiteit. Daarnaast vertoonden de inclusionbodies zelf ook enige enzymatische activiteit. Dit soort inclusionbodies, waarvan de activiteit gedeeltelijk behouden blijft, worden beschouwd als ‘niet-klassieke inclusion bodies’ (ncIBs). De verschillende GtfB enzymen (oplosbaar, hervouwen en ncIB) produceren allemaal IMMP met vergelijkbare grootte. Daarnaast hebben de ncIB enzymen een hogere thermostabiliteit. GtfB geproduceerd via hervouwing of in de vorm van ncIB kan daarmee een bijdrage leveren aan de productie van IMMP en gemodificeerd zetmeel op industriële schaal.

Van glucansucrases is bekend dat deletie van het N-terminale en/of C-terminale domein de heterologe expressie aanzienlijk kan verbeteren. GtfB, homoloog aan glucansucrases, verschilt onder meer van GS vanwege het ontbreken van het C-terminale deel van domein V. Deletie van het N-terminale gebied van GtfB (aminozuren 1-733) resulteerde in een verhoogde opbrengst van oplosbaar GtfB-ΔN enzym (75 voudige toename) zonder dat daardoor de activiteit van het enzym en de gevormde producten werden beïnvloed (Hoofdstuk 3).

De ontwikkeling van geschikte assays voor bepaling van de activiteiten van 4,6-α-GTases is essentieel voor zowel hun gedetailleerde biochemische karakterisering als voor de ontwikkeling van industriële toepassingen. Bestaande assays kunnen geen onderscheid maken tussen de specifieke hydrolyse en transglycosylerings activiteit van zulke enzymen, waardoor het niet mogelijk is om een kinetische analyse te doen. Daarom hebben we een nieuwe methode

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ontwikkeld die is gebaseerd op de traditionele amylose jodium kleuringsassay (Hoofdstuk 3). Met deze assay kan zowel de hydrolyse als de totale activiteit nauwkeurig bepaald worden, waarna de transglycosylerings activiteit kan worden berekend. Hiermee hebben we de 4,6-α-GTases biochemisch gekarakteriseerd, waaronder bepaling van hydrolyse (H) en transglycosylerings (T) activiteit, substraatspecificiteit en acceptor efficiëntie. De verhouding T/H voor GtfB-ΔN is met behulp van deze assay vastgesteld op 4,3:1.

Kristalstructuur van GtfB

Om beter inzicht te krijgen in het katalytische mechanisme van 4,6-α-GTases hebben we de kristalstructuur bepaald van een (volledig actief) fragment van GtfB (GtfB-ΔNΔV) uit L. reuteri 121 (Hoofdstuk 6). Zoals voorspeld lijkt de structuur van GtfB-ΔNΔV sterk op die van glucansucrases, met een vergelijkbare domeinstructuur. In tegenstelling tot GS bevat het katalytisch centrum een verrassend lange groef met daarin meerdere donor substraat bindingsplaatsen. De aanwezigheid van deze tunnel, die gevormd wordt door drie eiwitlussen die uniek zijn in 4,6-α-GTases, verklaart dat GtfB alleen de lineaire ketens van zetmeel als donorsubstraat kan gebruiken, en verklaart ook de voorkeur van het enzym voor substraten met een hoog amylose gehalte. Door inzicht in de enzymstructuur te combineren met kennis over de acceptor specificiteit, kunnen we nu het productspectrum van 4,6-α-GTase verklaren met een nieuw mechanisme, waarbij het enzym één of meerdere glucosyl eenheden tegelijk naar een acceptor substraat kan overbrengen. Daarnaast ondersteunt de kristalstructuur van GtfB-ΔNΔV het idee dat de 4,6-α-GTase subfamilie een evolutionair intermediair vormt tussen α-amylase en glucansucrase enzymen.

Biologische rol van 4,6-α-GTases in Lactobacillus reuteri

Glucansucrases komen voor in melkzuurbacteriën en staan erom bekend bij te dragen aan de vorming van exopolysaccharides (EPS), die een rol spelen bij de vorming van biofilms. Bacteriële biofilms zorgen voor hechting van bacteriën aan bijvoorbeeld het tandoppervlak wat uiteindelijk kan leiden tot cariës. Voor de bacteriën heeft EPS ook nog andere functies zoals waterretentie, gebruik als reserve voedingsbron, bescherming tegen externe factoren, opname van organische verbindingen, immobilisatie van enzymen, enzovoort. 4,6-α-GTase vormt in vitro een polysaccharide uit maltodextrinen of zetmeel, daarom werd in

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hoofdstuk 4 onderzocht of 4,6-α-GTase ook in vivo bijdraagt aan de vorming van EPS in melkzuurbacteriën (Hoofdstuk 4). In de modelstam L. reuteri 121 is vastgesteld dat het gen dat codeert voor GtfB van essentieel belang is voor de vorming van EPS bij groei op maltodextrines of zetmeel als koolstofbron. L. reuteri 121 kan in deze condities amylose en amylopectine omzetten waarbij nieuwe met name α1 → 6 glycosidische bindingen worden gevormd. Het in vivo gevormde EPS bestaat uit dezelfde fragmenten als IMMP dat in vitro wordt geproduceerd door GtfB. Het gevormde EPS kan worden gebruikt als koolstof- en energiebron voor de groei van sommige prebiotische bifidobacteriën; het EPS is dus een potentieel prebioticum. De L. reuteri stammen die deze voedingsvezels en potentiële prebiotica kunnen produceren bieden interessante mogelijkheden voor toepassingen in de productie van gefermenteerde levensmiddelen.

Bacteriën behorende tot de geslachten Lactobacillus en Streptococcus komen voor in de mondholte, waar ze EPS produceren uit sucrose met hun glucansucrase enzymen. Sucrose en zetmeel zijn belangrijke koolhydraten in het moderne humane dieet. Van sucrose is bekend dat het cariogeen is, terwijl de rol van zetmeel hierin nog ter discussie staat. Wel is vastgesteld dat een combinatie van zetmeel en sucrose de vorming van biofilms beïnvloedt en daarmee een effect heeft op cariës. Het mechanisme hiervan is niet duidelijk. De ontdekking van de aanwezigheid van glucansucrase GtfA én 4,6-α-GTase GtfB in L. reuteri stammen kan leiden tot nieuwe inzichten ten aanzien van de vorming van biofilms en cariogeniciteit. Zowel in vivo als in vitro resultaten laten een synergistische werking zien van GtfB en GtfA in de aanwezigheid van zowel zetmeel als sucrose (Hoofdstuk 5). Daarmee is een nieuwe route aangetoond voor de vorming van EPS door melkzuurbacteriën uit verschillende koolhydraten; bestudering hiervan kan bijdragen aan het onderzoek naar tandcariës.

Kansen en uitdagingen

4,6-α-GTases produceren oplosbare voedingsvezels en potentiële prebiotica uit zetmeel of maltodextrinen, die mogelijk toepasbaar zijn in de voedingsindustrie. Andere mogelijke toepassingen van IMMP zijn het stabiliseren van eiwitten tijdens vriesdrogen, verbeteren van de oplosbaarheid van eiwitten en

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geneesmiddelen, vervangen van vet in voedingsmiddelen en vervanging van zetmeel met een caloriearme vulstof. Het is mogelijk dat 4,6-α-GTase-gemodificeerd zetmeel speciale rheologische eigenschappen heeft die specifieke toepassingen mogelijk maakt. De driedimensionale structuur van GtfB-ΔNΔV is van belang voor de verdere engineering van dit eiwit, bijvoorbeeld voor de aanpassing van de substraatspecificiteit of synthetiseren van producten met verschillende percentages glycosidische bindingen. De ontdekking van Lactobacillus stammen met een gen coderend voor 4,6-α-GTase opent nieuwe perspectieven voor de toepassing van deze bacteriën in levensmiddelen.

Er bestaan echter ook nog veel onzekerheden rond de industriële toepassing van 4,6-α-GTases. Ten eerste, het is onduidelijk hoe 4,6-α-GTase precies de modificatiereactie op zetmeel uitvoert. Eén van de mogelijkheden is dat de α1→4 gelinkte amylopectin zijketens worden omgezet in α1→6 gelinkte ketens, welke zich nog steeds aan de buitenkant van de amylopectin moleculen bevinden. De andere mogelijkheid is dat de α1→4 gelinkte zijketens van amylopectin worden gekloofd zodat losse α1→6 gelinkte ketens worden gevormd. Ten tweede, het mechanisme en lokatie van de afbraak van IMMP in de spijsvertering van de mens moet nog nader worden onderzocht. Ten derde, de thermostabiliteit en de activiteit van 4,6-α-GTase moeten aanzienlijk worden verbeterd voordat gebruik op industriële schaal mogelijk wordt. Hiervoor, heeft een computerontwerp de voorkeur. Voor directe toepassing van Lactobacillus stammen met 4,6-α-GTase is het noodzakelijk dat ook het expressieniveau en de activiteit onder fermentatiecondities worden verbeterd. Deze relatief grote uitdagingen vormen tegelijkertijd ook de charme van het onderzoek aan 4,6-α-GTases.

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Acknowledgements

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At this very moment of completing the thesis, the precious memories generated in the past three years all come to my mind. From China to the Netherlands, from a joint PhD candidate of Jiangnan University and Groningen University to a regular PhD student of Groningen University, from food science to microbial physiology, from analyzing physiochemical properties of food components to characterizing the carbohydrate acting enzymes, all these transformations are full of difficulties. I cannot imagine that I could succeed in these transformations without the efforts from the friendly people surrounding me. I feel lucky that I can meet so many nice people in such a lovely country. I would like to take this opportunity to say “thanks” to all of them.

Firstly and most importantly, I would like to express my sincerest appreciation and deepest gratitude to my promoter, Prof. Lubbert Dijkhuizen. Thank him for kindly accepting me to join Microbial Physiology group as a joint PhD student and providing me with a full PhD position in Groningen. Thank him for giving me a promising topic, which let me involve in the knowledgeable Carbohydrate Competence Center and made me love in scientific research. I will never forget all the fruitful and inspiring discussions with him in the past years. His erudition, wisdom, enthusiasm, preciseness and kindness deeply influenced me and helped me to develop a positive attitude towards scientific research. It is a great treasure and fortune of mine to have him as my supervisor during this precious period of my life. I really appreciate that he had always tried to make his word simple and easy to be understandable in my first studying year. I will never forget his kindness to my requests. I will also keep in mind that he always encouraged me to be active in showing my research by posters and oral presentations in all possible occasions and to communicate with other scientists from various institutes or companies. I appreciate that he, as a director of many reputable organizations, offered me opportunities to meet and discuss with so many important persons from Universities, industries, and governments in many high level meetings. I will always remember that he worked on my chapters and discussed with me via email in many midnights and non-working days. He is a father-like supervisor who tries to make me independent, strict and responsible. I sincerely wish him and his family a healthy and happy life.

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I would like to give my grateful thank and appreciation to my copromoter and daily supervisor, Dr. Rachel van der Kaaij. Without her help, I cannot finish the thesis efficiently and I may still be struggling with my research at this moment. I will never forget that she had spent a lot of time on revising and reconstructing my manuscripts and chapters time by time. She taught me many basic concepts in microbiology and enzymology. She is not only a supervisor but also a kind friend of mine. I appreciate her patience when listening my reports and complaints. I will remember every inspired discussion in her offices in either Linnaeusborg or Bernoulliborg. I appreciate all the constructive and valuable advices from her. I also thank her for helping me to translate the summary of my thesis in Dutch.

My sincere and grateful thanks go to Prof. Zhengyu Jin, who is my Chinese supervisor in my joint PhD program in Jiangnan University. Thank him for providing me with the hard-won position to work in his group as a master student. Thank him for giving me the opportunity to be a combined PhD/master student in his group. Thank him for supervising me and introducing me to the carbohydrate field since my master period. I appreciate that he provided me with the scarce opportunity to study abroad, indicated the direction of my oversea research, and encouraged me to finish my PhD study in Groningen. His kindness to all of my requests in the past years will be kept in my mind forever. I also appreciate that he gave me opportunities to visit various companies in China and to participate international conferences, which broadened my vision and helped me to establish a right attitude to scientific research.

My deep gratitude also goes to Prof. Xueming Xu, who had helped me a lot during my master and PhD periods. Thank him for providing me with many brilliant ideas on my topics and encouraging me to continue my study abroad.

To my Chinese daily supervisor, Prof. Yaoqi Tian, I am deeply grateful for his support and guidance during my master and early PhD periods. I will never forget all the moments we discussed together in the studying room when he was still a PhD candidate. I was so impressed by his intelligence, diligence and kindness. I appreciate that he taught me how to organize a story for publication in international journals and helped me to correct my sentences word by word during my master period.

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Here, I also would like to thank Prof. Johannis P. Kamerling. I will cherish all the times we met in the past years. I will remember all the great ideas from him. I also appreciate that you kindly wrote a recommendation letter for me when I was looking for a position in China.

My gratitude goes to the members of the reading committee Prof. T. Desmet, Prof. D.B. Janssen, and Prof. M.J.E.C. van der Maarel. Thank them for spending time on critically reading my thesis and giving valuable comments, and approving the thesis for the public defence.

My deep appreciation goes to Dr. Tjaard Pijning. His profound knowledge on the structure of carbohydrate acting enzymes is so impressive. I thank him for teaching me so much on understanding the crystal structure of carbohydrate acting enzymes. I really enjoyed all the inspired discussions with him. I will never forget the days we sit together and discuss on how to organize and write a excellent story from our results. I also appreciate his help on designing the beautiful cover of my thesis.

I would like to thank Joana Gangoiti and Markus Böger, who are willing to be my paranimfs. I am glad to work and give talk together with Joana in the same project for AVEBE in my first two years. Thank her for many wonderful discussions on our topics. I appreciate Markus’ help on growth of anaerobic bacteria using the system established by him and his contribution on the chapter 4 of this thesis.

Of course, my thanks also go to my other two officemates, Ana Ceniceros and Tim Devlamynck, who brought me so much warmness. Thank them for chatting with me on various topics including research, cultures and politics. Thank them for providing me so much useful information to make me live here happy. Thank them for accompanying with me in the office after working time in many days. I wish you all a nice promotion in the near future.

I would like to thank Justyna Dobruchowska, Sander van Leeuwen and Gerrit Gerwig for the structural analysis of the saccharides. They always offered kind supports even though they are very busy with other work at the same time. I appreciate their patient explanation on interpreting the NMR spectra and contribution to some chapters in this thesis.

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I would like to express my appreciation to Piet Buwalda and Piet van der Zaal from AVEBE starch company and Wageningen University for all the worthy discussion and knowledge sharing in the project meetings.

Grateful thanks go to Albert Woortman and Gert Alberda van Ekenstein for helping to do the SEC and DSC analysis. Thank Albert for contributing to some of my chapters. I appreciate that they patiently explained the principle of all the technologies and provided me with many useful suggestions.

My thanks go to Prof. Jan Kok and Harma Karsens for kindly offering me help on gene inactivation in Lactobacillus.

My gratitude goes to Pieter. I appreciate his technical support at my first year in the lab. I will never forget the word “backup” from you.

Sincere thanks go to our secretaries Bea, Manon, and Anmara, and managing director of GBB, Engel for their kind help on a series of issues such as identity transformation, extension, arrangement of conference and so on.

I should not forget to thank Tseard de Graaf, who is expertise on learning and teaching foreign languages. Thank him for giving me a warm welcome, chatting with me in my first journey from Amsterdam to Groningen, and showing me around the Museum of Groningen University.

I would like to thank Xiangfeng Meng for helping me to settle quickly into the Dutch life-style and laboratory work. I thank him for all the valuable discussions on the work. Also, thank Huifang Yin for the work discussion during lunchtime.

I am grateful to all the former and present colleagues in MicFys including Slavko, Hans, Jelle, Jolanda, Evelien, Cecile, Alle, Alicia, Elena, Lara, Mark, Mirjan, Hans-Jörg, Marnix, Veronica, Gerda, Maarten, Vincent, Laura, Hien, Rivca, Sahar, Ahmed, Sebastiaan, Geralt, Marta. I will never forget their supports, which made my PhD fruitful. My deep gratitude also goes to my colleagues in Jiangnan University including Xing Zhou, Jianwei Zhao, Jinpeng Wang, Zhengjun Xie, Aiquan Jiao, Huanxin Zhang, Bo Yu, Wangyang Shen, Fengfeng Wu, Yisheng Chen, Huanlong Liu, Na Yang, Jin Xu, Xu Cao, Guangyao Chen, Xuexia Ji, Jing Chen, Yawei Ning, Jun Nie, Xiuting Hu, Benxi

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Wei, Chunsen Wu, Jie Long, Hongyan Li, Xiuchao Zhai, Yan Xu, Yan Yue, Haining Zhuang, Yamei Jin, Zhixuan Sheng, Qianqian Jiang, Lulu Zhang, Tiantian Wang, Bao Zhang, Tao Feng. I appreciate all the support from them and I really enjoyed the happy time with them.

I would like to thank all my Chinese friends in the Netherlands: Wanli Zheng, Peng Peng, Feng Hu, Bian Wu, Wenjuan Ma, Yanping Geng, Qiuyan Yang, Pei Zhao, Nana Yu, Youtao Liu, Min Wu, Xuewen Zhang, Mengqiu Ge, Yan Zeng, Jiquan Wu, Xiaoyu Ruan, Di Wang, Yuxi Tang, Fei Deng, Jinfeng Shao, Pu Yang, Yirenjile, Jiawei Chen, Wenjun Wang, Tao Zhang, Wenxuan Zhang, Sheng He, Yinxing Jin, Dongdong Mu, Gang Huang, Juanjuan Su, Qian Li, Haojie Cao, Yanglei Yi, Jingjing Deng, Xin Zhao, Boqun Liu, Zhiwei Zhou, Ximin Wang, Shengcheng Ren, Fangjie Gu, Sheng He, Mingbo Liu, Michael, Jia Liu. Bin Jiang, Yuanyuan Ju, Jinfeng Peng, Xuan Xu, Qihan Zhang and so on. Their enthusiasm and goodness let me feel warm and unlonely. I feel lucky to have them in this lovely country.

Last but not least, I would like to express my appreciation to my dear family, my parents, parents in law, grandma in law, sisters, brothers in law and nephews. I would never achieve my study abroad without their understanding, encouragement, support and love. Thank my sisters and brothers in law for taking care of my parents in the past years and visiting them in all the important festivals. Finally and especially, I should not forget to thank my dearest wife. I appreciate that she travelled so frequently between China and the Netherlands and spend all of her holidays with me together. Her companionship relieved my pressure from work and my homesickness. It is a sweet memory that we got the marriage certificate in Chinese embassy in the Netherlands. Travelling around many beautiful European countries and wonderful cities in the Netherlands with her will be kept in my mind forever. Her love, trust and understanding made my PhD life bright and colorful!

Yuxiang Bai University of Groningen

Janurary 2016

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Yuxiang Bai was born on Septermber 27th 1986 in Yancheng, Jiangsu Province, China. After obtaining the B.Sc of Food Science and Engineering in Nanjing Tech University in September 2008, he got full scholarship and moved to the School of Food Science and Technology in Jiangnan University in Wuxi city to be a master student supervised by Prof. Zhengyu Jin, the vice-president of Jiangnan University and the director of National Key Laboratory of Food Science and Technology. Two years later, he directly got a PhD position in the same group, headed by Prof. Zhengyu Jin, investigating the physiochemical properties and application of the starch and starch-derived carbohydrates. In 2011, his research focus switched to the starch-acting enzymes. At the same year, he got the invitation from Prof. Lubbert Dijkhuizen, the head of Microbial Physiology group in University of Groningen and scientific director of Carbohydrate Competence Center and Protein Competence Center in the Netherlands. After obtaining a 2-year scholarship from Chinese Scholarship Council in early 2012, he arrived in the Netherlands on September 2012 and started his project of characterization of 4,6-α-glucanotransferase enzymes and their functional role in Lactobacillus reuteri. In 2014, he obtained the scholarship from University of Gronigen and became a regular PhD student supervised by Prof. Lubbert Dijkhuizen and Dr. R.M. van der Kaaij. Since 2016, Yuxiang will work in Jiangnan University.

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List of Publications: 1) Bai, Y. X.; van der Kaaij, R. M.; Leemhuis, H.; Pijning, T.; van Leeuwen, S.

S.; Jin, Z. Y.; Dijkhuizen, L., Biochemical characterization of Lactobacillus reuteri glycoside hydrolase family 70 GTFB type of 4,6-α-glucanotransferase enzymes that synthesize soluble dietary starch fibers. Appl Environmen Microbiol 2015. 81, 7223-7232.

2) Bai, Y. X.; van der Kaaij, R. M.; Woortman, A. J. J.; Jin, Z. Y.; Dijkhuizen, L., Characterization of the 4,6-α-glucanotransferase GTFB enzyme of Lactobacillus reuteri 121 isolated from inclusion bodies. BMC Biotechnol 2015, 15.

3) Bai, Y. X.; Wang, J. P.; Bashari, M.; Hu, X. T.; Feng, T.; Xu, X. M.; Jin, Z. Y.; Tian, Y. Q., A thermogravimetric analysis (TGA) method developed for estimating the stoichiometric ratio of solid-state α-cyclodextrin-based inclusion complexes. Thermochim Acta 2012, 541, 62-69.

4) Bai, Y. X.; Yu, B.; Xu, X. M.; Jin, Z. Y.; Tian, Y. Q.; Lu, L., Comparison of encapsulation properties of major garlic oil components by hydroxypropyl β-cyclodextrin. Eur Food Res Technol 2010, 231, 519-524.

5) Bai, Y. X.; Gangoiti, J.; Dijkstra B.; Dijkhuizen, L.; Pijning, T., Crystal structure of the 4,6- α-glucanotransferase: evidence that dietary changes triggered the evolution of glucansucrases from α-amylases. To be Submitted

6) Bai, Y. X.; Boger, M.; van der Kaaij, R. M.; Woortman, A.; Pijning, T.; van Bueren, A.; Dijkhuizen, L., Characterization of homo-exopolysaccharides produced by Lactobacillus reuteri 121 from maltodextrins and starch. To be Submitted

7) Bai, Y. X.; Dobruchowska, J.; van der Kaaij, R. M; Gerwig, G.; Dijkhuizen, L., Structural basis for the roles of starch and sucrose in homo-exopolysaccharide formation in Lactobacillus reuteri 35-5. To be Submitted

8) Tian, Y. Q.; Bai, Y. X.; Li, Y.; Xu, X. M.; Xie, Z. J.; Jin, Z. Y., Use of the resistance effect between retrograded starch and iodine for evaluating retrogradation properties of rice starch. Food Chemistry 2011, 125, 1291-1293.

9) Yu, B.; Bai, Y. X.; Wu, Y. C.; Jin, Z. Y., Preparation and Identification of 6(2)-α-Maltotriosyl-Maltotriose Using a Commercial Pullulanase. Int J Food Prop 2015, 18, 186-193.

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10) Yang, T. Y.; Bai, Y. X.; Wu, F. F.; Yang, N. J.; Zhang, Y. J.; Bashari, M.; Jin, Z. Y.; Xu, X. M., Combined effects of glucose oxidase, papain and xylanase on browning inhibition and characteristics of fresh whole wheat dough. J Cereal Sci 2014, 60, 249-254.

11) Xu, Y.; Zhou, X.; Bai, Y. X.; Wang, J. P.; Wu, C. S.; Xu, X. M.; Jin, Z. Y., Cycloamylose production from amylomaize by isoamylase and Thermus aquaticus 4-α-glucanotransferase. Carbohydrate Polymers 2014, 102, 66-73.

12) Zhang, H. X.; Tian, Y. Q.; Bai, Y. X.; Xu, X. M.; Jin, Z. Y., Structure and properties of maize starch processed with a combination of α-amylase and pullulanase. Int J Biol Macromol 2013, 52, 38-44.

13) Wang, J. P.; Ji, X. X.; Bai, Y. X.; Jin, Z. Y.; Xu, X. M.; Li, Y., Effects of fermentation conditions on the production of 4-α-glucanotransferase from recombinant Escherichia coli. Afr J Biotechnol 2011, 10, 17519-17531.

14) Bashari, M.; Nikoo, M.; Jin, Z. Y.; Bai, Y. X.; Xu, X. M.; Yang, N., Thermal and rheological properties of the supersaturated sucrose solution in the presence of different molecular weight fractions and concentrations of dextran. Eur Food Res Technol 2012, 234, 639-648.

15) Hu, X. T.; Wei, B. X.; Li, H. Y.; Wu, C. S.; Bai, Y. X.; Xu, X. M.; Jin, Z. Y.; Tian, Y. Q., Preparation of the β-cyclodextrin-vitamin C (β-CD-Vc) inclusion complex under high hydrostatic pressure (HHP). Carbohydrate Polymers 2012, 90, 1193-1196.

16) Jiang, Q. Q.; Xu, X. M.; Jin, Z. Y.; Tian, Y. Q.; Hu, X. T.; Bai, Y. X., Physico-chemical properties of rice starch gels: Effect of different heat treatments. J Food Eng 2011, 107, 353-357.

17) Hu, X. T.; Xu, X. M.; Jin, Z. Y.; Tian, Y. Q.; Bai, Y. X.; Xie, Z. J., Retrogradation properties of rice starch gelatinized by heat and high hydrostatic pressure (HHP). J Food Eng 2011, 106, 262-266.

18) 庄海宁, 冯涛, 金征宇, 谢正军, 徐学明, 高明, 柏玉香, 挤压加工参数对

重组米生产过程及产品膨胀度的影响, 农业工程学报, 2011 年 9 期. 19) 金征宇(导师), 柏玉香, 王金鹏, 环糊精葡萄糖基转移酶的筛选及其定向

改造, 食品与生物技术学报, 2012 年 2 期. 20) 于博, 柏玉香, 金征宇, 吴月纯, 汤尚文, 吴进菊, 麦芽三糖基-β-环糊精的

结构, 食品与发酵工业, 2013 年 7 期

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21) 金征宇, 王金鹏, 柏玉香, 环糊精制备过程的催化机制及其影响因素, 食品与生物技术学报, 2013年 1 期

22) 王田田, 周星, 柏玉香, 王金鹏, 谢正军, 金征宇, 超高压对 α-CGTase 产物专一性的影响, 食品与发酵工业, 2013 年 12 期

23) 岳艳, 王金鹏, 周星, 柏玉香, 王亚敏, 金征宇, 棕帘固定化环状芽孢杆菌

ATCC21783 制备 CGTase, 食品与发酵工业, 2013年 11 期

List of Patents (Chinese): 1) 金征宇,柏玉香,田耀旗,徐学明,谢正军,一种热分析判定固态 β-

环糊精包合物中主客体摩尔比的方法.CN 102095660 B 2) 金征宇,田耀旗,柏玉香,徐学明,谢正军,赵建伟,一种节能环保

的大蒜油环糊精包合物的制备方法. CN 101816744 A 3) 金征宇,翟秀超,王金鹏,柏玉香,焦爱权,李亦璇,徐学明,谢正

军,周星,一种具有缓释性能的食品抗菌包装材料及其制备方法 CN 102604343 A

4) 金征宇,田耀旗,朱艳巧,柏玉香,徐学明,谢正军,赵建伟,杨哪,

一种抑菌淀粉包装纸的制备方法. CN 101886356 B 5) 金征宇,李亦璇,王金鹏,焦爱权,徐学明,谢正军,田耀旗,杨哪,

柏玉香,翟秀超,一种环糊精抑制大豆脂肪氧合酶活性的方法.CN 102653753 A

Chapter of English Book:

Yuan, C; Bai, Y. X.; Jin, Z. Y., “Preparation and Analysis of Cyclodextrin Derivatives”, one chapter in “Cyclodextrin Chemistry” edited by Prof. Zhengyu Jin (World Scientific Publishing Co Pte Ltd)

Grants: 1) Fundamental Research Funds for the Central Universities (No.

JUDCF10051), 2010; 2) Graduate Student Innovation Project of Jiangsu Province (No.

CXZZ11_0486), 2011.

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Awards: 1) 7th “Challenge Cup” college students business plan competition in Jiangsu

province, Gold Award (Team leader, No. 1) 2) 8th National “Challenge Cup” college students business plan competition,

the 3rd Prize (Team leader, No. 1) 3) Scientific and Technical Academay Awards in Jiangsu Province, the 3rd

Prize (No. 11) 4) 1st college students business plan competition in Huishan district, Wuxi

city, Grand Prize (Team leader, No. 1) 5) 3rd Prize in the poster session of “11th Carbohydrate Bioengineering

Meeting-CBM 11” (Espoo, Finland) 6) Scholarship from Chinese Scholarship Council, 2012-2014 7) Scholarship from University of Groningen, 2014-2016

Article reviewing experience: 1) Food Hydrocolloid 2) Microbial Cell Factories 3) International Journal of Biological Macromolecules 4) 3 Biotech