Novel enzymes for functional carbohydrates production: From scientific research to application in health food industry 9813360208, 9789813360204

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Wanmeng Mu Wenli Zhang Qiuming Chen Editors

Novel enzymes for functional carbohydrates production From scientific research to application in health food industry

Novel enzymes for functional carbohydrates production

Wanmeng Mu • Wenli Zhang • Qiuming Chen Editors

Novel enzymes for functional carbohydrates production From scientific research to application in health food industry

Editors Wanmeng Mu Jiangnan University Wuxi, China

Wenli Zhang Jiangnan University Wuxi, China

Qiuming Chen Jiangnan University Wuxi, China

ISBN 978-981-33-6020-4 ISBN 978-981-33-6021-1 https://doi.org/10.1007/978-981-33-6021-1

(eBook)

© The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Singapore Pte Ltd. The registered company address is: 152 Beach Road, #21-01/04 Gateway East, Singapore 189721, Singapore

Preface

Carbohydrates are a key form of energy for most organisms. From the point of trendy diets, carbohydrates are divided into “good” or “bad” groups. The wellrefined carbohydrates with labels of high calories or high glycemic index are considered to be “bad.” The “good” carbohydrates are generally called the functional carbohydrates. The term “functional carbohydrate” is used not only to describe the edible carbohydrates but also to describe the ones with some physical functions other than the energy-supplying function, such as hyaluronic acid and curdlan with special solubilities and rheological properties. In the context of this book, only the carbohydrates with health-promoting effects especially prebiotic actions are referred to as functional carbohydrates. They have more nutritious value than traditional carbohydrates. Several enzymatic methods for the synthesis of such carbohydrates have been discovered and developed in recent decades, which provide a new range of application areas for these novel enzymes. The research on novel enzymes will remove many of the limitations currently restricting the manufacture of some functional carbohydrates. However, the lack of knowledge of scientific progress in enzyme technology might lead to slow progress of functional carbohydrate production. Therefore, to facilitate tracking the progress in the study of functional carbohydrate-related enzymes, the book will cover 13 chapters including the scientific progresses and recent applications of a selected number of emerging enzymes. The discussion on catalytic mechanism of the enzymes is also covered in this book. This book addresses the classification of functional carbohydrate-related enzymes and the overall development of food enzymes in Chap. 1. There are various types of functional carbohydrates with health-promoting effects including monosaccharides, oligosaccharides, and polysaccharides. The monosaccharides of such carbohydrates are collectively called as rare sugars. The common sugars that abundantly exist in nature are used as the starting materials to produce rare sugars. Chapters 2–5 describe the isomerases or epimerases involved in the production of rare sugars, such as D-allulose, D-mannose, D-tagatose, and D-allose. While the studies of the enzymes related to fructo-oligosaccharides (FOS) and galacto-oligosaccharides v

vi

Preface

(GOS) dominate the scientific literature in the field of enzymatic production of health-functional oligosaccharides, some enzymes also show promise for the emerging oligosaccharide production, which are introduced in Chaps. 6–9. In Chaps. 10 through 13, we summarize the new enzymatic technologies and applications in fructan- and glycan-related industries. We will give an overall perspective on the trends of enzymatic functional carbohydrate production in the last chapter. Wuxi, China Wuxi, China Wuxi, China

Wanmeng Mu Wenli Zhang Qiuming Chen

Contents

1

2

3

4

5

Development and Classification of Functional Carbohydrate Processing Enzymes in the Food Industry . . . . . . . . . . . . . . . . . . . . Wanmeng Mu and Qiuming Chen

1

Recent Advances in Ketose 3-Epimerase and Its Application for D-Allulose Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wenli Zhang, Jiajun Chen, and Wanmeng Mu

17

D-Mannose-Producing Isomerases and Epimerases: Properties, Comparisons, and Different Strategies . . . . . . . . . . . . . . . . . . . . . . Hao Wu and Qiuming Chen

43

L-Arabinose Isomerase: Sources, Biochemical Properties, and Its Use to Produce D-Tagatose . . . . . . . . . . . . . . . . . . . . . . . . . Hao Wu and Wei Xu

63

Various Enzymes for the Biotechnological Production of D-Allose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ziwei Chen, Wenli Zhang, and Wanmeng Mu

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6

Characteristics of Cellobiose 2-Epimerase and Its Application in Enzymatic Production of Lactulose and Epilactose . . . . . . . . . . . 105 Qiuming Chen, Yaqin Xiao, and Yanchang Wu

7

Enzymatic Production of Lactosucrose by Levansucrase, β-Fructofuranosidase, and β-Galactosidase . . . . . . . . . . . . . . . . . . . 125 Wei Xu, Wenli Zhang, and Hao Wu

8

Difructose Anhydrides-Producing Fructotransferase: Characteristics, Catalytic Mechanism, and Applications . . . . . . . . . 147 Mei Cheng, Yingying Zhu, and Wanmeng Mu

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Contents

9

Characteristics of Levansucrase and Its Application for the Preparation of Levan and Levan-Type Oligosaccharides . . . 175 Wei Xu, Wenli Zhang, and Wanmeng Mu

10

Inulosucrase, an Efficient Transfructosylation Tool for the Synthesis of Microbial Inulin . . . . . . . . . . . . . . . . . . . . . . . . 199 Dawei Ni, Wei Xu, and Wanmeng Mu

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Amylosucrase: A Versatile Sucrose-Utilizing Transglucosylase for Glycodiversification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 223 Yuqing Tian, Qiuming Chen, and Wenli Zhang

12

Glucansucrases Derived from Lactic Acid Bacteria to Synthesize Multitudinous α-Glucans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 251 Ziwei Chen, Dawei Ni, and Wanmeng Mu

13

Trends in Enzymology for Functional Carbohydrate Production . . 275 Qiuming Chen and Wanmeng Mu

Chapter 1

Development and Classification of Functional Carbohydrate Processing Enzymes in the Food Industry Wanmeng Mu and Qiuming Chen

1.1

Introduction of Modern Enzymology

As far back as 4000 years ago, the people in China began to use the enzymes in barley malt to make caramel and use fermentation to make wine and make vinegar and sauce. But at that time, people did not intentionally use enzymes. It was a natural behavior, only knowing the results but not knowing the principle. Moreover, the enzymes were mostly used to carry out hydrolysis reactions for debranching and improve solubility and clarity of food ingredients. In 1926, Sumner (1926) first obtained urease crystals that can decompose urea. This research has proved that it is a protein and has all the properties of protein. Afterward, researches on enzymes developed and became an independent discipline. With the development of physics and chemistry, enzymes became powerful technical methods for application. Due to the rapid development of the antibiotic industry stimulated by the Second World War, submerged fermentation was developed and introduced to the enzyme production industry, which can be regarded as a sign of the modernization of the enzyme industry. Enzymology research and application really entered a period of rapid development. Amylase and glucoamylase are first used industrially to produce glucose and fructose (Souza 2010; James and Lee 1997). High-fructose syrup is an important sweetener used in the food and beverage industry. Its sweetness is comparable to that

W. Mu (*) State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China e-mail: [email protected] Q. Chen State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_1

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of sucrose. It is currently the largest product produced using immobilized enzymes and one of the representatives of high-tech industries applying immobilized enzyme technology. Since then, lipase, lactase, glucose isomerase, glucose oxidase, aminoacylase, rennet, phosphodiesterase, etc. have also begun to be industrially produced. Especially in the late 1950s, the mass production of glucoamylase and the successful application of enzymatic production of glucose replaced the acid hydrolysis process that has been used for hundreds of years and greatly stimulate the development of the enzyme industry, thus becoming the first milestone in the modern enzyme history. In 1960, Denmark’s Novozymes implemented submerged fermentation to produce proteases on a large scale (Singh and Bajaj 2017). Since then several other enzymes have been developed for removing protein stains, which further stimulated the development of the enzyme industry. At the same time, since the 1950s, enzyme immobilization technology has been rapidly developed and gradually become mature (Robinson 2015). Especially in the early twentieth century, immobilized glucose isomerase was successfully used in the production of high-fructose syrup, with an annual output of more than 10 million tons. By creating a new way to make sugar from starch, it has effectively stimulated the development of the enzyme industry. The immobilization technology makes the biocatalyst possess completely different characteristics from its free state: the immobilized enzymes are easy to be separated from the product; the enzymes can be recovered or recycled; the stability of the enzymes is greatly improved; the reaction process can be strictly controlled. These characteristics greatly reduce the application cost of expensive enzymes, thus making it possible to be applied in large-scale industrial production. The immobilization technology is the second milestone in the enzyme industry. In 1973, Cohen et al. (1973) established DNA recombination technology, which opened the prelude to the era of genetic engineering. The gradual maturity and development of DNA recombination technology has revolutionized many fields of life science, becoming one of the fastest-growing disciplines since the twentieth century. The development of DNA recombination technology allows people to obtain many kinds of natural enzyme genes by cloning and express them efficiently in heterologous microbial receptors. Maltose amylase was first successfully produced by genetically modified microorganisms in the starch industry, and the heterologous high-efficiency expression technology of enzyme protein has been popularized as a conventional technical method in the enzyme industry. Since then, protein engineering technologies such as site-directed mutations of proteins were developed and implemented for tailoring their protein sequence and redesigning the functionalities of enzymes, creating the third milestone in the industrial production of enzyme. At present, there are more and more varieties of enzyme. The application scope of enzyme preparations is getting wider, and the quality of enzyme is getting better.

1 Development and Classification of Functional Carbohydrate Processing Enzymes in. . .

1.2

3

Current Commercial Carbohydrate-Related Enzymes

Because of the rapid development of biotechnology in the recent decades, heterogeneous production of enzymes from microbial sources became the method of choice for the production of industrial enzymes. The application of microbial enzymes usually shows lower production cost, better performance for genetic manipulation, and stronger physical and chemical resistances than their equivalents from plant or animal sources. Many microbial enzymes have been used variously in the industry for carbohydrate production (Table 1.1). The identification of new microbial sources is of high strategic interest. The amylases, glucoamylase, and glucose isomerase used to hydrolyze starch or convert glucose into fructose brought revolutionary changes to the sugar industry. Starch processing enzymes occupy the largest share of the market. Commercial enzymes are also widely used in the baking industry. Amylases can catalyze the breakdown of starch when baking. Glucose oxidase is used to improve stability of dough, prevent food browning, etc. Overall, most of the current commercial enzymes in food industry are used to hydrolyze biological polysaccharide or to improve the texture of foods.

1.3

Enzymatic Production of Functional Carbohydrate

With the increasing requirements for nutrition, in addition to the main function of nutrition supply, people have also paid great attention to the functionality of food. The latest trend in food industry is the development of functional foods such as prebiotics and low-calorie sweeteners. Carbohydrates and derivatives are important nutrients in the human diet. For humans 50–60% of total energy should come from carbohydrates according to dietary advice. They also serve as structural element and play important roles in metabolism processes. Carbohydrates are maligned from the mainstream perspective because conventional high carbohydrate diets increase the risk of chronic disease. The quality of carbohydrates is important to a healthy diet. Foods that are high in added carbohydrate should be avoided, and consumers should consider to improve the nutritional quality when choosing the types of sugar content in foods. With the continuous improvement of people’s living standards and the enhancement of health care awareness, people’s demand for functional sugar has greatly increased. Functional carbohydrates are carbohydrates that offer health benefits beyond basic nutrition. They have physiological effects such as low calorie, anti-caries, and intestinal flora regulation and are widely used in many fields such as food and pharmaceutical industries, which attract considerable interest over decades. Conventional carbohydrates such as glucose, fructose, sucrose, lactose, and rapidly digested starch are of high glycemic index (GI) and usually cause faster rise in blood glucose and insulin levels, which are harmful to human health.

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Table 1.1 List of the current commercial carbohydrate-related enzymes Enzyme activity Amylase (alpha)

Production organism Aspergillus niger, Aspergillus oryzae, Bacillus subtilis

Amylase (beta)

Bacillus flexus, barley, soybean, sweet potato Bacillus amyloliquefaciens, Bacillus subtilis, Geobacillus stearothermophilus Fusarium venenatum

Branching glycosyltransferase Cellobiose dehydrogenase Cellulase

Cyclodextrin glucanotransferase Dextranase Dextrin dextranase Glucanase (endo1,3(4)-beta) Galactosidase (alpha) Glucanase (endo1,3-beta) Glucanotransferase Glucoamylase

Glucose isomerase Glucose oxidase Glucosidase (alpha) Glucosidase (beta)

Glucosidase (exo-1.3-beta) Glucosyltransferase Hemicellulase

Aspergillus niger*, Bacillus amyloliquefaciens, Bacillus subtilis, Penicillium funiculosum, etc. Bacillus licheniformis, Bacillus macerans, Geobacillus stearothermophilus Chaetomium erraticum Chaetomium erraticum Aspergillus niger, Aspergillus oryzae, Bacillus subtilis, Cellulosimicrobium cellulans Aspergillus niger, Saccharomyces cerevisiae Streptomyces violaceoruber Bacillus amyloliquefaciens, Bacillus subtilis, Geobacillus pallidus Aspergillus niger, Rhizopus delemar, Rhizopus oryzae, Trichoderma reesei Streptomyces murinus, Streptomyces rubiginosus Aspergillus niger, Aspergillus oryzae, Penicillium chrysogenum Aspergillus niger, Trichoderma reesei Aspergillus niger, Penicillium decumbens, Penicillium multicolor, Trichoderma reesei Trichoderma harzianum Aspergillus niger Aspergillus niger, Bacillus amyloliquefaciens, Bacillus subtilis, Trichoderma reesei, Trichoderma longibrachiatum

Donor Bacillus sp., Thermoactinomyces sp. None

EC number 3.2.1.1

3.2.1.2

Rhodothermus sp.

2.4.1.18

Microdochium sp.

1.1.99.18

Bacillus sp., Streptomyces sp.

3.2.1.4

Thermoanaerobacter sp.

2.4.1.19

None None Thermoascus sp. Bacillus sp. Trichoderma sp. Guar plant

3.2.1.11 2.4.1.2 3.2.1.6

Streptomyces sp.

3.2.1.39

Thermus sp.

2.4.1.25

Aspergillus sp., Rhizomucor sp., Talaromyces sp. Streptomyces sp.

3.2.1.3, 3.2.1.39

Aspergillus sp., Penicillium sp. Aspergillus sp.

1.1.3.4

Trichoderma sp.

3.2.1.21

None

3.2.1.58

None Aspergillus sp.

2.4.1.24 –

3.2.1.22

5.3.1.5

3.2.1.20

(continued)

1 Development and Classification of Functional Carbohydrate Processing Enzymes in. . .

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Table 1.1 (continued) Enzyme activity Fructofuranosidase (beta) Galactosidase (beta) Maltodextrin α-Dglucosyltransferase Pullulanase Mannanase (endo1,4-beta) Rhamnosidase Xylanase

EC number 3.2.1.26

Production organism Saccharomyces cerevisiae

Donor None

Aspergillus oryzae, Bacillus circulans, Kluyveromyces lactis Streptomyces violaceoruber

Aspergillus sp., Kluyveromyces sp. Streptomyces sp.

3.2.1.23

Bacillus acidopullulyticus, Bacillus licheniformis, Bacillus subtilis Aspergillus niger, Trichoderma reesei, Trichoderma longibrachiatum Penicillium decumbens Aspergillus niger, Aspergillus oryzae, Bacillus subtilis, Bacillus licheniformis

Bacillus sp.

3.2.1.41

Trichoderma sp.

3.2.1.78

None Aspergillus sp., Talaromyces sp., Bacillus sp., etc.

3.2.1.40 3.2.1.8

5.4.99.15

However, these carbohydrates can serve as raw materials for production of functional carbohydrates. In addition to a small amount of natural extraction, functional carbohydrates are mainly manufactured by different types of reactions chemically or enzymatically. Because of the notorious difficulties in the chemical synthesis of carbohydrates, enzymatic methods will offer much more competitive processes for carbohydrates production in the future. Enzymes have strength not only in catalytic efficiency but also in chemoselectivity, regioselectivity, and stereoselectivity, which makes enzymatic reactions more precise, efficient, and reproducible. Therefore, the development of enzymes to produce or process functional carbohydrate becomes a research hotspot. Functional carbohydrates can be produced enzymatically through polymerization, isomerization, transglycosylation, or oxidation/reduction reactions. A number of novel enzymes have been discovered other than the current commercial carbohydrate-related enzymes.

1.4

Classification of the Functional Carbohydrate-Related Enzymes Based on Catalytic Mechanism

According to the induced fit model proposed by Daniel Koshland in 1958, the shape of enzyme and substrate will be slightly changed once the substrate is bound into the enzyme. During a reaction, the activation energy can be reduced to a certain extent by the enzyme (Fig. 1.1). Even a fairly modest reduction in the activation energy may result in a substantial increase in the reaction rate. The speed of enzymecatalyzed reaction is higher than that of inorganic catalysts 108 to 1020 times. Inorganic catalysts are usually used in bulk chemicals production, fine chemicals

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Fig. 1.1 Schematic diagram for describing the progress of reactions with and without an enzyme. The free energy is plotted as a function of a generalized reaction coordinate. E, S, and P represent enzyme, substrate, and product, respectively. The reaction pathway catalyzed by an enzyme goes through the transition states TS’ with the free energy of activation Ea0 , while the un-catalyzed reaction goes through the transition state TS with the free energy of activation Ea. The standard free energy changes of reaction (ΔG*) remain unchanged

production, and energy production processing, while enzymes are mostly used in food processing and pharmaceutical industry. The enzymatic catalysis is highly specific. Enzymes have high selectivity and specificity for the substrate or reaction type it catalyzes. Enzymes can only act on specific type of compounds through certain mechanism. In the early days, most of the names of enzymes were named by enzyme discoverers or based on the substrate catalyzed by the enzyme, the type of reaction, or the source of the enzyme. Enzymes catalyzing the hydrolysis of starch are called amylases. However, there are thousands of known enzymes. This nomenclature lacks systematic rules and cannot explain the nature of enzymatic reactions. To be able to accurately distinguish and correctly identify a certain enzyme, the Enzyme Commission (EC) proposed a systematic nomenclature according to the type of enzyme substrate and catalytic reaction and stipulated that each enzyme corresponds to a system name (Webb 1992). The system naming is more detailed and accurate to describe the type of enzyme catalytic reaction. Currently, enzymes are divided into seven categories according to their catalytic reaction types: oxidoreductases (EC 1, oxidation/reduction reactions), transferases (EC 2, transfer of a functional group), hydrolases (EC 3, hydrolysis), lyases (EC 4, non-hydrolytic removal of groups from substrates), isomerases (EC 5, intramolecular rearrangement), ligases (EC 6,

1 Development and Classification of Functional Carbohydrate Processing Enzymes in. . .

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synthesis of new bonds with the help of adenosine triphosphate), and translocases (EC 7, movement reaction across membranes). A limited number of lyases is developed for production of functional carbohydrates, e.g., alginate lyases (EC 4.2.2.3) catalyzing the formation of alginate oligosaccharides. Besides this, the hydrolysis, transfer, isomerization, and hydrogenation reactions present most of the reaction types catalyzed by the functional carbohydrate-related enzymes. Therefore, hydrolases, transferases, isomerases, epimerases, hydrogenase, and dehydrogenase are the primary focus of this chapter.

1.4.1

Hydrolases and Transferases Involved in Functional Carbohydrate Production

The first foundation for a family classification for carbohydrate-related enzymes was conducted for glycoside hydrolases (GHs). In 1989, Henrissat et al. classified 21 amino acid sequences of β-glycanase through hydrophobic cluster analysis and divided them into six cellulase families based on the similarity of amino acid sequence (Henrissat et al. 1989). Afterward, Henrissat et al. classified 291 GH sequences from different sources out of 301 sequences in the SWISS-PROT databases based on the similarity of amino acid sequence (Henrissat 1991). The classification system of GHs was established after a few updating studies by Henrissat and Bairoch (1996), Henrissat and Romeu (1995), Henrissat and Bairoch (1993)). Carbohydrate-active enzymes that can synthesize or decompose carbohydrates are collectively called CAZymes. With the accumulation of released sequential and structural information on CAZymes, the classification system was further extended to all CAZymes. In September 1998, this classification of CAZymes was available on the web, forming the CAZy database (http://www.cazy.org/) (Cantarel et al. 2009). GHs are widely used in biotechnological and biomedical applications, of which the genes are abundant and present in the majority of the CAZy database. GHs are important industrial enzymes used to breakdown biomass for production of biofuels or small sugars. This type of enzyme is currently one of the most commercialized biocatalysts. Some GHs are used to synthesize functional carbohydrates in the industry. For instance, maltooligosyltrehalose trehalohydrolase (EC 3.2.1.141) was used for production of trehalose from starch or maltodextrins with the help of maltooligosyltrehalose synthase (EC 5.4.99.15). It is worth pointing out that the GH family classified in CAZy database also includes the transferases utilizing the unmodified saccharides as the donors. This class of transferases is also referred to as transglycosidases, which are structurally and evolutionarily related to the GH family. Transglycosidases are an important type of enzymes for production of functional carbohydrates. Transglycosidases can cleave the glycosidic bond of their substrate and couple the subunit to a growing polymer chain (transglycosylation) or a water molecule (hydrolysis). Industrial

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interest in transglycosidases arises from their preference for transglycosylation over hydrolysis. Some representative promising hydrolases and transferases for functional carbohydrates production are briefly introduced as follows (Table 1.2).

1.4.1.1

Cyclodextrin Glucanotransferase

Cyclodextrin (CD) is a type of cyclic oligosaccharide, composed of multiple D-glucopyranose units connected by α-1,4-glycosidic bonds. CD includes α-cyclodextrin, β-cyclodextrin, and γ-cyclodextrin, which are composed of 6, 7, and 8 glucose units, respectively. CD molecules possess the structure of a truncated cone with a hydrophobic internal cavity, which can accommodate hydrophobic molecules or groups to form cyclodextrin inclusion complexes (Wang et al. 2016). The inclusion form can change the physical and chemical properties of the embedded objects, such as volatility, solubility, and chemical reaction performance (Li et al. 2019). CD has been widely used in many fields such as food, medicine, cosmetics, textile, and biotechnology. Cyclodextrin glycosyltransferase (EC 2.4.1.19, CGTase) belonging to GH family 13 can catalyze synthesis of cyclodextrin from starch or starch derivates. Since the natural strains capable of producing CGTase generally have relatively strict regulatory mechanisms, the yield of CD produced by the natural strains is relatively low. Therefore, genetic engineering methods are usually used to construct the recombinants containing CGTase genes with high expression efficiency (Chen et al. 2018). This enzymatic production of CD by the recombinant CGTase is mostly used in the present industry.

1.4.1.2

Sucrose-Utilizing Enzymes

Sucrose (a-D-glucopyranosyl-1,2-b-D-fructofuranoside) is one of the most widely consumed natural sweetener. Sucrase can not only catalyze the hydrolysis of sucrose to its subunits, fructose and glucose, but also they can catalyze synthesis of polysaccharides through transfer reaction. A large number of sucrases have been identified and divided into groups based on the types of synthesized polymers. Enzymes that synthesize the polysaccharides consisting of either glucose residues (glucans and amylose) or fructose residues (fructans) are classified into three general categories, fructansucrases (FSase), glucansucrases (GSase), and amylosucrase (ASase). Inulosucrase (EC 2.4.1.9, sucrose, 2,1-β-D-fructan 1-β-D-fructosyltransferase) and levansucrase (EC 2.4.1.10, sucrose, 2,6-β-D-fructan 6-β-D-fructosyltransferase) are two distinguished types of FS for synthesis of fructan or fructan-oligosaccharide (FOS) (Xu et al. 2019; Van Hijum et al. 2006). These enzymes are two key enzymes for sucrose metabolism (KEGG database, https://www.genome.jp/kegg/pathway). Inulin and levan are prebiotic polysaccharides showing versatile physicochemical properties and physiological functions (Ni et al. 2019). A variety of functional carbohydrates or glycoconjugates such as FOS can be produced from the abundant resource of sucrose utilizing FS enzymes. It is worth mentioning that difructose

Depolymerizing Transfering

Transglycosylation and hydrolysis

Reaction type Hydrolysis

Inulosucrase Levansucrase Dextransucrase Alternansucrase Reuteransucrase Amylosucrase β-Galactosidase β-Glucosidases Inulin fructotransferase β-1,3-Nacetylglucosaminyltransferase β-1,3-Galactosyltransferase β-1,4-Galactosyltransferase α-1,2-Fucosyltransferases α-1,3-Fucosyltransferases

Enzyme Maltooligosyltrehalose trehalohydrolase Cyclodextrin glucanotransferase Sucrose Sucrose Sucrose Sucrose Sucrose Sucrose Lactose Lactose Inulin Lactose LNT II LNT II Lactose Lactose

2.4.1.2.4.1.22 2.4.1.69 2.4.1.-

Substrate Starch, maltodextrins Starch

2.4.1.9 2.4.1.10 2.4.1.5 2.4.1.140 2.4.1.2.4.1.4 3.2.1.23 3.2.1.21 2.4.1.93 2.4.1.149

2.4.1.19

EC number 3.2.1.141

Table 1.2 Hydrolases and transferases for functional carbohydrate production

LNT LNnT 20 -FL 3-FL

α-Cyclodextrin, β-cyclodextrin, and γ-cyclodextrin Inulin and inulin-type oligosaccharide Levan and levan-type oligosaccharide Dextran-type polymer Alternansucrase-type polymer Reuteran-type polymer Amylose-type polymer Galacto-oligosaccharides Galacto-oligosaccharides Difructose dianhydrides LNT II

Product of functional carbohydrate Trehalose

1 Development and Classification of Functional Carbohydrate Processing Enzymes in. . . 9

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dianhydrides can be decomposed from inulin fructotransferase (IFTase, EC 2.4.1.93) (Cheng et al. 2017; Mu et al. 2019). Difructose dianhydrides are a family of cyclic fructose-type disaccharides which shows promising prebiotic effect and non-cariogenic property. Thus, difructose dianhydrides can be directly biosynthesized from sucrose by a continuous enzymatic production strategy utilizing both inulosucrase and IFTase (Yu et al. 2016). The nomenclature for GSase is comparably complicated, and some work has revealed that a limited number of amino acid substitutions may cause changes in glucosidic linkages in the synthesized products (Chen et al. 2019). GSase can catalyze the synthesis of various glucans varying from the type of glucosidic linkages, the length of the chains to the type of branching. A large number of GSase genes are presented in the CAZy database, and most of them are classified into GH 13, GH 70, and GH 77 families. In general, GSase enzymes, such as dextransucrase (EC 2.4.1.5), alternansucrase (EC 2.4.1.140), and reuteransucrase (EC 2.4.1.-), are named depending on the main glucosidic linkages present in their synthesized glucan (dextran with >50% α-1,6 linkage, alternan with alternating α-1,3/α-1,6 linkages, and reuteran with >50% α-1,4 linkage, respectively) (André et al. 2010). Different types of α-glucans with various sizes, branch degrees, and linkages can be synthesized by GSase, giving them distinct physicochemical properties (Van Hijum et al. 2006). ASase (EC 2.4.1.4) is another type of transglucosylase that can catalyze the synthesis of α-1,4-glucans, namely, amylose, and catalyze the transglucosylation of different exogenous acceptors (Tian et al. 2018). ASase belongs to GH 13 family in the classification system of CAZy database. It catalyzes the polymerization reaction forming only α-1,4 linkages (amylose-like polymer) using sucrose as the sole substrate. It has numerous applications in producing functional sweeteners (e.g., turanose and maltooligosaccharides), dietary fibers, carbohydrate-based carrier materials, and bioactive compounds (e.g., arbutin and flavonoids).

1.4.1.3

Lactose-Utilizing Transglycosidases

Lactose is an inexpensive disaccharide produced in large amounts in industry. It can be converted to value-added products such as lactulose, epilactose, and galactooligosaccharides (GOS) by isomerization, epimerization, or transgalactosylation reactions, respectively. Transglycosidases including β-galactosidase (EC 3.2.1.23), β-glucosidases (EC 3.2.1.21), and β-mannosidases (EC 3.2.1.25) are renowned for their hydrolytic activities, and they can also oligomerize GOS (Torres et al. 2010). GOSs are not digested by humans but selectively increase the beneficial microflora in the intestine, which can provide health benefits when utilized as food ingredients. Another bioactive derivative compound lactosucrose (O-β-D-galactopyranosyl(1,4)-O-α-D-glucopyranosyl-(1,2)-β-D-fructofuranoside) can be synthesized using both lactose and sucrose as substrates. The transgalactosylation can be catalyzed by β-fructofuranosidase, levansucrase, or β-galactosidase.

1 Development and Classification of Functional Carbohydrate Processing Enzymes in. . .

1.4.1.4

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Transferase Acting on Activated Sugars

In the CAZy classification system, only the transferase (EC, 2.4.1.) catalyzing the transfer of sugar moieties from activated donor to nucleophilic acceptor molecules are classified into glycosyl transferase (GT) family. Leloir (sugar-nucleotide-dependent) glycosyltransferases and non-Leloir glycosyltransferases are classified depending on the donor substrates. More than 100 glycosyltransferase families including about 400,000 enzyme modules are in the CAZy database currently (Nidetzky et al. 2018). GTs generally show good reaction selectivity and cover a broad range of donors and acceptors, leading to promising applications of the glycoside synthesis with a high precision. Therefore, biosynthesis of functional oligosaccharides such as human milk oligosaccharides (HMOs) can be performed directly from the sugar nucleotide. LNT II (Lacto-N-triose II), LNT (lacto-N-tetraose), LNnT (lacto-N-neotetraose), 20 -FL (20 -Fucosyllactose), and 3-FL (3-Fucosyllactose) might be the best exemplified by biosynthesis from activated sugars catalyzed by transferase. β-1,3-Nacetylglucosaminyltransferase (EC 2.4.1.149) can catalyze the transglycosylation of N-acetyl-D-glucosaminyl residue from UDP-GlcNAC to biosynthesize LNT II. LNT II can be used as a common precursor to produce LNT and LNnT by β-1,3-galactosyltransferase (EC 2.4.1.-) and β-1,4-galactosyltransferase (EC 2.4.1.22), respectively. LNT and LNnT are important core structures to those HMOs with more complex structures. Fucosyltransferase (EC 2.4.1.) can catalyze the transfucosylation of fucosyl residue to a series of saccharides including LNB, LacNAc units, or lactose to synthesize fucosylated HMOs and Lewis blood group antigens. In particular, α-1,2-fucosyltransferase (EC 2.4.1.69) catalyzes the transfer of the L-fucosyl moiety from guanosine diphosphate (GDP)-fucose to an acceptor lactose to form 20 -FL, which is the most abundant oligosaccharide in human milk. These fucosylated HMOs have been widely proved to have many physiological functions and health effects. In summary, the field of HMOs is developing very rapidly, with industrialization fueling biological research and novel findings in turn giving rise to new commercial opportunities.

1.4.2

Isomerases and Epimerases Involved in Functional Carbohydrate Production

Isomerases have been assigned EC number of EC 5. Epimerases (EC 5.1) are isomerase enzymes that catalyze the inversion of stereochemistry at the target chiral carbon in biomolecules. The isomerases that act on carbohydrates are usually used to describe the ones that catalyze structural isomerization reactions forming different sequences and/or different connectivity of bonds, such as the interconversion of an aldose to a ketose. Up to now, the application of isomerases is mainly present in

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Fig. 1.2 Izumoring strategy to link the process of all hexose production

sugar manufacturing. High-fructose corn syrup (HFCS) is a mixture of glucose and fructose, which can be used as a sweetener. Glucose isomerase (EC 5.3.1.5) is mainly used to catalyze the isomerization of D-glucose to D-fructose. It is the key enzyme for preparing HFCS by bioconversion in industry. Glucose isomerase, protease, and amylase are extremely important industrial enzymes in the current food industry. Epimerases and isomerase are attracting great attention because of their abilities to convert traditional carbohydrates to functional carbohydrates, especially rare sugars. Only D-glucose, D-galactose, D-mannose, D-fructose, D-xylose, L-arabinose, and D-ribose present in nature in large amounts. Rare sugars are the monosaccharides, and their derived polyols rarely exist in nature. All the hexose sugars including 16 aldohexoses, 8 ketohexoses, and 10 hexitols can theoretically be interconverted through a few enzymatic steps, of which the concept is called the Izumoring strategy (Izumori 2006). Thus, some low-calorie rare sugars possessing prebiotic properties can be synthesized by this strategy, leading to a huge application prospect in the health food industry. There is a strong evidence that D-tagatose and D-allulose have already been approved to be “generally recognized as safe” (GRAS) by the US Food and Drug Administration (Zhang et al. 2017, 2016). In Izumoring strategy (Fig. 1.2), ketose 3-epimerization and aldose-ketose isomerization are reversible enzymatic reactions catalyzed by families of ketose 3-epimerase and aldose isomerase, respectively. Ketose 3-epimerase catalyzes reversible C-3 epimerization between D-tagatose, D-allulose, D-fructose, and L-ribulose. There are three types of epimerases in the family of ketose 3-epimerase in terms of their substrate specificities: D-tagatose 3-epimerase, D-psicose 3-epimerase, and L-ribulose 3-epimerase. Aldose isomerase can catalyze reversible isomerization reactions between aldoses and ketoses. Aldose isomerase generally exhibits

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broad substrate specificities and allows for a wide range of application for rare sugar production. For instance, L-rhamnose isomerase catalyzes the isomerization between several aldoses and ketoses. It shows the potential to be applied in production of D-allose (Lin et al. 2011), L-lyxose (Granström et al. 2005), L-mannose (Park 2014), D-gulose (Bhuiyan et al. 1999), and L-galactose (Leang et al. 2004). L-arabinose isomerase can catalyze biosynthesis of D-tagatose from D-galactose (Xu et al. 2014; Tien-Kieu et al. 2018) and bioconversion of L-ribulose from L-arabinose (Liu et al. 2019). Relatively fewer studies have reported on the epimerases that act on inactivated disaccharides. Cellobiose 2-epimerase (CEase, EC 5.1.3.11) is currently the only reported epimerase that catalyzes epimerization reaction toward unsubstituted disaccharides. It can be used for production of prebiotic derivatives (epilactose and lactulose) of lactose. Further information on CEase will be discussed in Chapter 6.

1.4.3

Hydrogenase and Dehydrogenase Involved in Functional Carbohydrate Production

The nomenclature for the enzymes that catalyze the conversion of aldehydes and/or ketones to their corresponding alcohols is quite complex. Aldose reductases (ARase) can catalyze the NADPH-dependent conversion of glucose to sorbitol. It is the prototypical enzyme of the aldo-keto reductase (AKRase) superfamily (Petrash 2004). The members in the AKRase superfamily can metabolize a range of substrates. ARase and AKRase are well studied as a drug target in developing therapeutic strategies for several diseases such as diabetic complications, Alzheimer’s disease, and atherosclerosis (Barski et al. 2008). The nomenclature system for the AKRase superfamily is available at www.med.upenn.edu/akr/. Another class of enzymes catalyzing oxidation-reduction reactions between ketohexoses and the corresponding hexitols is referred to as polyol dehydrogenases (PDHs) (Lu et al. 2019; Izumori 2006). PDH plays a pivotal role in Izumoring strategy for catalyzing the reversible biotransformation between rare sugar and alcohol. Therefore, mannitol 2-dehydrogenase (EC 1.1.1.67), galactitol 2-dehydrogenase (EC 1.1.1.16), ribitol 2-dehydrogenase (EC 1.1.1.56), arabitol 2-dehydrogenase (EC 1.1.1.12), and xylitol 4-dehydrogenase (EC 1.1.1.14) belonging to the family of PDH were extensively studied to produce sugar alcohols. Mannitol 2-dehydrogenase can catalyze the biosynthesis of mannitol from D-fructose (Fig. 1.2). Galactitol 2-dehydrogenase can catalyze the production of L-/D-tagatose from galactitol. Allitol can be produced from D-fructose utilizing ribitol 2-dehydrogenase. Arabitol 2-dehydrogenase and xylitol 4-dehydrogenase are involved in the production of five-carbon sugar alcohols such as L-xylulose and D-xylulose.

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Conclusion

Functional carbohydrates have a wide range of applications in food, pharmaceuticals, and cosmetics sectors. Enzymes involved in the production of functional carbohydrates are attracting increasing attention recently. Many novel enzymes are extensively discovered and investigated, which provides a feasible enzymatic approach to produce functional carbohydrates industrially in the near future. Carbohydrates can be classified into three major groups based on their chemical structure, and so do the functional carbohydrates. They are rare sugars, prebiotic oligosaccharides, and functional polysaccharides. Selected novel enzymes are discussed based on the types of carbohydrates they act on in the following sections.

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Henrissat B, Claeyssens M, Tomme P, Lemesle L, Mornon J-P (1989) Cellulase families revealed by hydrophobic cluster analysi. Gene 81(1):83–95 Izumori K (2006) Izumoring: a strategy for bioproduction of all hexoses. J Biotechnol 124 (4):717–722. https://doi.org/10.1016/j.jbiotec.2006.04.016 James JA, Lee BH (1997) Glucoamylases: microbial sources, industrial applications and molecular biology—a review. J Food Biochem 21(6):1–52 Leang K, Maekawa K, Menavuvu BT, Morimoto K, Granström TB, Takada G, Izumori K (2004) A novel enzymatic approach to the massproduction of L-galactose from L-sorbose. J Biosci Bioeng 97(6):383–388 Li Y, Li C, Gu Z, Cheng L, Hong Y, Li Z (2019) Digestion properties of corn starch modified by α-D-glucan branching enzyme and cyclodextrin glycosyltransferase. Food Hydrocoll 89:534–541 Lin C-J, Tseng W-C, Fang T-Y (2011) Characterization of a thermophilic L-rhamnose isomerase from Caldicellulosiruptor saccharolyticus ATCC 43494. J Agric Food Chem 59(16):8702–8708 Liu X, Li Z, Chen Z, Wang N, Gao Y, Nakanishi H, Gao X-D (2019) Production of l-ribulose using an encapsulated l-arabinose isomerase in yeast spores. J Agric Food Chem 67(17):4868–4875 Lu F, Xu W, Zhang W, Guang C, Mu W (2019) Polyol dehydrogenases: intermediate role in the bioconversion of rare sugars and alcohols. Appl Microbiol Biotechnol 103(16):6473–6481. https://doi.org/10.1007/s00253-019-09980-z Mu W, Jiang B, Shuhuai Y, Zhu Y, Zhang T (2019) Highly efficient method for synthesizing difructose anhydride III. Google Patents Ni D, Xu W, Zhu Y, Zhang W, Zhang T, Guang C, Mu W (2019) Inulin and its enzymatic production by inulosucrase: characteristics, structural features, molecular modifications and applications. Biotechnol Adv 37(2):306–318. https://doi.org/10.1016/j.biotechadv.2019.01.002 Nidetzky B, Gutmann A, Zhong C (2018) Leloir glycosyltransferases as biocatalysts for chemical production. ACS Catal 8(7):6283–6300. https://doi.org/10.1021/acscatal.8b00710 Park C-S (2014) Characterization of a recombinant L-rhamnose isomerase from Bacillus subtilis and its application on production of L-lyxose and L-mannose. Biotechnol Bioprocess Eng 19 (1):18–25 Petrash JM (2004) All in the family: aldose reductase and closely related aldo-keto reductases. Cell Mol Life Sci 61(7-8):737–749. https://doi.org/10.1007/s00018-003-3402-3 Robinson PK (2015) Enzymes: principles and biotechnological applications. Essays Biochem 59:1–41. https://doi.org/10.1042/bse0590001 Singh S, Bajaj BK (2017) Potential application spectrum of microbial proteases for clean and green industrial production. Energy Ecol Environ 2(6):370–386. https://doi.org/10.1007/s40974-0170076-5 Souza PMD (2010) Application of microbial α-amylase in industry-A review. Braz J Microbiol 41 (4):850–861 Sumner JB (1926) The isolation and crystallization of the enzyme urease preliminary paper. J Biol Chem 69(2):435–441 Tian Y, Xu W, Zhang W, Zhang T, Guang C, Mu W (2018) Amylosucrase as a transglucosylation tool: From molecular features to bioengineering applications. Biotechnol Adv 36 (5):1540–1552. https://doi.org/10.1016/j.biotechadv.2018.06.010 Tien-Kieu N, Moon-Gi H, Pahn-Shick C, Byung-Hoo L, Sang-Ho Y, Martins LO (2018) Biochemical properties of L-arabinose isomerase from Clostridium hylemonae to produce D-tagatose as a functional sweetener. PLoS One 13(4):e0196099 Torres DPM, Gonçalves MDPF, Teixeira JA, Rodrigues LR (2010) Galacto-oligosaccharides: production, properties, applications, and significance as prebiotics. Compr Rev Food Sci Food Saf 9(5):438–454. https://doi.org/10.1111/j.1541-4337.2010.00119.x Van Hijum SAFT, Kralj S, Ozimek LK, Dijkhuizen L, Van Geel-Schutten IGH (2006) Structurefunction relationships of glucansucrase and fructansucrase enzymes from lactic acid bacteria. Microbiol Mol Biol Rev 70(1):157–176. https://doi.org/10.1128/mmbr.70.1.157-176.2006

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Wang L, Duan X, Wu J (2016) Enhancing the α-cyclodextrin specificity of cyclodextrin glycosyltransferase from paenibacillus macerans by mutagenesis masking subsite 7. Appl Environ Microbiol 82(8):2247–2255 Webb EC (1992) Enzyme nomenclature 1992. Recommendations of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology on the Nomenclature and Classification of Enzymes, vol 6. Academic, New York Xu Z, Li S, Feng X, Liang J, Xu H (2014) L-Arabinose isomerase and its use for biotechnological production of rare sugars. Appl Microbiol Biotechnol 98(21):8869–8878 Xu W, Ni D, Zhang W, Guang C, Zhang T, Mu W (2019) Recent advances in levansucrase and inulosucrase: evolution, characteristics, and application. Crit Rev Food Sci Nutr 59 (22):3630–3647. https://doi.org/10.1080/10408398.2018.1506421 Yu S, Zhu Y, Zhang T, Jiang B, Mu W (2016) Facile enzymatic production of difructose dianhydride III from sucrose. RSC Adv 6(105):103791–103794. https://doi.org/10.1039/ c6ra23352j Zhang W, Yu S, Zhang T, Jiang B, Mu W (2016) Recent advances in d-allulose: physiological functionalities, applications, and biological production. Trends Food Sci Technol 54:127–137. https://doi.org/10.1016/j.tifs.2016.06.004 Zhang W, Zhang T, Jiang B, Mu W (2017) Enzymatic approaches to rare sugar production. Biotechnol Adv 35(2):267–274. https://doi.org/10.1016/j.biotechadv.2017.01.004

Chapter 2

Recent Advances in Ketose 3-Epimerase and Its Application for D-Allulose Production Wenli Zhang, Jiajun Chen, and Wanmeng Mu

2.1

Rare Sugar

In the past few decades, the number of people dealing with obesity, diabetes, hyperlipidemia, and hypertension has grown dramatically throughout the world. The main reason leading to this case is the high-sugar and high-fat diets (Van Laar et al. 2020). With the continuous development of human’s living standards, the health awareness of the publics has gradually risen. As this result, the low-calorie rare sugars with special physiological functions have become popular, which are widely used as the sweeteners and flavor enhancers in food industries, such as healthy foods, infant formula, dairy products, baked goods, and beverage. According to the definition by the International Society of Rare Sugars (ISRS) in 2002, rare sugars are defined as the monosaccharides and their derivatives that occur in extremely small quantities in nature (Granström et al. 2004). Over 50 types of simple sugars exist in the natural world; only seven types occur abundantly, containing D-glucose, D-fructose, D-xylose, D-mannose, D-galactose, D-ribose, and L-arabinose. Despite its scarcity, rare sugars display great biological and functional potentials. For example, D-allulose displayed the weight loss, hypotensive, antitumor, anti-inflammatory, cryoprotective, and immunosuppressant effects (Mu et al. 2012); D-tagatose exhibited prebiotic activity, anticariogenic property,

W. Zhang (*) · J. Chen State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China e-mail: [email protected] W. Mu State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_2

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anti-glycemic activity (Xu et al. 2018); L-sugars were often used as the key precursors of pharmaceutical drug molecules (Chen et al. 2016); the sugar alcohols were often regarded as the low-calorie sweeteners with the effect of dental caries prevention, constipation treatment, and reducing insulin response (Rice et al. 2020). Usually, chemical synthesis of rare sugars needs multistep reactions and functional group protection-deprotection manipulation and brings chemical waste and by-product. By comparison, biotransformation method of rare sugars shows many advantages, for instance, mild reaction conditions, environmentally friendly, strong specificity, good sustainability, and high efficiency (Emmadi and Kulkarni 2014). After many years of research and exploration, Prof. Izumori from the International Institute of Rare Sugar Research and Education (Kagawa University, Japan) proposed the systematic biosynthesis methods of all rare sugars, named “Izumoring strategy.” Based on this strategy, all monosaccharides could be linked and cycled through ketose C-3 epimerization, aldose-ketose isomerization, and oxidationreduction catalyzed by ketose 3-epimerase (KEase), aldose isomerase (AIase), aldose reductase (ARase), and polyol dehydrogenase (PDH) (Granström et al. 2004; Izumori 2006). In the past three decades, most rare sugar productions were based on this method. In addition, some non-Izumoring methods have emerged, which is the good complement to “Izumoring strategy” (Zhang et al. 2017b), such as aldose C-2 epimerization, enzymatic aldol condensation, phosphorylationdephosphorylation cascade reaction, transglycosylation, ulosonic acid decarboxylation, and sucrose phosphorylase catalyzed by disaccharide biosynthesis.

2.2

Introduction of D-Allulose

D-Allulose, also named as D-psicose or D-ribo-2-hexulose, is a low-calorie rare hexose. It is the C-3 epimer of D-fructose with reducing activity, containing one ketone group (Fig. 2.1), and systematically named as (3R,4R,5R)-2(hydroxymethyl) oxane-2,3,4,5-tetrol by the International Union of Pure and Applied Chemistry (IUPAC), with the molecular formula of C6H12O6 (CAS No. 551-68-8) and molecular weight of 180.16 g/mol, respectively. Normally, D-allulose exists as a white odorless crystalline compound, crystallized solely as β-D-pyranose with the 1C (1C4 (D)) conformation. It is well soluble in water, with solubility of 291 g/100 g water (25  C) (Fukada et al. 2010). The melting points of Fig. 2.1 The structural formula of D-fructose and D-allulose

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D-allulose are measured as 109  C. D-Allulose is about 70% as sweet as sucrose and shows some cooling sensation and no bitterness. According to the regulation of US Food and Drug Administration (FDA), D-allulose provides dietary energy of 0.4 kcal/g (FDA-2019-D-0725). Owing to its high sweetness and ultralow energy, D-allulose has been widely recognized as an excellent low-calorie sweetener and the ideal substitute for sucrose. Now D-allulose is globally used as a bulk sweetener in beverage, bakery, candy, and dessert manufacturing. Allulose was first discovered through psicofuranine isolation; hence it was originally called “psicose” (Eble et al. 1959). In 2014, it was officially renamed as “D-allulose” by ISRS. D-Allulose is not naturally present in animals and occurred in small quantities in a few plants (Itea, wheat and figs) and certain bacteria. In addition, D-allulose also exists in some high-sugar foodstuffs, for example, maple syrup, commercial high-fructose corn syrup, processed molasses, steam-treated coffee, raisins, and long-heated fruit juice (Zhang et al. 2016b). The D-allulose contents in manufactured food were closely associated with the sugar concentrations, processing time, and heating temperature. D-Allulose is one of the most intensively studied monosaccharides among rare sugars, containing its healthy functions, biocatalysis, and industrial applications.

2.3

Physiological Functions of D-Allulose

Through the 14C-labeled method, the absorption, distribution, and metabolism of D-allulose were evaluated by rat experiments. It was found that, in the form of oral administration, about 70% D-allulose was absorbed in the small intestine and excreted out via urine without metabolism. A small amount of D-allulose was not absorbed and transferred to the large intestine. The unabsorbed D-allulose was partly fermented into short-chain fatty acids by cecal microbes, while the residual parts were directly eliminated via feces (Kimoto-Nira et al. 2017). In the form of intravenous administration, 50% D-allulose was excreted through urine within 1 h. The D-allulose was detected to accumulate in the liver, kidney, and urinary bladder, except for the brain (Tsukamoto et al. 2014). Although D-Allulose could not be metabolized and converted into energy in vivo, it shows a key role in the physiological activities. During the monosaccharide absorption, D-allulose shares the passive transporter GLUT5 (glucose transporter 5) with D-glucose during the its uptake, while in the efflux process, D-allulose and D-glucose, D-galactose, D-mannose, and D-fructose are effluxed via the same transporter GLUT2 (glucose transporter 2). Hence, D-allulose could partially suppress the uptake of D-glucose and D-fructose through competition effect (Hossain et al. 2015). As reported, D-allulose could stimulate the translocation of glucokinase (GK) and enhance hepatic glucose utilization and inhibit the increase in plasma glucose (Hossain et al. 2011). Additionally, D-allulose could greatly suppress the intestinal α-glucosidase and α-amylase activities, which delayed the digestion of sucrose and digestion and reduced the postprandial hyperglycemia (Matsuo and

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Izumori 2009). The antihyperlipidemic mechanisms of D-allulose have been well studied by a lot of researches; however, the antihyperlipidemic mechanism is not very clear. The possible reason might be that D-allulose influenced the activities of lipogenic (fatty acid synthase and glucose-6-phosphate dehydrogenase) and lipolytic enzymes and thus altered the triglyceride metabolism (Han et al. 2016). Besides, it was also reported that serum cholesterol metabolism could be altered by D-allulose through reducing serum PCSK9 levels (Kanasaki et al. 2019). The anti-obesity effect of D-allulose was not simply due to reducing food intake but also related to decreasing adipose tissue weight and fat mass, by enhanced fat oxidation and reduced carbohydrate oxidation (Ochiai et al. 2017). What’s more, D-allulose also exhibited many other physiological functions, for example, the anti-inflammatory effects by suppressing the serum proinflammatory cytokines (Han et al. 2020), the anti-oxidation effects by scavenging reactive oxygen species (ROS) (Sun et al. 2007), neuroprotective effects by scavenging reactive oxygen species and protecting the 6-hydroxydopamine-induced apoptosis in PC12 cells (Takata et al. 2005), and atherosclerosis treatment effects by inhibiting the expression of monocyte chemoattractant protein-1 (MCP-1) (Murao et al. 2007).

2.4

Application of D-Allulose

According to the toxicity rating chart, D-allulose is classified as the lowest toxicity category of “relatively harmless.” The acceptable daily consumption of D-allulose was 0.55 g/kg body weight, without undesirable diarrhea symptoms. For the regulation status, D-allulose has been approved as generally regarded as safe (GRAS) by FDA in the USA (GRN No. 828) and applied for approval as food ingredient in the European Union, Japan, and China. Recently, D-allulose has be excluded from the sugar count and can be used with a “no sugar added” claim according to the new regulation of FDA (FDA-2019-D-0725). Owing to its high sweetness, low calories, and good food safety, D-allulose could be used as a bulk sweetener and ideal sucrose substitute in beverage, bakery, candy, and dessert manufacture. Not only this, the addition of D-allulose could also improve the senses and quality during food processing. During caramelization and the Maillard reaction, the D-allulose content food was decreased as the reaction time, temperature, and pH increased, especially for pH (Oshima et al. 2014). Through the Maillard reactions, D-allulose could obviously improve the food properties of egg white protein and whey protein, including the gel strength, antioxidant activities, foaming properties, and emulsifying stability (Sun et al. 2008; Puangmanee et al. 2008). Besides, D-allulose exhibited the ability to promote the gelatinization and suppress retrogradation of rice flour (Ikeda et al. 2014). During the long-term frozen storage of chicken breast sausage, the presence of D-allulose could greatly reduce quality deteriorations, including viscosity, elasticity, and water-holding capacity (Hadipernata et al. 2016). In the heat-induced gelation of surimi, D-allulose was used as a gel improver to enhance the mechanical properties and water-holding capacity (Ogawa et al. 2017). In the

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yogurt fermentation, D-allulose supplement could suppress the acid production in some lactic acid bacteria and improve the taste, without changing the probiotic activity (Kim and Han 2019). D-Allulose also showed many non-food applications. Because of its excellent physiological functions, D-allulose displayed great potentials for the treatment of diabetes, obesity, hyperlipidemia, hypertension, and atherosclerotic diseases (Van Laar et al. 2020). Besides, D-allulose was widely used in the production of other rare sugars, such as D-allose (Seo et al. 2018), D-allitol (Hassanin et al. 2016), D-altrose (Menavuvu et al. 2006), D-tagatose, and D-talitol (Yoshihara et al. 2006). In addition, the chitosan biotransformation could be improved in Rhizopus oryzae by D-allulose supplement (Yoshihara et al. 2003). Moreover, through nanoimprint lithography, the plant-based materials derived from D-allulose were prepared, which was regarded as permanent, eco-friendly, water-repellent, light-transparent films for liquid crystal displays and optical devices (Takei and Hanabata 2015). What’s more, D-allulose could inhibit the motility and growth of parasite, suggesting that D-allulose was a potential anthelmintic (Sato et al. 2008).

2.5

Bioproduction of D-Allulose

Initially, D-allulose was obtained through chemical synthesis from 1,2,4,5-di-Oisopropylidene-β-D-fructopyranose (McDonald 1967). After that, D-allulose was obtained through chemical catalysis from D-fructose catalyzed by a molybdate ion catalyst (Bilik and Tihlarik 1973), or heat-boiling in the mixture of ethanol and trimethylamine (Doner 1979). However, similar with all chemical catalysis methods, chemical synthesis of D-allulose inevitably suffers from the complicated purification steps, organic reagent waste, as well as unwanted by-product formation. Therefore, the more moderate and efficient biocatalysis method becomes the research and production trends of D-allulose (Mu et al. 2012). The development and application of the simulation of the simulated moving bed (SMB) further has further promoted the bioproduction of D-allulose, with the complete separation purity of 99.36% (Long et al. 2009). Based on the Izumoring strategy proposed by Prof. Izumori, ketose 3-epimerase (EC 5.1.3.31) occupies the crucial position in the monosaccharide cycle and plays a key role in the epimerization at the C-3 position between the corresponding ketoses.

2.5.1

Microbial Source of KEase

Ketose 3-epimerase could catalyze the reversible epimerization between D-fructose and D-allulose. KEases are family enzymes, and according to the various optimum substrates, it could be classified as D-tagatose 3-epimerase (DTEase), D-allulose 3-epimerase (DAEase), D-fructose 3-epimerase (DFEase), and L-ribulose

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3-epimerase (LREase) (Zhang et al. 2016b). In 1994, the first KEase was isolated from Pseudomonas cichorii ST-24 and characterized as DTEase, with the optimum substrate specificity for D-tagatose (Itoh et al. 1994). Until 2006, the second KEase from Agrobacterium tumefaciens was reported and identified as DAEase, which could efficiently catalyze D-allulose from D-fructose (Kim et al. 2006a). In 2009, a novel KEase from Rhodobacter sphaeroides was discovered that showed best substrate specificity for D-fructose and was classified as DFEase (Zhang et al. 2009). Different with other KEases, the LREase from Mesorhizobium loti was also identified and displayed the best substrate specificity for a pentose, L-ribulose (Uechi et al. 2013b). In the past 10 years, the researches about KEases have developed rapidly, including heterologous expression, characterization, immobilization, crystal structure analysis, molecular modification, and food-grade expression. So far, more than 20 kinds of KEases from various microorganisms have been identified and characterized (Table 2.1). Although these KEases belong to one family enzyme, the multiple amino acid sequence homology varied from to 17.2% to 94.2% (data not shown), and the phylogenetic tree of ketose 3-epimerases was displayed in Fig. 2.2.

2.5.2

Enzyme Properties of KEase

The enzyme properties, including temperature, stability, pH, and metal ions, are the key factors in biocatalytic synthesis. The summary of the enzyme properties of the aforementioned KEases is shown in Table 2.2. As displayed in Table 2.2, the KEase showed the highest catalytic activity at the temperature ranges between 40 and 80  C. Among all these KEases, The DaeM DAEase (Patel et al. 2020) and T. maritima DAEase (Shin et al. 2017) displayed highest optimum temperature of 80  C, while R. sphaeroides DTEase showed the lowest optimum temperature of 40  C (Zhang et al. 2009). It is well-known that the higher operation temperature could accelerate the reaction speed, increase the solubility, lower the viscosity, reduce microbial contamination, and enhance the favorable equilibrium ratio of endothermic reactions, which is more preferred in the industrial production. However, if the rare sugar production is carried out at too high temperature under alkaline pH, the nonenzymatic browning reactions are easier to occur. Hence, an efficient and thermostable KEase with acidic optimum pH is needed. Additionally, the reaction temperature might influence the equilibrium ratio between D-allulose and D-fructose. For example, with the temperature increase from 30  C to 60  C, the equilibrium ratio between D-allulose and D-fructose catalyzed by C. bolteae DAEase was changed from 23:77 to 32:68 (Jia et al. 2014; Mu et al. 2011; Zhu et al. 2012), while, for Dorea sp. DAEase, the effect of temperature on the equilibrium ratio between D-allulose and D-fructose was very small, maintaining around 29:71 to 32:68 (Zhang et al. 2015). Thermostability is a key index of enzyme properties; however, most of the reported KEases displayed poor thermostability. Usually, they were relative thermostable under 50  C, while at higher temperature over 55  C, the catalytic activity

2 Recent Advances in Ketose 3-Epimerase and Its Application for D-Allulose. . .

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Table 2.1 KEases from different microorganisms Ketose 3-epimerase Agrobacterium sp.

Optimum substrate D-Allulose

GeneBank accession EGL65884.1

A. tumefaciens

D-Allulose

AAK88700.1

C. bolteae

D-Allulose

EDP19602.1

C. cellulolyticum

D-Allulose

ACL75304.1

C. scindens

D-Allulose

QBF73304.1

Clostridium sp.

D-Allulose

WP_014314767.1

DaeM Desmospora sp.

D-Allulose D-Allulose

QHD25651.1 WP_009711885.1

Dorea sp.

D-Allulose

CDD07088.1

F. plautii

D-Allulose

EHM40452.1

P. senegalensis

D-Allulose

WP_010270828.1

R. baltica

D-Allulose

WP_007330622.1

Ruminococcus sp. T. primitia

D-Allulose

ZP_04858451.1

D-Allulose

WP_010256447.1

A. globiformis

L-Ribulose

BAW27657.1

M. loti

L-Ribulose

BAB50456.1

S. aureus

L-Ribulose

SQA09501.1

T. maritima

L-Ribulose

NP_228226.1

C. fortuita

D-Tagatose

WP_061137998.1

Bacterium source Agrobacterium sp. ATCC 31749

Agrobacterium tumefaciens ATCC 33970 Clostridium bolteae ATCC BAA-613 Clostridium cellulolyticum H10 Clostridium scindens ATCC 35704 Clostridium sp. BNL1100 NRa Desmospora sp. 8437 Dorea sp. CAG317 Flavonifractor plautii ATCC 29863 Paenibacillus senegalensis Rhodopirellula baltica Ruminococcus sp. 5_1_39BFAA Treponema primitia ZAS-1 Arthrobacter globiformis M30 Mesorhizobium loti Staphylococcus aureus Thermotoga maritima MSB8 Caballeronia fortuita

Research institution National Taiwan University of Science and Technology Sejong University

Jiangnan University

Jiangnan University

Jiangnan University

Jiangnan University Panjab University Jiangnan University Jiangnan University Konkuk University

Chinese Academy of Sciences Tianjin University of Science and Technology Chinese Academy of Sciences Jiangnan University Kagawa University Kagawa University Tianjin University of Science and Technology Kyungpook National University Jiangnan University (continued)

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Table 2.1 (continued) Ketose 3-epimerase P. cichorii

a

Optimum substrate D-Tagatose

GeneBank accession BAA24429.1

Sinorhizobium sp.

D-Tagatose

WP_069063284.1

R. sphaeroides

D-Fructose

ACO59490.1

Bacterium source Pseudomonas cichorii ST-24 Sinorhizobium sp. Rhodobacter sphaeroides SK011

Research institution Kagawa University Tianjin University of Science and Technology Jiangnan University

NR not reported

drastically lost. Only DaeM DAEase, T. maritima LREase and M. loti LREase exhibited good thermostability. The half-life (t1/2) values of DaeM DAEase and M. loti LREase at 60 and 70  C were determined to be 165 h and 48 h at 60  C and 54 h and 2.3 h at 70  C, respectively (Patel et al. 2020; Uechi et al. 2013a). But the catalytic activity toward D-allulose of these KEases was ultralow (Table 2.2). At present, the poor thermostability of KEases became the critical limitation factor for the industry production of D-allulose. The top priority is to get good KEases mutants with high catalytic efficiency and strong thermal stability, through molecular modification or directed evolution. Generally, acidic reaction conditions were preferred in commercial bioproduction of rare sugars, because in the slightly acidic reaction conditions, the nonenzymatic browning could be significantly suppressed, and the unwanted by-product formation could be reduced. However, as shown in Table 2.2, most identified KEases exhibited optimum activity in weakly alkaline conditions (pH 7.5 to 9.0). C. bolteae DAEase (Jia et al. 2014), DaeM DAEase (Patel et al. 2020), F. plautii DAEase (Park et al. 2016), and T. maritima DAEase (Shin et al. 2017) displayed optimum pH in neutral conditions (pH 7.0). Only Dorea sp. DAEase showed optimum pH under weak acidic conditions (pH 6.0). Interestingly, the catalytic activity toward D-fructose of Dorea sp. DAEase was the highest among all the reported KEases with the specific activity of 803 U/mg, which exhibited great potentials in the industrial production of D-allulose (Zhang et al. 2015). It was found that metal ions played an important role in the C-3 epimerization by anchoring the bound of substrate. Many KEases were strictly Co2+ or Mn2+ metaldependent, which were completely inactivated in the absence of metal ions. The other KEases were not metal-dependent, and they could display catalytic capability without any metal ions. But the enzyme activity could be remarkably enhanced by the supplement of Co2+, Mn2+, or Mg2+. In contrast, the addition of Zn2+, Cu2+, as well as EDTA might remarkably suppress the enzyme e activity of the most KEases. Not only the different types of metal ions but also the concentration might affect the enzyme activity of the KEases. The effects of the metal ion concentration on some KEases were investigated, for example, C. scindens DAEase (Zhang et al. 2013a), Clostridium sp. DAEase (Mu et al. 2013), C. bolteae DAEase (Jia et al. 2014), and

2 Recent Advances in Ketose 3-Epimerase and Its Application for D-Allulose. . . 100 45

25

A. tumefaciens Agrobacterium sp. C. cellulolyticum

94 100 90

Clostridium. sp. C. scindens Dorea. sp.

100

Ruminococcus. sp. 99 C. bolteae

100

T. primitia

83

83

Desmospora. sp. 100

P. senegalensis

100

DaeM F. plautii P. cichorii

88 100

C. fortuita R. sphaeroides

100

Sinorhizobium sp.. M. loti

75

A. globiformis

100 100

S. aureus T. maritima R. baltica

99

0.1

Fig. 2.2 The phylogenetic tree of ketose 3-epimerases

T. primitia DAEase (Zhang et al. 2016c). Under ultralow concentrations, the catalytic activity could not be detectable; subsequently, similar with the first-order reaction, the catalytic activity increased linearly to the maximum with the metal ion concentration increase; finally the catalytic activity became stationary as the concentration increased. The overall effects of metal ion concentration were shown as an S-shaped trend (Zhang et al. 2013a). What’s more, the thermal and structure stability of KEases were also influenced by metal ions. It was reported that the supplement of metal ions during heating process could significantly increase the t1/2 values of C. cellulolyticum DAEase (Mu et al. 2011), Ruminococcus sp. DAEase (Zhu et al. 2012), and C. bolteae DAEase (Jia et al. 2014). Not only that, the addition of Mn2+ greatly improved the structure stability of C. scindens DAEase, with the

8.0

50

55

55

60

65

80

60

70

65

55

60

60

C. bolteae

C. cellulolyticum

C. scindens

Clostridium sp.

DaeM

Desmospora sp.

Dorea sp.

F. plautii

P. senegalensis

R. baltica

Ruminococcus sp.

7.5–8.0

8.0

8.0

7.0

6.0

7.5

7.0

8.0

7.5

8.0

7.0

Optimum pH 7.5–8.0

Optimum temperature ( C) 55–60

Ketose 3-epimerase Agrobacterium sp. A. tumefaciens Metal dependence Yes No Yes Yes Yes Yes Yes Yes Yes Yes No No No

Optimum metal ion Co2+ Mn2+ Co2+ Co2+ Mn2+ Co2+ Co2+ Co2+ Co2+ Co2+ Mn2+ Mn2+ Mn2+

Table 2.2 Comparison of enzyme properties of various KEases

1.14 252.1 803.5 NR

31:69 (80  C) 30:70 (60  C) 30:70 (70  C) 32:68 (65  C)

8.95

249.5

28:72 (65  C)

28:72 (60  C)

171

28:72 (50  C)

11.7

287

32:68 (55  C)

23:77 (60  C)

150.7

32:68 (60  C)

25.2

8.89

33:67 (40  C)

NR

Specific activity to D-fructose (U/mg) 253

Equilibrium ratio between D-allulose and D-fructose 30:70 (55  C)

References Tseng et al. (2018) Kim et al. (2006a) Jia et al. (2014) Mu et al. (2011) Zhang et al. (2013a) Mu et al. (2013) Patel et al. (2020) Zhang et al. (2013b) Zhang et al. (2015) Park et al. (2016) Yang et al. (2019) Mao et al. (2020) Zhu et al. (2012)

26 W. Zhang et al.

8.0

9.0

60

70

80

65

60

50

40

M. loti

S. aureus

T. maritima

C. fortuita

P. cichorii

Sinorhizobium sp. R. sphaeroides

7.5

7.5

7.0

8.0

8.0

7.5–8.0

70

A. globiformis

8.0

70

T. primitia

No

No

Co2+

Mn2+

Yes

Mn2+

No

No

Mg2+

Mn2+

No

Mn2+

No

No

Mg2+

None

Yes

Co2+

4 NR 0.1 270 NR NR NR

28.3:71.7 (60  C) 80:20 (70  C) 29.5:70.5 (65  C) 20:80 (30  C) 25.3:74.7 (50  C) 23:77 (40  C)

22.3

27:73 (70  C) NR

234.9

28:72 (70  C) Zhang et al. (2016c) Yoshihara et al. (2017) Uechi et al. (2013b) Zhu et al. (2019a) Shin et al. (2017) Li et al. (2019) Itoh et al. (1994) Zhu et al. (2019b) Zhang et al. (2009)

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urea mid-transition concentration (Cm) and melting temperature (Tm) increasing by 2.68 mM and 6.1  C, respectively (Zhang et al. 2013a).

2.5.3

Crystal Structure and Catalytic Mechanism

Until now, six crystal structures from A. tumefaciens DAEase (PDB No. 2HK0) (Kim et al. 2006b), P. cichorii DTEase (PDB No. 2QUL) (Yoshida et al. 2007), C. cellulolyticum DAEase (PDB No. 3VNI) (Chan et al. 2012), M. loti LREase (PDB No. 3VYL) (Uechi et al. 2013a), T. maritima (PDB No. 5H6H) (Shin et al. 2017), and A. globiformis (PDB No. 5ZFS) (Yoshida et al. 2018) have been already determined. As displayed in Fig. 2.3, A. tumefaciens DAEase, C. cellulolyticum DAEase, and M. loti LREase were determined to be dimers, while P. cichorii DTEase, T. maritima DTEase, and A. globiformis DTEase were homotetramers. By subunit superimposition of these six KEases, it could be found that the overall structures of the monomer were extremely similar. The subunit structure is a typically TIM-(β/α)8 barrel fold, and each monomer contains 12 α-helices and 8 β-strands as the main structural motif. Interestingly, the size of the α8 helix at the C-terminal tail of M. loti LREase was much bigger than other KEases, promoting to the further intermolecular interactions of different subunits. It was confirmed by the experiments that the long C-terminal tail greatly improved the thermostability of M. loti LREase (Uechi et al. 2013a, b). In 2006, according to the crystal structure analysis of A. tumefaciens DAEase and site-directed mutation experiments of two Glu residues (Glu150 and Glu244), Kim et al. proposed a possible catalytic mechanism. At first, one Glu residue coordinated

Fig. 2.3 (a) Superimposed diagram of monomers of the reported KEases. A. tumefaciens DAEase (red, 2HK0), C. cellulolyticum DAEase (yellow, 3VNI), M. loti LREase (blue, 3VYL), P. cichorii DTEase (gray, 2QUL), T. maritima LREase (purple, 5H6H), and A. globiformis LREase (cyan, 5ZFS), respectively. (b) Stereo views of the overall structures of reported apo-form KEases. (I) A. tumefaciens DAEase, (II) C. cellulolyticum DAEase, (III) M. loti LREase, (IV) P. cichorii DTEase, (V) T. maritima LREase, and (VI) A. globiformis LREase. Subunits Mol A, Mol B, Mol C, and Mol D were shown in gold, blue, orange, and magenta, respectively

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with Mn2+ removed a proton from C-3 of the substrate to form a cis-enediolate intermediate; subsequently the other Glu residue protonated at C-3 position in the opposite direction (Kim et al. 2006b). Later, the crystal structure of P. cichorii DTEase was solved, and the catalytic mechanism was further verified and improved. Yoshida et al. proposed the C3–O3 proton-exchange mechanism, which the Glu246 residue firstly removed a proton from C-3 of the substrate and produced a cis-enediolate intermediate with the plane structure of O2–C2–C3–O3; then the proton possibly transferred from Glu246 to Glu152 via O3; finally the Glu152 residue protonated at C-3 position, and the two Glu residues achieved the negatively ionized status (Yoshida et al. 2007). The crystal structure solved later further provided more evidence of the C3–O3 proton-exchange mechanism (Chan et al. 2012). Despite the low homology identity of amino acid sequences, the amino acids located at the catalytic sites and metal-binding sites were strictly conserved (Fig. 2.4). The metal ions in the active site were octahedrally coordinated to two water molecules and four residues (Glu, Asp, His, and Glu) that were strictly conserved among all the KEases. What’s more, three residues (Glu, His, and Arg) located at the substrate binding sites around O-1, O-2, and O-3 positions of the substrate were also conserved. However, these residues located at the substrate binding sites around the O-4, O-5, and O-6 positions were obviously different. This difference significantly influenced the substrate specificity and catalytic efficiency of KEases. The key residues located at active sites were compared and summarized in Table 2.3.

2.5.4

Molecular Modification of KEase

Because of its low catalytic efficiency and weak thermostability, the wild-type KEases usually could not meet the requirement of practical applications. In the past 10 years, the molecular modification of KEases through irrational, semirational, and rational design was developed rapidly. For the molecular modification of thermostability, in 2011, a double-site I33L/S213C mutation of A. tumefaciens KEase with great thermostability improvement was obtained through random mutagenesis. Compared with the wild-type enzyme, the optimum temperature increased by 7.5  C, t1/2 value increased by 29.9-fold at 50  C, and Tm value increased by 7.6  C (Choi et al. 2011). Based on the crystal structure of P. cichorii DTEase, a semi-rational surface engineering strategy for thermostability improvement of multimeric enzymes was proposed. The interface interactions of dimeric subunits were systematic optimized through iterative saturation mutagenesis, and the thermostable variant (Var8) of P. cichorii DTEase was constructed, with a T5020 value increase of 21  C (Bosshart et al. 2013). By N-terminal fusion of the SUMO homolog (Smt3) with A. tumefaciens DAEase, the optimum temperature (from 50 to 65  C) and thermostability were greatly improved. After heating at 21  C for

Fig. 2.4 Multiple sequence alignment of amino acids from six KEases with solved crystal structure. The GenBank accession numbers of KEases were as follows: A. tumefaciens DAEase (AAK88700.1), C. cellulolyticum DAEase (ACL75304.1), P. cichorii DTEase (BAA24429.1), A. globiformis LREase (BAW27657.1), M. loti LREase (BAB50456.1), and T. maritima LREase (NP_228226.1). The alignment was performed by the program ESPript

30 W. Zhang et al.

Residues involved in the O-4, O-5, and O-6 binding sites with D-fructose

Residues involved in the O-1, O-2, and O-3 binding sites with D-fructose

The active sites Residues involved in the metal coordinating sites

P. cichorii DTEase Glu152 Asp185 His211 Glu246 Glu158 His188 Arg217 Phe7 Trp15 Cys66 Leu108 Trp113 Phe248

Table 2.3 Summary of the residues located at active sites in KEases A. tumefaciens DAEase Glu150 Asp183 His209 Glu244 Glu156 His186 Arg215 Tyr6 Trp14 Gly65 Ala107 Trp112 Phe246

C. cellulolyticum DAEase Glu150 Asp183 His209 Glu244 Glu156 His186 Arg215 Tyr6 Trp14 Gly65 Ala107 Trp112 Phe246

A. globiformis LREase Glu146 Asp179 His205 Glu240 Glu152 His182 Arg211 His6 Phe14 Ser63 Val105 Met110 Phe242

M. loti LREase Glu147 Asp180 His206 Glu241 Glu153 His183 Arg212 His7 Leu252 Ser64 Ile106 His111 Phe243

T. maritima LREase Glu149 Asp182 His208 Glu243 Glu155 His185 Arg214 Val6 Phe14 Ala66 Leu113 Arg118 Leu245

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12 h, the SUMO-fusion A. tumefaciens DAEase retained its 45% residual activity, while the wild-type enzyme was rapidly inactive after 2 h in the same conditions. Besides, the catalytic efficiency was also improved, with a D-allulose space-time yields of 8.5 kg/L day (Patel et al. 2016). Besides, the hydrophobic substitutions of C. bolteae DAEase around the substrate pocket increased the structural rigidity of the substrate pocket and improved the thermostability. The double-site variant Y68I/ G109P of C. bolteae DAEase displayed the t1/2 value increase from 2.6 to 4.4 h and Tm value increase from 61.1 to 63.5  C (Zhang et al. 2016a). By site-directed mutagenesis of Dorea sp. DAEase at the interface regions, the F154Y/E191D/ I193F variant of Dorea sp. DAEase exhibited a significant improvement in thermostability, with t1/2 value increase by 5.4-fold at 50  C, and the Tm value increased by 17.54  C, respectively (Zhang et al. 2018). Based on the structure analysis of reported KEases variant and amino acid alignments, Zhu et al. redesigned the S. aureus LREase and got the more efficient and thermostable variants (Zhu et al. 2019a). To improve the robustness and thermostability of R. baltica DAEase, the residual substitution was carried out, which were located at the flexible regions. It provided an efficient and easy strategy for rational design and thermostability molecular modification of KEases (Mao et al. 2020). For the molecular modification of catalytic efficiency, using the thermostable Var8 variant of P. cichorii DTEase as the template, the iterative divergent evolution around the around the substrate binding site was carried out to improve the catalytic activity for D-fructose and L-sorbose. The eight-site mutant IDF8 and the six-site mutant ILS6 were obtained, which could efficiently convert D-fructose to D-allulose and L-sorbose to L-tagatose, respectively. Compared with those of Var8 template, the kcat of IDF8 mutant for D-fructose and the kcat of ILS6 mutant for L-sorbose showed 9-fold and 14-fold increases, respectively. (Bosshart et al. 2015). Using a laboratory-scale enzyme-membrane reactor (EMR), the maximum space-time yields of IDF10-3 P. cichorii DTEase variant for D-allulose production could reach 10.6 kg/L day, and ISL6 variant for L-tagatose production could reach 478 g/ L day (Bosshart et al. 2016). Through semi-rational design mutagenesis around the substrate binding pocket with O-4, O-5, and O-6 positions of substrate, the catalytic activity of C. bolteae DAEase variant was enhanced, with the Km value decreased by 17.9% and kcat/Km value increased by 1.2-fold (Zhang et al. 2016a). Furthermore, through the molecular modeling methods combined with protein structure networks, the reason for the enhanced connectivity between the substrate D-fructose and Y68I/G109P variant of C. bolteae DAEase was explained theoretically (Zhu et al. 2018).

2.5.5

Bioproduction of D-Allulose

Usually, the maximum equilibrium ratio between D-allulose and D-fructose catalyzed by KEases was nearly 30:70 (Table 2.2), except for P. cichorii DTEase (20:80 at 30  C) (Itoh et al. 1994), R. sphaeroides DFEase (23:77 at 40  C)

2 Recent Advances in Ketose 3-Epimerase and Its Application for D-Allulose. . .

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Fig. 2.5 The bioproduction strategy of D-allulose from D-glucose by coupling D-GIase and KEase

(Zhang et al. 2009), and T. maritima LREase (80:20 at 70  C) (Shin et al. 2017). It’s quite strange that the equilibrium ratio catalyzed by T. maritima LREase was so different from others, and the possibility might be that the catalytic activity of T. maritima LREase is ultralow, and the equilibrium ratio was measured by using D-allulose as the substrate. As mentioned above, the equilibrium ratio between D-allulose and D-fructose could be influenced by the reaction temperature. Similar with D-tagatose production, the conversion rate of D-allulose could be significantly increased by the addition of borate. The borate supplement could form the borate complex with monosaccharide, and the binding affinity between borate and D-allulose was much stronger than that between borate and D-fructose. In the presence of borate at a borate-to-fructose ratio of 0.6, the maximum conversion rate of D-allulose from D-fructose could reach 64% catalyzed by A. tumefaciens DAEase (Kim et al. 2008). Up to now, among all the reported KEases, Dorea sp. DAEase displayed the highest specific activity of 803 U/mg for D-fructose, in the acidic and elevated temperature conditions (pH 6.0 and 70  C) (Zhang et al. 2015). As we all know, D-glucose is a more common and cheaper monosaccharide. Based on the Izumoring strategy, D-fructose could be produced from D-glucose catalyzed by D-glucose isomerase (or D-xylose isomerase, GIase). As a result, D-allulose could be produced from D-glucose by coupling D-glucose isomerase and KEase, with D-fructose as the intermediate (Fig. 2.5). Men et al. constructed the one-step enzymatic process of D-allulose production, which inserted together the genes of Bacillus sp. GIase and Ruminococcus sp. DAEase in pCDFDuet-1 plasmid and co-expressed them in E. coli cells. The conversion rate of D-allulose from D-glucose could reach 16%, and the equilibrium ratio of D-glucose, D-fructose, and D-psicose was 3.0:2.7:1.0 (Men et al. 2014). Chen et al. built the co-expression system of the E. coli GIase and A. tumefaciens DAEase in E. coli that could efficiently produce D-allulose utilizing the sugarcane bagasse and microalgae with a yield of 1.42 and 1.69 g/L, respectively (Chen et al. 2017). By co-expression of A. cellulolyticus GIase and Dorea sp. DAEase, it could produce 204.3 g/L D-fructose and 89.1 g/L D-allulose from 500 g/L D-glucose, with the ratio of 6.5:7:3 between D-glucose, D-fructose, and D-allulose (Zhang et al. 2017a).

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Besides, Li et al. immobilized T. thermophilus GIase and I33L/S213C variant of A. tumefaciens DAEase on the wall of Saccharomyces cerevisiae spore spores, respectively. By this method, 12% of the D-glucose could be converted into D-allulose (Li et al. 2015). Interestingly, by the fusion of B. amyloliquefaciens exo-inulinase and A. tumefaciens DAEase through linker, D-allulose could be produced from inulin in one step, in which inulin was firstly converted to D-fructose and subsequently converted to D-allulose (Huang et al. 2020). In industrial production, enzyme immobilization is an efficient way for recycling enzyme and reducing the manufacturing cost. In 1995, Itoh et al. firstly immobilized P. cichorii DTEase on the Chltopearl beads of BCW 2503. By this method, it could produce 90 g D-allulose from 500 g D-fructose for 10 day at 45  C (Itoh et al. 1995). Subsequently, the mass production of D-allulose was improved by continuous bioreactor system using immobilized P. cichorii DTEase. The conversion rate of D-allulose could reach 25%, and the substrate D-fructose was removed by baker’s yeast treatment. Through the crystallization process with ethanol, 20 kg pure D-allulose could be produced after 60 day (Takeshita et al. 2000). In the presence of borate, 441 g/L D-allulose was from 700 g/L D-fructose in 2 h by A. tumefaciens DTEase immobilization on Duolite A568 beads. In the continuous packed-bed bioreactor reaction with borate, it could produce 325 g/L D-allulose from 500 g/L D-fructose at a dilution rate of 1.62/h in 236 h, with a D-allulose productivity of 527 g/L h (Lim et al. 2009). By N-terminal fusion of oleosin with C. cellulolyticum DAEase, the fusion protein was immobilized on Artificial Oil Bodies (AOBs), which retained over 50% of residual activity after five cycles (Tseng et al. 2014). He et al. fused the C. scindens DAEase gene at the C-terminal of CotZ protein and integrated into B. subtilis chromosome for immobilization on the surface of B. subtilis spores, which produce 85 g/L D-allulose from 500 g/L D-fructose after 12 h (He et al. 2016a). In recent years, with the development of nanomaterial technology, the immobilization of KEases on nanomaterials has been well studied. For example, A. tumefaciens DTEase was immobilized on graphene oxide, and it could be reused ten times efficiently with a D-allulose conversion rate of 40% (Dedania et al. 2017); besides, Patel et al. immobilized SUMO-fusion A. tumefaciens DAEase on iron oxide magnetic nanoparticles, which displayed four- to fivefold enhancement in its t1/2 values at 50–65  C and maintained over 90% residual activity after ten cycles (Patel et al. 2018); Zhu et al. immobilized C. cellulolyticum DAEase by creating hybrid organic-inorganic nanoflowers, which kept its 90% initial activity after six cycles (Zheng et al. 2018); the IDF10-3 variant of P. cichorii DTEase was fused to N-terminal of polyhydroxyalkanoate (PHA) synthase, and immobilized on the surface of PHA nano-beads, with 80% residual activity left after eight cycles (Ran et al. 2019); in addition, A. tumefaciens DAEase was also immobilized on titanium dioxide (TiO2) surface, which displayed bioconversion efficiency more than nine cycles of reusability (Dedania et al. 2020). The expression level in the native strains is very low and could not meet the requirement of industrial application. As this result, the heterologous expression of KEases in genetically engineered strains has been widely studied now. Because of its low cost, simple operation, fast growth speed, and high expression efficiency, in the

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earlier studies, E. coli was the most widely used host bacteria for the expression and properties of KEases. However, E. coli expression system usually produced endotoxins and led to food safety problems. The food-grade expression in Bacillus subtilis, Bacillus pumilus, Corynebacterium glutamicum, and Kluyveromyces marxianus has become the research hotspot. Based on the replicative plasmids with a D-alanine racemase gene marker, He et al. constructed a food-grade recombinant B. subtilis, which had no antibiotic resistance genes as well as no antibiotic supplement during fermentation (He et al. 2016b). To solve the instability problem of the plasmids, they constructed chromosome-integrated expression with tandem repeat genes in B. subtilis (He et al. 2016c). In addition, they expressed and displayed the C. scindens DAEase on the surface of B. subtilis spores (He et al. 2016a). Similarly, using alanine racemase-encoding gene as the selection marker, the food-grade expression cells of Ruminococcus sp. DAEase in B. subtilis was constructed and regulated by segmented fermentation (Fu et al. 2019). It was also reported that the C. cellulolyticum DAEase (Su et al. 2018), R. baltica DAEase (Zhang et al. 2020), C. bolteae DAEase, and Dorea sp. DAEase (Wei et al. 2020) were overexpressed in B. subtilis for D-allulose production. Besides, Li et al. overexpressed Ruminococcus sp. DAEase in B. pumilus, as well as D-allulose production, separation, and crystallization (Li et al. 2018). Moreover, A. tumefaciens DAEase was also overexpressed in K. marxianus, and it could efficiently produce D-allulose through cell regeneration and cyclic catalysis (Peizhou et al. 2018). In 2019, Yang et al. presented a tandem isoenzyme gene expression strategy in C. glutamicum, which could solve the problem of internal homologous recombination and express multiple DAEases in one cell (Yang et al. 2019).

2.6

Conclusion and Future Scope

Because of its low calories and high sweetness, as well as the excellent physiochemical properties, D-allulose has attracted more and more attention and displayed great potentials in food, pharmaceutical, cosmetic, and material fields. In the industrial bioproduction of D-allulose, the biocatalyst KEase plays the vital role. The poor thermostability, alkaline optimal pH, and low catalytic inefficiency of KEases have become the major restrictions for D-allulose. Therefore, the directed screening and molecular modification are needed. The task that is of top priority is to establish a high-throughput screening (HTS) method with simple operation, high sensitivity, and low cost. Moreover, the food-grade expression, high-density fermentation, and immobilization need further researching in the future.

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Sun Y, Hayakawa S, Ogawa M, Fukada K, Izumori K (2008) Influence of a rare sugar, D-PSicose, on the physicochemical and functional properties of an aerated food system containing egg albumen. J Agric Food Chem 56(12):4789–4796. https://doi.org/10.1021/jf800050d Takata MK, Yamaguchi F, Nakanose K, Watanabe Y, Hatano N, Tsukamoto I, Nagata M, Izumori K, Tokuda M (2005) Neuroprotective effect of D-psicose on 6-hydroxydopamineinduced apoptosis in rat pheochromocytoma (PC12) cells. J Biosci Bioeng 100(5):511–516 Takei S, Hanabata M (2015) Eco-friendly, water-repellent, light-transparent film derived from psicose using nanoimprint lithography. Mater Lett 143:197–200. https://doi.org/10.1016/j. matlet.2014.12.086 Takeshita K, Suga A, Takada G, Izumori K (2000) Mass production of D-psicose from D-fructose by a continuous bioreactor system using immobilized D-tagatose 3-epimerase. J Biosci Bioeng 90(4):453–455. https://doi.org/10.1263/jbb.90.453 Tseng C-W, Liao C-Y, Sun Y, Peng C-C, Tzen JTC, Guo R-T, Liu J-R (2014) Immobilization of Clostridium cellulolyticum D-psicose 3-epimerase on artificial oil bodies. J Agric Food Chem 62(28):6771–6776. https://doi.org/10.1021/jf502022w Tseng W-C, Chen C-N, Hsu C-T, Lee H-C, Fang H-Y, Wang M-J, Wu Y-H, Fang T-Y (2018) Characterization of a recombinant D-allulose 3-epimerase from Agrobacterium sp ATCC 31749 and identification of an important interfacial residue. Int J Biol Macromol 112:767–774. https:// doi.org/10.1016/j.ijbiomac.2018.02.036 Tsukamoto K, Hossain A, Yamaguchi F, Hirata Y, Dong Y, Kamitori K, Sui L, Nonaka M, Ueno M, Nishimoto K, Suda H, Morimoto K, Shimonishi T, Saito M, Song T, Konishi R, Tokuda M (2014) Intestinal absorption, organ distribution, and urinary excretion of the rare sugar D-psicose. Drug Des Dev Ther 8:1955–1964. https://doi.org/10.2147/dddt.S60247 Uechi K, Sakuraba H, Yoshihara A, Morimoto K, Takata G (2013a) Structural insight into L-ribulose 3-epimerase from Mesorhizobium loti. Acta Crystallogr Sect D 69:2330–2339. https://doi.org/10.1107/s0907444913021665 Uechi K, Takata G, Fukai Y, Yoshihara A, Morimoto K (2013b) Gene cloning and characterization of L-ribulose 3-epimerase from Mesorhizobium loti and its application to rare sugar production. Biosci Biotech Bioch 77(3):511–515. https://doi.org/10.1271/bbb.120745 Van Laar ADE, Grootaert C, Van Camp J (2020) Rare mono- and disaccharides as healthy alternative for traditional sugars and sweeteners? Crit Rev Food Sci Nutr. https://doi.org/10. 1080/10408398.2020.1743966 Wei H, Zhang R, Wang L, Li D, Hang F, Liu J (2020) Expression of d-psicose-3-epimerase from Clostridium bolteae and Dorea sp. and whole-cell production of d-psicose in Bacillus subtilis. Ann Microbiol 70(1). https://doi.org/10.1186/s13213-020-01548-x Xu W, Zhang W, Zhang T, Jiang B, Mu W (2018) L-arabinose isomerases: characteristics, modification, and application. Trends Food Sci Technol 78:25–33 Yang J, Tian C, Zhang T, Ren C, Zhu Y, Zeng Y, Men Y, Sun Y, Ma Y (2019) Development of food-grade expression system for d-allulose 3-epimerase preparation with tandem isoenzyme genes in Corynebacterium glutamicum and its application in conversion of cane molasses to D-allulose. Biotechnol Bioeng 116(4):745–756 Yoshida H, Yamada M, Nishitani T, Takada G, Izumori K, Karnitori S (2007) Crystal structures of D-tagatose 3-epimerase from Pseudomonas cichorii and its complexes with D-tagatose and D-fructose. J Mol Biol 374(2):443–453. https://doi.org/10.1016/j.jmb.2007.09.033 Yoshida H, Yoshihara A, Gullapalli PK, Ohtani K, Akimitsu K, Izumori K, Kamitori S (2018) X-ray structure of Arthrobacter globiformis M30 ketose 3-epimerase for the production of D-allulose from D-fructose. Acta Crystallogr 74:669–676. https://doi.org/10.1107/ s2053230x18011706 Yoshihara K, Shinohara Y, Hirotsu T, Izumori K (2003) Chitosan productivity enhancement in Rhizopus oryzae YPF-61A by D-psicose. J Biosci Bioeng 95(3):293–297. https://doi.org/10. 1016/s1389-1723(03)80032-4

2 Recent Advances in Ketose 3-Epimerase and Its Application for D-Allulose. . .

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Yoshihara K, Shinohara Y, Hirotsu T, Izumori K (2006) Bioconversion of D-psicose to D-tagatose and D-talitol by Mucoraceae fungi. J Biosci Bioeng 101(3):219–222. https://doi.org/10.1263/ jbb.101.219 Yoshihara A, Kozakai T, Shintani T, Matsutani R, Ohtani K, Iida T, Akimitsu K, Izumori K, Gullapai PK (2017) Purification and characterization of (D)-allulose 3-epimerase derived from Arthrobacter globiformis M30, a GRAS microorganism. J Biosci Bioeng 123(2):170–176. https://doi.org/10.1016/j.jbiosc.2016.09.004 Zhang LT, Mu WM, Jiang B, Zhang T (2009) Characterization of d-tagatose-3-epimerase from Rhodobacter sphaeroides that converts d-fructose into d-psicose. Biotechnol Lett 31 (6):857–862. https://doi.org/10.1007/s10529-009-9942-3 Zhang W, Fang D, Xing Q, Zhou L, Jiang B, Mu W (2013a) Characterization of a novel metaldependent D-psicose 3-epimerase from Clostridium scindens 35704. PLoS One 8(4):e0062987. https://doi.org/10.1371/journal.pone.0062987 Zhang W, Fang D, Zhang T, Zhou L, Jiang B, Mu W (2013b) Characterization of a metal-dependent D-Psicose 3-epimerase from a novel strain, Desmospora sp 8437. J Agric Food Chem 61 (47):11468–11476. https://doi.org/10.1021/jf4035817 Zhang W, Li H, Zhang T, Jiang B, Zhou L, Mu W (2015) Characterization of a D-psicose 3-epimerase from Dorea sp CAG317 with an acidic pH optimum and a high specific activity. J Mol Catal B 120:68–74. https://doi.org/10.1016/j.molcatb.2015.05.018 Zhang W, Jia M, Yu S, Zhang T, Zhou L, Jiang B, Mu W (2016a) Improving the thermostability and catalytic efficiency of the D-psicose 3-epimerase from Clostridium bolteae ATCC BAA-613 using site-directed mutagenesis. J Agric Food Chem 64(17):3386–3393 Zhang W, Yu S, Zhang T, Jiang B, Mu W (2016b) Recent advances in D-allulose: physiological functionalities, applications, and biological production. Trends Food Sci Technol 102:7283–7292 Zhang W, Zhang T, Jiang B, Mu W (2016c) Biochemical characterization of ad-psicose 3-epimerase from Treponema primitia ZAS-1 and its application on enzymatic production of d-psicose. J Sci Food Agric 96(1):49–56 Zhang W, Li H, Jiang B, Zhang T, Mu W (2017a) Production of d-allulose from D-glucose by Escherichia coli transformant cells co-expressing d-glucose isomerase and d-psicose 3-epimerase genes. J Sci Food Agric 97(10):3420–3426 Zhang W, Zhang T, Jiang B, Mu W (2017b) Enzymatic approaches to rare sugar production. Biotechnol Adv 35(2):267–274. https://doi.org/10.1016/j.biotechadv.2017.01.004 Zhang W, Zhang Y, Huang J, Chen Z, Zhang T, Guang C, Mu W (2018) Thermostability improvement of the d-allulose 3-epimerase from Dorea sp. CAG317 by site-directed mutagenesis at the interface regions. J Agric Food Chem 66(22):5593–5601 Zhang J, Xu C, Chen X, Ruan X, Zhang Y, Xu H, Guo Y, Xu J, Lv P, Wang Z (2020) Engineered Bacillus subtilis harbouring gene of D-tagatose 3-epimerase for the bioconversion of D-fructose into D-psicose through fermentation. Enzym Microb Technol 136:109531. https://doi.org/10. 1016/j.enzmictec.2020.109531 Zheng L, Sun Y, Wang J, Huang H, Geng X, Tong Y, Wang Z (2018) Preparation of a flower-like immobilized D-psicose 3-epimerase with enhanced catalytic performance. Catalysts 8(10). https://doi.org/10.3390/catal8100468 Zhu Y, Men Y, Bai W, Li X, Zhang L, Sun Y, Ma Y (2012) Overexpression of D-psicose 3-epimerase from Ruminococcus sp. in Escherichia coli and its potential application in D-psicose production. Biotechnol Lett 34(10):1901–1906. https://doi.org/10.1007/s10529012-0986-4 Zhu J, Li Y, Wang J, Yu Z, Liu Y, Tong Y, Han W (2018) Adaptive steered molecular dynamics combined with protein structure networks revealing the mechanism of Y68I/G109P mutations that enhance the catalytic activity of D-psicose 3-epimerase from Clostridium Bolteae. Front Chem 6:437. https://doi.org/10.3389/fchem.2018.00437 Zhu Z, Gao D, Li C, Chen Y, Zhu M, Liu X, Tanokura M, Qin H-M, Lu F (2019a) Redesign of a novel d-allulose 3-epimerase from Staphylococcus aureus for thermostability and efficient

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biocatalytic production of D-allulose. Microb Cell Factories 18:59. https://doi.org/10.1186/ s12934-019-1107-z Zhu Z, Li C, Liu X, Gao D, Wang X, Tanokura M, Qin H-M, Lu F (2019b) Biochemical characterization and biocatalytic application of a novel d-tagatose 3-epimerase from Sinorhizobium sp. RSC Adv 9(6):2919–2927. https://doi.org/10.1039/c8ra10029b

Chapter 3

D-Mannose-Producing Isomerases and Epimerases: Properties, Comparisons, and Different Strategies Hao Wu and Qiuming Chen

3.1

Introduction

Functional sugars are some of monosaccharides, oligosaccharides, polysaccharides, and sugar alcohols with special care and superior physiological functions on people’s health. Within the emerging development of modern food industry, many functional monosaccharides have been recognized by the public due to the attractive benefits and crucial roles they play in daily life. Different from common monosaccharides named D-glucose and D-fructose used mainly for providing energy, other functional sugars, such as D-mannose, D-tagatose, and D-allulose, possess many excellent properties of low calories and low sweetness, providing vast applications in food, medicinal, and beverage industries (Huang et al. 2018a). For example, D-allulose, generally regarded as safe (GRAS) by the US Food and Drug Administration (FDA), showed physiological functions of suppressing postprandial blood glucose (Hayashi et al. 2010) and anti-hyperlipidemic (Matsuo et al. 2001) and anti-inflammatory effects (Moller and Berger 2003). D-Tagatose also exhibited antidiabetic and obesity effects (Lu et al. 2008), flavor enhancement, and prebiotic and anti-biofilm properties (Bertelsen et al. 1999; Oh 2007). D-Mannose is another attractive functional sugar widely used in the food, medical, chemical, and feed industry (Wu et al. 2019). It has been reported that D-mannose can be used as a prebiotic (Korneeva et al. 2012), a facilitator for insulin secretion (Machicao et al. 1990), and a drug for phosphomannose isomerase deficiency (Lonlay and Seta 2009). Besides, D-mannose can be used to synthesize many different compounds which are very important in the medical industry as a raw material. For example, researchers have successfully achieved the chemical

H. Wu (*) · Q. Chen State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_3

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H. Wu and Q. Chen

synthesis of vitamins from D-mannose material (Chen et al. 2007). Also, some antitumor drugs (El-Nakkady et al. 2012) and immunostimulatory agents (Ranta et al. 2012) could be produced using D-mannose as a starting material. Due to these excellent benefits and applications, the production of D-mannose has received considerable attentions and interest by food scientists and companies. Currently, production of D-mannose was mainly through plant extraction, chemical synthesis, and biological production using enzymes. Some D-mannosecontaining plants are once the main source of D-mannose because D-mannose is the component of mannan, hemicellulose, and cellulose in the plant cell wall. The block of plant cell wall usually requires a specific condition of high temperature and acid hydrolysis or synergistic action of enzymes because of the high strength and high crystallinity in cell wall, which inevitably increased the production costs. Besides, the content of D-mannose was relatively low in some plants, resulting in the low yield of D-mannose. There is no exception that D-mannose can be chemically synthesized from D-glucose or D-mannitol by chemical scientists. However, the chemical production of D-mannose shows many shortcomings which are not friendly to the environment, especially in the sustainable development society advocated by people now. Chemical synthesis usually happened in specific equipment which can bear the strict conditions of high temperature, high pressure, and low acid environment and required the participation of inorganic catalysts. Furthermore, due to the weak selectivity of inorganic catalysts, many by-products are produced in this reaction system, which invisibly increased the separation costs and complexity during the subsequent process in food industry. Therefore, an alternative production method for D-mannose under mild reaction condition represents the future perspectives and trends when compared with aforementioned plant extraction and chemical synthesis. Biological production for D-mannose using enzyme approaches receives great attentions and interest because this reaction system is easy to operate and the enzyme reaction condition is mild and easy to achieve. The structure of D-mannose is very similar to D-fructose and D-glucose (Fig. 3.1). Detailedly, D-mannose is an epimer of glucose at the C-2 position and the aldose isomer of D-fructose. Theoretically, D-mannose can be produced from D-glucose or D-fructose raw materials using some epimerases and isomerases. Besides, mannitol-1-dehydrogenase (MDH, EC 1.1.1.255) can also catalyze the conversion of mannitol and D-mannose. These isomerases, epimerases, and dehydrogenases play crucial roles in the enzymatic production of D-mannose from other sugars and sugar alcohols. Currently, D-lyxose isomerase (LIase, EC 5.3.1.15) and D-mannose isomerase (MIase, EC 5.3.1.7) have been reported to have potential application in production of D-mannose from D-fructose by isomerization, whereas cellobiose 2-epimerase (CEase, EC 5.1.3.11) and D-mannose 2-epimerase (MEase, EC 5.3.1) are able to catalyze D-glucose to D-mannose by epimerization reaction. To efficiently produce D-mannose through biological methods, scientists tried their best to discover new D-mannose-producing enzymes with better catalytic efficiency and high thermostable properties. Different enzymes are introduced in this chapter in terms of their biochemical characteristics, kinetic parameters, and

3 D-Mannose-Producing Isomerases and Epimerases: Properties, Comparisons, and. . .

45

Fig. 3.1 The structure of D-glucose, D-mannose, D-fructose, and mannitol

reaction conditions. Besides, various strategies for D-mannose production including free enzymes, free cells, immobilization enzymes, and immobilization cells are compared and evaluated for its possibility. Furthermore, the physiological functions and benefits, determination methods, and applications of D-mannose are briefly discussed and outlined to enrich the knowledge about this functional sugar.

3.2 3.2.1

Overview of D-mannose Brief Description of D-mannose

The “mannose” term is derived from biblical term manna (Tonnesen 1983); D-mannose is also named D-mannoseis and D-mannopyranose. D-Mannose is a six-carbon hexose. Its chemical structure is displayed in Fig. 3.1. The molecular formula and weight are C6H12O6 and 180.16 g/mol, respectively. The density of D-mannose is 1.539 g/m3. Similar to D-glucose and D-fructose, D-mannose is a water-soluble compound. A maximum of 248 g D-mannose can be dissolved in 100 g water at 17
C. In a solution, 60% of D-mannose is in the crystallized form of α-D-pyranose, while the rest is β-D-pyranose form (Hu et al. 2016a). In terms of sweetness, D-mannose is 60% and 86% sweeter than sucrose and glucose, respectively. D-mannose has a low caloric value of 3.75 kcal/g (Pohl et al. 2012). D-mannose is an important intermediate metabolic product in the human body, which is found during the process of secreting some glycoproteins. It has been reported that a small amount of 2% D-mannose can be converted into glycogen for

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energy storage after being absorbed in the small intestine through free diffusion, thus not resulting in higher blood glucose level (Sharma et al. 2014).

3.2.2

Functions and Health Effects

Since its discovery, researchers are interested in this natural sugar for its functions and benefits. As D-mannose has hemiacetal hydroxyl group in its structure, it has a certain degree of reduction property and can increase the color when used in the food industry. In addition, in the food industry, D-mannose also has the functions of adjusting solubility, serving as a sweetener, increasing melting point, and improving flavor. As a dietary supplement influencing glyconutrient, D-mannose is essential for human health. It has been verified that D-mannose plays an important role in modulating systemic immunity, protecting against bacterial and viral infections, and promoting the normal synthesis of glycoproteins in the body (Sharma et al. 2014). Mannose-binding lectin can accurately recognize the mannose on the surface of pathogens that invade the human body, thereby initiating human immunity to resist pathogens and playing an important role in the human innate immune system (Turner 2003).

3.2.3

Determination Methods

As a concerned and attractive sugar, the detection of D-mannose becomes an important step before conducting qualitative and quantitative analysis. Currently, many different determination methods were developed to better assay the amount of D-mannose, including UV/visible spectrophotometry (Etchison and Freeze 1997), high-performance liquid chromatography (HPLC) (Jensen et al. 2010), gas-liquid chromatography (Kumar et al. 2014), potentiometric sensor (Grobler and Rechnitz 1980), and capillary electrophoresis electrospray mass spectrometry (Yeung et al. 1997). These methods have their own advantages and disadvantages. The UV/visible spectrophotometry can commonly be used for qualitative analysis without prior purification, whereas the tested results will be disturbed by many factors (Etchison and Freeze 1997). The HPLC method and gas-liquid chromatography are time-consuming and expensive. Samples need to be purified in advance before analyzing. Although the potentiometric sensor shows the advantage of a lower detection limit of 104 to 103 M, the narrow range of application is obvious (Grobler and Rechnitz 1980).

3 D-Mannose-Producing Isomerases and Epimerases: Properties, Comparisons, and. . .

3.3 3.3.1

47

Production of D-mannose Plant Extraction

In the past few decades, D-mannose was mainly extracted from some D-mannosecontaining plants, such as palm kernel, spend coffee grounds, Chinese jujube, orange peel, litchi pericarp, cranberries, apple flesh, mango, and so on (Hu et al. 2016a). The percentage of D-mannose varies in these plants. For example, the content of D-mannose in spent coffee grounds could reach at 21.2%, and this plant was an important material for extracting D-mannose (Mussatto et al. 2011). However, the content of D-mannose in cranberries (Johnson-White et al. 2006), apple flesh (Gheyas et al. 1997), and mango (Yashoda et al. 2007) was as low as 0.04–0.14%, 0.04–0.08%, and 0–0.03%, respectively. The various contents of D-mannose in different plants may be affected by the species, ripening stages, climate, and soil fertility. The extraction of D-mannose from palm kernel was conducted by Zhang and his workers under mild conditions (Zhang et al. 2009). This purification process involves acid hydrolysis, thermal hydrolysis, and enzyme hydrolysis. Firstly, the palm kernel should be treated by sulfuric acid at 100
C. Then, the hydrolyzed solution is further catalyzed by endo-β-mannanase. Thereafter, D-mannose was obtained through a silica gel column and ion exchange resin with a yield of 48.4% (Zhang et al. 2009). Additionally, microwave-assisted method combined with sulfuric acid hydrolysis was also employed to extract D-mannose from palm kernel (Fan et al. 2014). Under a condition of 148
C for 10 min and 31 s at a substrate-tosolvent ratio (w/v) of 1:69.69, a final yield of 92.11% D-mannose was obtained from the hydrolyzed solutions. Although D-mannose can be successfully gained from these plants, the extraction conditions are relatively harsh and unfriendly to the environment, which may not be completely accepted in the resource-saving society.

3.3.2

Chemical Synthesis

D-mannose can also be synthesized using inorganic catalysts from D-glucose material. Using 1% ammonium molybdate as catalyst, D-mannose could be produced from D-glucose through epimerization under the optimal condition of pH 3.0 and 150
C for 120 min and a conversion rate of 32%, D-mannose was achieved (Zhao et al. 2005). A similar conversion rate of D-mannose was also reported in another study, but the reaction condition was pH 2.0 and 98
C for 150 min (Zhang et al. 2017). If using ammonium molybdate and calcium oxide, a higher 44.8% conversion rate of D-mannose was obtained at 150
C, pH 3.0, for 80 min (Xu et al. 2014). It can be seen from the above that although D-mannose can be synthesized by chemical methods, the chemical synthesis has obvious disadvantages. The chemical reactions often happened by inorganic catalysts, such as ammonium molybdate, in a strong acid and high temperature environment. This process is full of high energy

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consumption and not economical. Furthermore, used inorganic catalysts and formed by-products should be removed from the reaction system, which increases the costs and difficulty of downstream separation.

3.3.3

Biological Synthesis

As mentioned above, biological production of D-mannose is an attractive and promising approach as this method happens in a mild condition which is easy to operate and achieve when compared with plant extraction and chemical synthesis methods. Currently, abilities of various isomerases and epimerases from different microbes to catalyze some monosaccharides including D-fructose and D-glucose to D-mannose through isomerase reaction and epimerase reaction were discovered. These D-mannose-producing isomerases and epimerases mainly included but are not limited to D-lyxose isomerase (D-LIase, EC 5.3.1.15), D-mannose isomerase (D-MIase, EC 5.3.1.7), cellobiose 2-epimerase (CEase, EC 5.1.3.11), and D-mannose 2-epimerase (D-MEase, EC 5.1.3.-). From previous literature surveys, it is found that among these D-mannose-producing enzymes, the most studied are mainly D-LIase and D-MIase.

3.3.3.1

D-LIase

Sources As shown in Table 3.1, approximately 12 D-LIases have been reported and biochemically characterized. These D-LIases are identified from Cohnella laevoribosii (Cho et al. 2007), Providencia stuartii (Kwon et al. 2010; Park et al. 2010a), Serratia proteamaculans (Park et al. 2010b), Escherichia coli (van Staalduinen et al. 2010), Bacillus licheniformis (Patel et al. 2011), Dictyoglomus turgidum (Choi et al. 2012), Thermosediminibacter oceani (Yu et al. 2016), Bacillus velezensis (Guo et al. 2019), Thermoflavimicrobium dichotomicum (Zhang et al. 2019), Caldanaerobius polysaccharolyticus (Wu et al. 2020a), Thermoprotei archaeon (Wu et al. 2020b), and Peptococcaceae bacterium (Huang et al. 2019).

Temperature and pH Optimal temperatures of these D-LIases are 40–85
C, showing a broad temperature range. This is because these enzymes are from different microbes characterized by mesophilic, thermophilic, and hyperthermophilic behaviors. Such results provide a reliable experience for researchers to find enzymes with suitable properties which are favored for in the industry. Actually, high reaction temperatures are beneficial to the production of sugars and sugar alcohols because higher temperature can increase the

EDU58657

BAJ07463.1

Q8X5Q7

AAU22106.1

YP_002352606.1

ADL08607.1

WP013351044.1

KLU40859.1

SFJ88669.1

WP_026486773.1 C. polysaccharolyticus 6.5

D-LIase

D-LIase

D-LIase

D-LIase

D-LIase

D-LIase

D-LIase

D-LIase

D-LIase

T. dichotomicum

P. bacterium

B. velezensis

T. oceani

D. turgidum

B. licheniformis

E. coli

S. proteamaculans

P. stuartii

7.5

7.5

6.5

6.5

7.5

7.5-8.0

7.5

7.5

7.5

6.5

D-LIase

C. laevoribosii

ABI93960.1

Optimum pH

D-LIase

Microorganism

GenBank accession

Enzymes

65

60

70

55

65

75

40–45

50

40

45

70

Optimum temperature (
C)

Mn2+ (1 mM) Co2+ (0.5 mM) Mn2+ (1 mM) Co2+ (0.1 mM) Co2+ (1 mM) Mn2+ (1 mM) Mn2+ (1 mM) 0.99

52.36

5.66

20.22

94.8

2.99

41.6

12.7

16170

2640

38.82

80.8

33.47

55.8

32.8

13

26

19.8

32.2

22

0.03

0.65

0.17

0.36

2.9

0.23

1.6

0.64

502

116

1.4

21

22

21

20

22

22

19.5

19.3

27

22

21

34

Mn2+ (1 mM) Mn2+ (1 mM) Mn2+ (1 mM) Mn2+ (1 mM) 47.6

Molecular Weight (kDa) by SDS-PAGE

D-Mannose as substrate kcat/Km (s1 Km Optimum kcat (mM) mM1) metal ions (s1)

Table 3.1 The biochemical characteristics of different D-mannose-producing enzymes

Dimer

Dimer

Dimer

NR

Dimer

Dimer

NR

Dimer

Dimer

Dimer

Dimer

Polymer

(continued)

Cho et al. (2007) Kwon et al. (2010) Park et al. (2010b) van Staalduinen et al. (2010) Patel et al. (2011) Choi et al. (2012) Yu et al. (2016) Guo et al. (2019) Huang et al. (2019) Zhang et al. (2019) Wu et al. (2020a)

Reference

3 D-Mannose-Producing Isomerases and Epimerases: Properties, Comparisons, and. . . 49

NP_418316.4

AB761401

AJH12524.1

NR

NC_009437

D-MIase

D-MIase

D-MIase

D-MIase

CEase

C. saccharolyticus

M. mediterranea

E. coli

T. fusca

E. coli

A. radiobacter

8.1

7.8

7.5

7.3

7.0

8.0

7.0

8.0

8–8.5

6.5

Optimum pH

37

37

50

30

45

60

47

60

60

80–85

Optimum temperature (
C)

NR

NR

None

NR

None

None

None

NR

3.88

4.77

0.74

329

40.9

788

25.3

NR

NR

66.4

73.5

51.8

16.7

86.4

115

108

NR

NR

0.06

0.06

0.01

19.7

0.47

6.85

0.23

NR

NR

0.42

47

45

47

NR

45

41

NR

44

NR

21

111.5

Ni2+ (0.5 mM) NR 46.8

Molecular Weight (kDa) by SDS-PAGE

D-Mannose as substrate kcat/Km (s1 Km Optimum kcat (mM) mM1) metal ions (s1) Reference

Wu et al. (2020b) NR Takasaki et al. (1993) Dimer Hirose et al. (2001) NR Itoh et al. (2008) Dimer Kasumi et al. (2014) Hexamer Hu et al. (2016b) NR Saburi et al. (2018) Monomer Park et al. (2011) Dimer Saburi et al. (2019) Dimer Saburi et al. (2019)

Dimer

Polymer

D-LIase D-lyxose isomerase, D-MIase D-mannose isomerase, CEase cellobiose 2-epimerase, MEase D-mannose 2-epimerase C. laevoribosii, Cohnella laevoribosii; P. stuartii, Providencia stuartii; S. proteamaculans, Serratia proteamaculans; T. fusca, Thermobifida fusca; M. mediterranea, Marinomonas mediterranea; C. saccharolyticus; Caldicellulosiruptor saccharolyticus; R. slithyformis, Runella slithyformis; D. fermentans, Dyadobacter fermentans; T. dichotomicum, Thermoflavimicrobium dichotomicum NR not reported, None not dependent

D. fermentans

NR

D-MIase

Pseudomonas sp.

D-MEase ACT96842.1

NR

D-MIase

T. archaeon

R. slithyformis

RLE72808.1

D-LIase

Microorganism

D-MEase AEI50834.1

GenBank accession

Enzymes

Table 3.1 (continued)

50 H. Wu and Q. Chen

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51

reaction rate, improve the solubility of products and substrates, and reduce the microbial contamination. However, excessively high temperature reactions may sometimes cause Maillard reaction, which can cause color browning and generate some undesirable by-products (Shen and Wu 2004). These color browning and unwanted by-products will increase the separation and decolorizing costs during the subsequent preparation of high-purity sugars. Interestingly, the browning degree in the acidic environment will be reduced because the produced by-products formed by carbonyl ammonia reaction caused by protein and reducing sugar interaction will be hydrolyzed (Wu et al. 2019; Shen and Wu 2004). Therefore, enzymes with optimal acid pH properties possess superior advantages for the production of sugar under higher temperature environment in the industry. As displayed in Table 3.1, the currently reported D-LIases showed a narrow optimal pH range of 6.5–8.0. Although some of D-LIases exhibited low-acid (pH 6.5) properties, such as D-LIase from T. archaeon (Wu et al. 2020b), C. polysaccharolyticus (Wu et al. 2020a), B. velezensis (Guo et al. 2019), T. oceani (Yu et al. 2016), and C. laevoribosii (Cho et al. 2007), the weakly acidic environment is not enough to inhibit the non-enzymatic browning reaction completely. Thus, searching for novel recombinant or wild-type D-LIases with eosinophilic behavior or conducting molecular modification of current D-LIases is a crucial direction in D-mannose production field.

Information About Crystal Structure At present, the crystal structure of a total of two D-LIases have been disclosed in the PDB database with numbers of 3MPB and 2Y0O, derived from E. coli and B. subtilis, respectively (Fig. 3.2). The crystal structure insights revealed that metal ions play an important part in the catalysis of D-fructose to D-mannose by D-LIase. The metal coordination sites are in the hydrophobic pocket which is formed by β-barrel. In E. coli D-LIase, four completely conserved residues named H103, H105, H171, and E110 are coordinated metal ions, whereas in B. subtilis D-LIase, the four conserved residues are changed to H69, H71, H137, and E82. It may be concluded that one glutamic acid residue and three histidines are responsible for the binding of metal ions among all the D-LIases. Furthermore, the experiment of optimal metal

Fig. 3.2 The crystal structure of D-LIase from (a) E. coli (PDB: 3MPB) and (b) B. subtilis (PDB: 2Y0O)

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ions determination also gave a direct evidence about its importance to D-LIase’ activity.

Metal Ions Dependence Seen from Table 3.1, all the reported D-LIases show metal ion-dependent properties. Among all the tested metal ions (Mg2+, Mn2+, Cu2+, Co2+, Ba2+, Zn2+, Ni2+, Ca2+), Mn2+, Co2+, and Ni2+ are verified as optimal metal ion for different sources of D-LIase. The optimal metal ion concentration ranges from 0.1 to 1.0 mM. For example, the activity of D-LIase from B. velezensis can increase 4.1-fold in the presence of 0.1 mM Co2+ than the group without adding any metal ion (Guo et al. 2019). With the presence of 1.0 mM Mn2+, the activity of C. polysaccharolyticus D-LIase is 7.5-fold increased than the group lacking the metal ion. For T. archaeon D-LIase, 0.5 mM Ni2+ enhances the enzyme activity greatly by 25-fold compared with the control group (Wu et al. 2020b). Although the subunit molecular weight of currently reported D-LIases determined by SDS-PAGE method is between 19.3 kDa and 27 kDa, they all exhibit a dimer structure. Using D-mannose as substrate, the kinetic parameters (Km, kcat, and kcat/Km) of these D-LIases are various. The highest kcat and kcat/Km values were 16,170 s1 and 502 mM1 s1, respectively, which were found in S. proteamaculans D-LIase (Table 3.1) (Park et al. 2010b). The lowest Km value was 13 mM, which was reported by D. turgidum D-LIase (Choi et al. 2012).

3.3.3.2

D-MIase

Sources Similar to the function of D-LIase, D-MIase can also catalyze the conversion reaction between D-fructose and D-mannose. To date, several D-MIases have been reported with the ability to produce D-mannose from D-fructose. These D-MIases are from Pseudomonas saccharophila (Palleroni and Doudoroff 1956), Xanthomonas rubrilineans (Takasaki and Takano 1964), Streptomyces aerocolorigenes (Takasaki 1967), Mycobacterium smegmatis (Hey-Ferguson and Elbein 1970), Escherichia coli (Stevens et al. 1981; Itoh et al. 2008; Hu et al. 2016b), Pseudomonas cepacia (Allenza et al. 1990), Pseudomonas sp. (Takasaki et al. 1993), Agrobacterium radiobacter (Hirose et al. 2001, 2003), Thermobifida fusca (Kasumi et al. 2014), and Marinomonas mediterranea (Saburi et al. 2018).

Biochemical Parameters The optimal temperatures of currently reported D-MIases are between 30 and 60
C, and the temperature range is lower than the D-LIases (Table 3.1). On the other hand, the optimal pH value of D-MIases ranges from 7.0 to 8.5, suggesting that all the

3 D-Mannose-Producing Isomerases and Epimerases: Properties, Comparisons, and. . .

53

reported D-MIases exhibit weak alkaline characteristic. As mentioned above, weakacid environment will be favored for the production of functional sugar because this environment can reduce the non-enzymatic browning effect. Therefore, searching for new D-MIases with weak-acid property or performing the molecular modification of presented D-MIase is a requirement in the future to meet industrial demands. For some D-MIases, it is not clear whether they are metal ion-dependent or not, such as Pseudomonas sp. D-MIase (Takasaki et al. 1993), A. radiobacter D-MIase (Hirose et al. 2001), and M. mediterranea D-MIase (Saburi et al. 2018); other identified D-MIases show metal ion-independent properties, which is greatly different from D-LIases (Table 3.1). In fact, metal ion-independent enzymes have certain benefits in the functional sugar industry, because metal ions must be removed from the reaction system during the downstream separation of functional sugar as the metal ions have the toxic effect on people’s health. Thus, this system without adding metal ions can reduce the unnecessary separation costs to some extent. Similar to D-LIase, the reported D-MIase is also a multi-subunit protein. It has been reported that A. radiobacter D-MIase (Hirose et al. 2001) and T. fusca D-MIase (Kasumi et al. 2014) displayed a dimer structure with subunit molecular weight of 44 and 41 kDa, respectively. However, the D-MIase from E. coli was a hexamer structure with subunit molecular weight of 45 kDa (Hu et al. 2016b). As for kinetic parameters of these reported D-MIases, the highest value of kcat of 788 s1 was found in D-MIase from T. fusca (Kasumi et al. 2014), whereas the highest value of kcat/Km of 19.7 mM1 s1 was M. mediterranea D-MIase (Saburi et al. 2018).

3.3.3.3

CEase

CEase was an aldose epimerase that could convert an aldose into its epimer aldose. This enzyme was first found in Ruminococcus albus as it could catalyze some β-1,4-linked disaccharides, such as mannobiose (mannose-β-1,4-mannose), cellobiose (glucose-β-1,4-glucose), and lactose (galactose-β-1,4-glucose) (Tyler and Leatherwood 1967). Under the action of CEase, mannobiose, cellobiose, and lactose could be converted to mannosyl glucose, glucosyl mannose, and galactosyl mannose, respectively. The action mechanism of CEase on these disaccharides is that these disaccharides have undergone epimerization reaction at their C-2 position. As it is known to us, D-mannose is an epimer of D-glucose at C-2 position. So, if D-mannose can be produced from D-glucose directly, the production advantages from D-glucose will be obvious than from D-fructose, because the price of D-glucose is cheaper than D-fructose. To test the potential ability for D-mannose production from D-glucose, Park et al. successfully identified a recombinant CEase from Caldicellulosiruptor saccharolyticus and investigated its application for D-mannose production from D-glucose (Park et al. 2011). This enzyme had an optimal temperature and pH of 50
C and 7.5. Its maximum enzyme activity does not require the participation of metal ions, which is similar to D-MIase but distinct from D-LIase. The half-lives of the enzyme were 142, 71, 35, 18, and 4.6 h at 60, 65, 70, 75, and 80
C, respectively. The C. saccharolyticus CEase has a kcat, Km, and

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kcat/Km values of 0.74 s1, 51.8 mM, and 0.01 mM1 s1, respectively. It is a monomer structure with a molecular weight of 47 kDa.

3.3.3.4

D-MEase

Recently, a cluster of uncharacterized proteins named MEase was identified and biochemically characterized as a new member of the acylglucosamine 2-epimerase (AGE) superfamily (Saburi et al. 2019). This enzyme could catalyze the intermolecular conversion of D-glucose to D-mannose at the C-2 position. However, the reported MEase did not exhibit the activity toward the substrates of β-1,4-mannobiose, N-acetyl-D-glucosamine, and D-fructose, which was different from the existing AGE superfamily enzymes that could catalyze these substrates. Therefore, the MEases from Runella slithyformis and Dyadobacter fermentans were the first reported AGE enzymes with potential application in producing D-mannose from D-glucose. Both MEases exhibited the same optimal temperature of 37
C and optimal weakly alkaline pH. The optimal pH value of R. slithyformis MEase and D. fermentans MEase were 7.8 and 8.1, respectively. Although the optimal metal ions experiment was not conducted in this literature, 24.4 and 22.8% yields of D-mannose were finally produced from 500 g L1 of D-glucose substrate at 50
C and pH 8.0. R. slithyformis MEase had a kcat, Km, and kcat/Km value of 4.77 s1, 73.5 mM, and 0.06 mM1 s1, respectively, whereas D. fermentans MEase had a kcat, Km, and kcat/Km value of 3.88 s1, 66.4 mM, and 0.06 mM1 s1. Both MEases showed a dimer structure with subunit molecular weight of 45 and 47 kDa for R. slithyformis MEase and D. fermentans MEase, respectively.

3.3.4

Strategies to Produce D-mannose Using Above Isomerases and Epimerases

As mentioned above, the biological production of D-mannose represents a promising method in the future market. There is a corresponding metabolic pathway of D-mannose in bacteria; therefore, the D-mannose can be absorbed and metabolized by microorganisms itself. Accordingly, there are no current reports of attempts to use microbial fermentation to produce D-mannose. The production of D-mannose from D-fructose or D-glucose using enzymatic isomerization or epimerization has been reported by many researchers (Fig. 3.3, Table 3.2). For example, Yu et al. characterized a thermostable D-LIase from a hyperthermophile T. oceani strain, which could produce 101.6 g L1 of D-mannose from 400 g L1 of D-fructose after 9 h of reaction under the conditions of pH 6.5, temperature of 65
C, and presence of 1 mM Mn2+ (Yu et al. 2016). The conversion yield and productivity were 25.4% and 11.28 g L1 h1, respectively. The D-LIase from C. polysaccharolyticus could produce 115 g L1 of D-mannose from 500 g L1 of D-fructose at 65
C, pH 6.5,

3 D-Mannose-Producing Isomerases and Epimerases: Properties, Comparisons, and. . .

a

b

or

or

or

+ D-mannose

D-fructose

D-LIase

55

D-glucose

D-MIase CEase D-MEase Glase

D-mannose

Cells Immobilized material

Fig. 3.3 Different strategies for production of D-mannose from D-fructose (a) or D-glucose (b) using various D-mannose-producing epimerases and isomerases

and 1 mM Mn2+ after a 24 h reaction with conversion yield of 23% (Wu et al. 2020a). Using free D-MIase from E. coli, 75 g L1 of D-mannose could be generated from 600 g L1 of D-fructose when reacted at 45
C and pH 7.0 for 2 h, and the conversion and productivity rates were 25% and 75 g L1 h1, respectively (Hu et al. 2016b). In addition to the free enzyme method, the immobilized strategy to produce D-mannose has also been employed. When immobilized D-LIase from P. stuartii on Duolite A568 beads was used, a highest conversion of 25% was achieved from D-fructose to D-mannose. At 35
C, pH 7.5, and 1 mM Mn2+ solutions, 75 g L1 of D-mannose was formed from 300 g L1 of D-fructose for 1 h, with productivity of 75 g L1 h1 after 23 cycles (Park et al. 2010a). The reverse reaction has also been investigated using immobilized D-MIase from Pseudomonas cepacia when producing D-fructose from D-mannose (Allenza et al. 1990). In another study, the A. radiobacter cells containing D-MIase were also immobilized by adsorption on chitosan or by glutaraldehyde crosslinking in the presence of albumin (Allenza et al. 1990). Results showed that 9 g of D-mannose could be continuously accumulated in the effluent (180 mL) from 200 g L1 of D-fructose for 14 days at 55
C and pH 8.0. As stated previously, research on the production of D-mannose from D-fructose using enzymatic methods lack extensive report. More enzymes should be screened from nature. Park et al. used the free CEase from C. saccharolyticus to produce D-mannose under the optimal reaction condition of 75
C and pH 7.5 (Park et al. 2011). After reaction of about 3 h, 75 g L1 of D-mannose was produced from 500 g L1 of D-fructose with conversion rate and productivity of 15% and 25 g L1 h1, respectively. Similarly, the free MEase from R. slithyformis could also produce 122 g L1 of D-mannose from 500 g L1 of D-fructose 48 h later with conversion rate and productivity of 24.4% and 2.54 g L1 h1 at pH 8.0, 50
C, respectively, whereas free D. fermentans MEase could generate 114 g L1 of D-mannose with conversion rate and productivity of 22.8% and 2.38 g L1 h1 under the same reaction condition (Saburi et al. 2019). In addition to single-enzyme technology, researchers have also used two enzyme co-expression strategies to produce D-mannose from D-glucose. For instance, Huang et al. adopted the free D-glucose isomerase (D-GIase) from Acidothermus cellulolyticus and free D-LIase from

Free D-LIase

Immobilization D-LIase Free D-MIase

Immobilization cells containing D-MIase

C. polysaccharolyticus

P. stuartii

A. radiobacter

Free CEase

Free D-MEase

Free D-MEase

Free D-GIase and Free D-LIase

C. saccharolyticus

R. slithyformis

D. fermentans

A. cellulolyticus T. dichotomicum

E. coli

Strategy Free D-LIase

Source T. oceani

400

500

500

500

200

600

300

500

pH 6.5, 65
C, 1 mM Co2+, 8 h

pH 8.0, 50
C, 48 h

pH7.5, 75
C, 3h pH 8.0, 50
C, 48 h

Reaction condition pH 6.5, 65
C, 1 mM Mn2+,9 h pH 6.5, 65
C, 1 mM Mn2+, 24 h pH 7.5, 35
C, 1 mM Mn2+, 1 h pH 7.0, 45
C, 2h pH 8.0, 55
C, 14 days

60

114

122

75

50 (9 g/180 mL)

15

22.8

24.4

15

25

25

25

75 150

23

Conversion (%) 25.4

115

D-mannose (g L1) 101.6

7.5

2.38

2.54

25

0.15

75

75

4.79

Productivity (g L1 h1) 11.28

Park et al. (2010a) Hu et al. (2016b) Hirose et al. (2003) Park et al. (2011) Saburi et al. (2019) Saburi et al. (2019) Huang et al. (2018b)

Reference Yu et al. (2016) Wu et al. (2020a)

D-LIase D-lyxose isomerase, D-MIase D-mannose isomerase, CEase cellobiose 2-epimerase; MEase, D-mannose 2-epimerase; D-GIase, D-glucose isomerase A. cellulolyticus, Acidothermus cellulolyticus

D-Glucose

Substrate D-Fructose

Substrate concentration (g L1) 400

Table 3.2 Production of D-mannose using different enzymes from different substrates under various reaction condition

56 H. Wu and Q. Chen

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T. dichotomicum to obtain D-mannose from D-glucose (Huang et al. 2018b). In which, the D-GIase can convert D-glucose to D-fructose, and then, the yield for D-fructose is further transformed to D-mannose through isomerization. Using this co-expressing system, 60 g L1 of D-mannose was gained from 400 g L1 of D-fructose after a reaction of about 8 h at pH 6.5, 65
C, and 1 mM Co2+. The conversion rate and productivity were 15% and 7.5 g L1 h1.

3.4

Applications

As a functional hexose with many excellent benefits and health effects, D-mannose has attracted many people’s attention and interest. Especially in modern society, many health problems, such as obesity and diabetes, emerged as unhealthy diet developed because of excessive consumption of high-fat and high-sugar foods. The low-sweetness, low-caloric health function of D-mannose displays its broad applications in food, medicines, and cosmetics and as feed additives. Due to its low sweetness and low calorie, D-mannose can be used as a dietary supplement for direct intake. In addition, D-mannose can improve the food texture as it exhibits a stable biochemical property, such as its use as stabilizer for ice cream (Sutton and Wilcox 2010). As described above, D-mannose can also be used as a cheaper material for synthesizing many high value-added medicinal products. For example, D-mannose can be used to produce mannitol through selective hydrogenation by ruthenium catalyst (Mishra and Hwang 2013). This sugar alcohol can help to minimize the risk of acute renal failure and be a highly effective dehydrating agent and osmotic diuretic (Chen et al. 2020). D-mannose is an auxiliary moisturizing agent used in skin care as the aloe polysaccharides are rich in mannan and glucomannan (Eshun and He 2004). In addition, D-mannose has found the ability to inhibit the infection of Salmonella typhi in chickens, indicating that D-mannose can be used to replace antibiotics to reduce environmental pollution and drug resistance in feed additives (van Immerseel et al. 2002).

3.5

Conclusion

As a functional sugar, D-mannose has broad applications in the food, cosmetics, and pharmaceutical industry. In particular, its low calorie and low sweetness properties make the market for D-mannose become more and more demanding. Highly efficient production of D-mannose has gained much attention and interest. In this chapter, the main content is focused on D-mannose-producing epimerases and isomerases including the biochemical characteristics and different strategies for the production of D-mannose. Also, a brief description on the functions and health effects, determination methods, and applications of D-mannose is also presented. Currently, different D-mannose-producing enzymes including D-LIase, D-MIase,

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CEase, and D-MEase are identified and characterized from many reports. However, there are currently insufficient work on the crystal structure information and analysis of these enzymes. And the research on the catalytic mechanism of these enzymes with their substrates urgently needs to be enhanced. An efficient enzyme suitable for industrial applications is a prerequisite for obtaining high yield of D-mannose. Of course, more D-mannose-producing enzymes should be screened from nature. Also, the food-grade expression system of these enzymes needs to be developed in the future.

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Mishra DK, Hwang J-S (2013) Selective hydrogenation of D-mannose to D-mannitol using NiO-modified TiO2 (NiO-TiO2) supported ruthenium catalyst. Appl Catal A General 453 (6):13–19 Moller DE, Berger JP (2003) Role of PPARs in the regulation of obesity-related insulin sensitivity and inflammation. Int J Obes 27:3 Mussatto SI, Carneiro LM, Silva JP, Roberto IC, Teixeira JA (2011) A study on chemical constituents and sugars extraction from spent coffee grounds. Carbohydr Polym 83(2):368–374 Oh D-K (2007) Tagatose: properties, applications, and biotechnological processes. Appl Microbiol Biotechnol 76(1):1–8. https://doi.org/10.1007/s00253-007-0981-1 Palleroni NJ, Doudoroff M (1956) Mannose isomerase of Pseudomonas saccharophila. J Biol Chem 218(1):535 Park CS, Kwon HJ, Yeom SJ, Oh DK (2010a) Mannose production from fructose by free and immobilized D-lyxose isomerases from Providencia stuartii. Biotechnol Lett 32(9):1305–1309. https://doi.org/10.1007/s10529-010-0300-2 Park CS, Yeom SJ, Lim YR, Kim YS, Oh DK (2010b) Substrate specificity of a recombinant D-lyxose isomerase from Serratia proteamaculans that produces D-lyxose and D-mannose. Lett Appl Microbiol 51(3):343–350. https://doi.org/10.1111/j.1472-765X.2010.02903.x Park CS, Kim J-E, Choi J-G, Oh D-K (2011) Characterization of a recombinant cellobiose 2-epimerase from Caldicellulosiruptor saccharolyticus and its application in the production of mannose from glucose. Appl Microbiol Biotechnol 92(6):1187–1196 Patel DH, Wi SG, Lee SG, Lee DS, Song YH, Bae HJ (2011) Substrate specificity of the Bacillus licheniformis lyxose isomerase YdaE and its application in in vitro catalysis for bioproduction of lyxose and glucose by two-step isomerization. Appl Environ Microbiol 77(10):3343–3350. https://doi.org/10.1128/AEM.02693-10 Pohl JB, Baldwin BA, Dinh BL, Rahman P, Smerek D, Sherazee N, Atkinson NS (2012) Ethanol preference in Drosophila melanogaster is driven by its caloric value. Alcohol Clin Exp Res 36 (11):1903–1912 Ranta K, Nieminen K, Ekholm FS, Poláková M, Roslund MU, Saloranta T, Leino R, Savolainen J (2012) Evaluation of immunostimulatory activities of synthetic mannose-containing structures mimicking the β-(1! 2)-linked cell wall mannans of Candida albicans. Clin Vaccine Immunol 19(11):1889–1893 Saburi W, Jaito N, Kato K, Tanaka Y, Yao M, Mori H (2018) Biochemical and structural characterization of Marinomonas mediterranea D-mannose isomerase Marme_2490 phylogenetically distant from known enzymes. Biochimie 144:63–73. https://doi.org/10.1016/j.biochi. 2017.10.016 Saburi W, Sato S, Hashiguchi S, Muto H, Iizuka T, Mori H (2019) Enzymatic characteristics of D-mannose 2-epimerase, a new member of the acylglucosamine 2-epimerase superfamily. Appl Microbiol Biotechnol 103:6559–6570. https://doi.org/10.1007/s00253-019-09944-3 Sharma V, Ichikawa M, Freeze HH (2014) Mannose metabolism: more than meets the eye. Biochem Biophys Res Commun 453(2):220–228. https://doi.org/10.1016/j.bbrc.2014.06.021 Shen SC, Wu J (2004) Maillard browning in ethanolic solution. J Food Sci 69(4):273–279 Stevens F, Stevens P, Hovis J, Wu T (1981) Some properties of D-mannose isomerase from Escherichia coli K12. Microbiology 124(1):219–223 Sutton RL, Wilcox J (2010) Recrystallization in model ice cream solutions as affected by stabilizer concentration. J Food Sci 63(1):9–11 Takasaki Y (1967) Kinetic and equilibrium studies on D-mannose-D-fructose isomerization catalyzed by mannose isomerase from Streptomyces aerocolorigenes. J Agric Chem Soc Jpn 31 (4):435–440 Takasaki Y, Takano S (1964) Studies on the isomerization of sugars by bacteria: part VIII. Purification and some properties of mannose isomerase from Xanthomonas rubrilineans S-48. Agric Biol Chem 28(9):605–609

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Takasaki Y, Hinoki K, Kataoka Y, Fukuyama S, Nishimura N, Hayashi S, Imada K (1993) Enzymatic production of D-mannose from D-fructose by mannose isomerase. J Ferment Bioeng 73(3):237–239 Tonnesen AS (1983) Clinical pharmacology and use of diuretics. Anesthesiology 11:217–236 Turner MW (2003) The role of mannose-binding lectin in health and disease. Mol Immunol 40 (7):423–429 Tyler TR, Leatherwood JM (1967) Epimerization of disaccharides by enzyme preparations from Ruminococcus albus. Arch Biochem Biophys 119(1):363–367 van Immerseel F, Cauwerts K, Devriese LA, Haesebrouck F, Ducatelle R (2002) Feed additives to control Salmonella in poultry. Worlds Poul Sci J 58(4):501–513 van Staalduinen LM, Park CS, Yeom SJ, Adams-Cioaba MA, Oh DK, Jia Z (2010) Structure-based annotation of a novel sugar isomerase from the pathogenic E. coli O157:H7. J Mol Biol 401 (5):866–881. https://doi.org/10.1016/j.jmb.2010.06.063 Wu H, Zhang W, Mu W (2019) Recent studies on the biological production of D-mannose. Appl Microbiol Biotechnol 103(21):8753–8761. https://doi.org/10.1007/s00253-019-10151-3 Wu H, Chen M, Guang C, Zhang W, Mu W (2020a) Characterization of a recombinant D-mannoseproducing D-lyxose isomerase from Caldanaerobius polysaccharolyticus. Enzym Microb Technol 138:109553. https://doi.org/10.1016/j.enzmictec.2020.109553 Wu H, Chen M, Guang C, Zhang W, Mu W (2020b) Identification of a novel recombinant D-lyxose isomerase from Thermoprotei archaeon with high thermostable, weak-acid and nickel ion dependent properties. Int J Biol Macromol 164:1267–1274. https://doi.org/10.1016/j.ijbiomac. 2020.07.222 Xu L, Guo D, Liu J, Zhao P, Wang Y (2014) A mixed catalyst for improving conversion rate of D-glucose to D-mannose. Chinese Patents. CN103831122A Yashoda HM, Prabha TN, Tharanathan RN (2007) Mango ripening – role of carbohydrases in tissue softening. Food Chem 102(3):691–698 Yeung B, Porter TJ, Vath JE (1997) Direct isoform analysis of high-mannose-containing glycoproteins by on-line capillary electrophoresis electrospray mass spectrometry. Anal Chem 69 (13):2510–2516. https://doi.org/10.1021/ac9611172 Yu L, Zhang W, Zhang T, Jiang B, Mu W (2016) Efficient biotransformation of D-fructose to D-mannose by a thermostable D-lyxose isomerase from Thermosediminibacter oceani. Process Biochem 51(12):2026–2033. https://doi.org/10.1016/j.procbio.2016.08.023 Zhang T, Pan Z, Qian C, Chen X (2009) Isolation and purification of D-mannose from palm kernel. Carbohydr Res 344(13):1687–1689 Zhang W, Zhang C, Liang G (2017) Study on increasing yield of D-mannose from D-glucose by epimerization conversion. Technol Dev Chem Ind 46(3):18–21 Zhang W, Huang J, Jia M, Guang C, Zhang T, Mu W (2019) Characterization of a novel D-lyxose isomerase from Thermoflavimicrobium dichotomicum and its application for D-mannose production. Process Biochem 83:131–136. https://doi.org/10.1016/j.procbio.2019.05.007 Zhao G, Wang G, Li J, Cao Y (2005) The orthogonal experiment analysis on the producing mannose from dextrose by catalyzing effect of ammonium molybdate. Contemp Chem Ind 34 (1):39–41

Chapter 4

L-Arabinose Isomerase: Sources, Biochemical Properties, and Its Use to Produce D-Tagatose Hao Wu and Wei Xu

4.1

Introduction

During long-term life, many people like to eat high-sugar and high-fat foods due to their better taste. However, this kind of preferential eating habits is not healthy. Long-term consumption of these high-calorie foods brings serious health problems, such as diabetes and obesity, and affects people’s quality of life. Therefore, demanding for alternative foods for sucrose with low calories and sweetness is gradually increasing and receiving great attention. Functional sweetener is one of the functional foods that have been widely studied in recent years. Its sweetness is similar to that of sucrose, but its energy is much lower than that provided by the same amount of sucrose, even without energy. Functional sweeteners include functional monosaccharides, oligosaccharides, and polyols. The most important type of monosaccharides in functional sugars is functional rare sugars. Rare sugars are defined as monosaccharides and their derivatives that rarely exist in nature, according to the International Society of Rare Sugars (ISRS) (Zhang et al. 2017). Recently, rare sugar has received increasing attention from scientific researchers and the public as it can be used as a low-calorie sweetener. For instance, rare sugar D-allulose (previously known as D-psicose) has the advantages of low energy, being difficult to absorb and utilize, and lower blood sugar. It can be used as a good substitute for edible sugar (Matsuo et al. 2001). Rare sugar D-allose can inhibit reactive oxygen species and hinder cancer cell proliferation (Murata et al. 2003). D-Tagatose is another attracted rare ketohexose firstly found in the gum of a tropical evergreen plant in nature. Recently, it has also been discovered in dairy products such as milk, milk power, yogurt, and cheese (Adachi 1958). D-Tagatose is

H. Wu (*) · W. Xu State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_4

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the epimer of D-fructose and the ketose isomer of D-galactose (Fig. 4.1). Its sweetness is 92% of that of sucrose, and its calories are only 1/3 of that of sucrose, which makes it an ideal sucrose substitute. D-Tagatose has been proven to possess the function of anti-diabetes and anti-obesity (Lu et al. 2008). Long-term intake of foods containing D-tagatose in patients with type 2 diabetes can reduce weight and increase the amount of high-density lipoprotein cholesterol and that no adverse effects have been found in blood sugar control and other biochemical indicators (Donner et al. 2010). In addition, D-tagatose can also inhibit dental caries, gum disease, and periodontitis and other diseases by changing the subgingival flora (Lu and Levin 2002). Due to it exhibiting many excellent functions on people’s health, production of rare sugar D-tagatose has received great interest. The efficient production of rare sugars has become an important goal as they have wide applications in food, medicinal, and pharmaceutical industries. In particular, the enzymatic approaches for rare sugar production show certain advantages. In the past few decades, Professor Ken Izumori who is the director at the Rare Sugar Research Center of Kagawa University in Japan creatively built a novel and systematic strategy to synthesize rare sugars. According to his theory, all rare sugars could be cyclically converted through epimerization, isomerization, and oxidationreduction (Izumori 2002, 2006). However, this biological process depends on various enzymes including aldose-ketose isomerases (EC 5.3.1), ketose C-3 epimerases (EC 5.1.3), polyol dehydrogenase, oxidoreductases (EC 1.1), and L-arabinose isomerase (L-AIase, EC 5.3.1.4). For example, rare sugar D-allulose could be converted from D-fructose under the action of D-psicose 3-epimerase (Fig. 4.1) (Zhang et al. 2016). D-Allose could be synthesized from D-allulose material by L-rhamnose isomerase (EC 5.3.1.14), whereas D-allulose could be obtained from the epimerization of D-fructose (Chen et al. 2018). Besides, D-tagatose could be produced from D-galactose using L-AIase (Oh 2007). This enzyme is an aldose-ketose isomerase which can also catalyze the isomerization reaction between L-arabinose and L-ribulose due to its broad spectrum of substrates. It was first purified from the strain Lactobacillus plantarum culture broth by Heath and its workers (Heath et al. 1958). Previous study has found that L-AIase exists in many microorganisms because this enzyme plays an important part in microbial metabolism. The strain Escherichia coli can use L-arabinose as a sole carbon source for growing because the encoded L-AIase can catalyze L-arabinose to L-ribulose. L-Ribulose is further transformed to L-ribulose-5-phosphate by L-ribulokinase (EC 2.7.1.16). Then, L-ribulose-5-phosphate is continued to convert to D-xylulose-5-phosphate under the action of L-ribulose-5-phosphate 4-epimerase (EC 5.1.3.4), which will enter the pentose phosphate pathway to provide energy and reducing force (Xu et al. 2018). Thus, many concerns are paid on the biological production of D-tagatose from D-galactose using L-AIases from various microorganisms by isomerization. Compared to the chemical production of D-tagatose using calcium catalyst and strong acid, which causes the heavy pollution to the environment and produces many by-products, biological methods show the superiority of environmental friendliness. It is worth mentioned that D-galactose can be produced from the hydrolyzed D-lactose by lactase (β-galactosidase), whereas lactose is the main component of

Fig. 4.1 The structure of D-fructose, D-psicose, D-allose, D-lactose, D-galactose, and D-tagatose

4 L-Arabinose Isomerase: Sources, Biochemical Properties, and Its Use to Produce. . . 65

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whey which is a cheap and abundant by-product generated from cheese or casein in industry. This method displays a great potential possibility for D-tagatose green production with regard to the production cost and economic considerations. An enzyme with superior industrial properties is essential for the efficient production of D-tagatose. In this chapter, the D-tagatose-producing L-AIase is introduced and described in detail. This article focuses on the sources of L-AIases and its characterization including temperature, pH, metal ion dependence, and other characteristics. In addition, the substrate specificity, kinetic parameters, and molecular modification of L-AIase are discussed and summarized. Finally, a brief introduction to D-tagatose and the different strategies for D-tagatose production are outlined as well.

4.2 4.2.1

Identification of L-Arabinose Isomerase The Sources

As we all know, the survival of microorganisms has certain adaptability to its environment. Microorganisms isolated in high-salt environments often possess halophilic properties, whereas enzymes secreted by microorganisms in hightemperature or low-temperature environment often show high thermostable or cold-tolerance behavior. Currently, approximately 30 L-AIases have been isolated and characterized from various environments which are psychrotolerant, mesophilic, thermophilic, and hyperthermophilic (Table 4.1) (Xu et al. 2018). This kind of work has been going on for more than 20 years. The microorganisms from the following section have been verified to produce L-AIase, including Thermotoga neapolitana from submarine thermal vent (Belkin et al. 1986; Kim et al. 2002), Thermotoga maritima from sea floor (Huber et al. 1986; Lee et al. 2004), Geobacillus thermodenitrificans from a high-temperature environment of Korean compost samples (Kim and Oh 2005), Bacillus stearothermophilus from Tunisian hot spring soil (Ali et al. 1999; Rhimi and Bejar 2006), Lactobacillus plantarum from grass silage (Chouayekh et al. 2007), Lactobacillus fermentum from traditional Chinese pickles (Xu et al. 2011), Bacillus coagulans from soil (Patel et al. 2006; Zhou and Wu 2012), Pediococcus pentosaceus from fermented pickles (Men et al. 2014), Alicyclobacillus hesperidum from hot spring sludge (Wang et al. 2012; Fan et al. 2014), Lactobacillus brevis from fermented cabbage (Du et al. 2019), and Enterococcus faecium from raw cow milk (Manzo et al. 2019).

4.2.2

Temperature

Temperature factor is an important biochemical property of enzyme. An optimal temperature and good thermostability are beneficial to maximize the enzyme’s

7.5

8.5

7.5–8.0

CAE46769

AAD35365

AM701769

NC

Sea floor

Unspecified

Fresh meat and fish

Unspecified

Chinese pickles

T. maritime

B. stearothermophilus Tunisian hot spring soil

Grass silage

Tan. mathranii

G. thermodenitrificans High tempera- AY302754 ture environment of Korean compost samples

AJ866972

Unspecified

Escherichia coli

L. plantarum

B. licheniformis

L. sakei

B. subtilis

L. fermentum

HM150718

NC

NC

NC

NC

6.5

7.5

5.0–7.0

7.5

7.5

8.0

8.0

AY0283379 7.0

Submarine thermal vent

Optimum pH

T. neapolitana

GenBank accession

Sources

Enzyme source

65

32

30–40

50

60

80

70

90

65

30

85

Optimum temperature ( C) 112.33

33.40

11.2 207.58 34.13 241.73 NR

Mn2+, Co2+ Mn2+, Co2+ Mn2+, Mg2+ Mn2+ Mn2+, Co2+

113.6

Mn2+

None

38.64

NR

Mn2+, Co2+

Mn2+

+

Mn2+, Fe2 NR

Co2+, Mn2+

Optimum metal ions L-Ara

kcat (s 1)

9.02

NR

10.13

NR

1.86

8.06

3.4

8.5

NR

NR

6.26

D-Gal

Table 4.1 The biochemical characteristics of different L-arabinoses from different sources

NR

119.87

31.6

369

43.4

28.57

142

31

80

NR

116

L-Ara

Km (mM) D-Gal

60

NR

59

NR

69.7

57

408

60

120

300

250

19

121

64.8

34

15.5

71.4

48

74.8

NR

NR

58.1

L-Ara

1

9.02

NR

10.3

NR

1.6

8.48

0.5

8.5

NR

NR

3.24

D-Gal

kcat/Km (min mM 1)

53

56

54

53

55

56

56

57

52

NR

57

Molecular weight by SDS-PAGE (kDa)

NC

Homodimer

Tetramer

Homodimer

Hexamer

Tetramer

Tetramer

Tetramer

Tetramer

NC

NC

Polymer

(continued)

Xu et al. (2011)

Kim et al. (2010)

Rhimi et al. (2010)

Prabhu et al. (2008)

Chouayekh et al. (2007)

Rhimi and Bejar (2006)

Kim and Oh (2005)

Lee et al. (2004)

Jorgensen et al. (2004)

Yoon et al. (2003)

Kim et al. (2002)

Reference

4 L-Arabinose Isomerase: Sources, Biochemical Properties, and Its Use to Produce. . . 67

Soil

NC

Fermented pickles

Hot spring sludge

NC

Fermented cabbage

Raw cow milk NC

B. coagulans

Arthrobacter sp.

P. pentosaceus

A. hesperidium

C. hylemonae

L. brevis

E. faecium

6.0

5.0-9.0

7.0

10.5

5.5

7.0

7.0–7.5

50

65

50

70

50

47–52

70

95

15–35

Optimum temperature ( C)

NR

NR 71.84 NR NR

NR

Mn2+, Co2+ Co2+ Mg2+ Mn2+, Co2+, Mg2+ Mn2+

60.95

Mn2+ None

0.88

Ni2+

2.52

0.26

28.39

1.13

3.19

NR

NR

2.17

D-Gal NR

Mn2+ NR

Optimum metal ions L-Ara

kcat (s 1)

NR

NR

NR

105.2

NR

NR

106

78.5

33.7

L-Ara

Km (mM) D-Gal

225

129

7.7

54.7

66

NR

NR

25.2

52.1

NR

NR

NR

41.0

NR

NR

34.5

0.67

NR

L-Ara

1

0.68

0.12

221.4

1.2

2.9

NR

NR

5.16

NR

D-Gal

kcat/Km (min mM 1)

55–56

60.1

57

55

54

55

56

56.3

55

Molecular weight by SDS-PAGE (kDa) Polymer

NC

Monomer

NC

NC

NC

Hexamer

NC

NC

Tetramer

Manzo et al. (2019)

Du et al. (2019)

Nguyen et al. (2018)

Fan et al. (2014)

Men et al. (2014)

Wanarska and Kur (2012)

Zhou and Wu (2012)

Li et al. (2011)

Rhimi et al. (2011)

Reference

T. neapolitana, Thermotoga neapolitana; Tan. mathranii, Thermoanaerobacter mathranii; T. maritim, Thermotoga maritime; G. thermodenitrificans, Geobacillus thermodenitrificans; B. stearothermophilus, Bacillus stearothermophilus; L. plantarum, Lactobacillus plantarum; B. licheniformis, Bacillus licheniformis; L. brevis, Lactobacillus brevis; P. pentosaceus, Pediococcus pentosaceus; A. hesperidum, Alicyclobacillus hesperidum; B. coagulans, Bacillus coagulans; L. fermentum, Lactobacillus fermentum; C. hylemonae, Clostridium hylemonae; E. faecium, Enterococcus faecium; A. flavithermus, Anoxybacillus flavithermus; L. sakei, Lactobacillus sakei; B. subtilis, Bacillus subtilis L-Ara L-arabinose, D-Gal D-galactose NR not reported, NC not clear

NC

NC

EJY56736.1 7.0

JN377428

JN642528

NC

HQ589928

Hot spring sludge

A. flavithermus

5.5–6.5

A0KWX7

Wooden pier located in a brackish estuary

Shewanella sp.

Optimum pH

GenBank accession

Sources

Enzyme source

Table 4.1 (continued)

68 H. Wu and W. Xu

4 L-Arabinose Isomerase: Sources, Biochemical Properties, and Its Use to Produce. . .

69

activity. Just like other isomerases, such as D-lyxose isomerase and L-rhamnose isomerase, a higher temperature and slightly acidic pH optimum are the two crucial enzymatic properties required for the production of D-tagatose using isomerase, which is generally recognized in the sugar industry. Therefore, the thermo and acid resistance L-AIase has become a research hotspot. A higher temperature reaction system can not only prevent the microbial contamination and increase the substrate and product solubility but also reduce the viscosity of the system solution, thereby increasing the reaction rate (Fan et al. 2014). More importantly, the isomerization reaction between D-galactose and D-tagatose catalyzed by L-AIase is an equilibrium process, suggesting that the final yield of D-tagatose is in a thermodynamic equilibrium. However, the equilibrium process can be affected by the temperature. Therefore, sugar isomerization reactions tend to initiate under higher-temperature environments in industry. However, a more contradictory point is that Maillard reactions caused by the interaction of protein and reducing sugar are prone to occur under high-temperature conditions. Thus, L-AIase with a slightly acidic pH optimum is more popular in sugar industry because this weakly acidic environment can reduce nonenzymatic browning and inhibit Maillard reaction. To identify suitable L-AIase with industrial properties, researchers have carried out unremitting work. Currently, the optimal temperatures of identified L-AIases are ranging from 15 to 95  C, exhibiting a wide temperature range (Fig. 4.2). This difference in optimum temperature is closely related to their sources. For example, the L-AIase with the highest optimum temperature of 95  C was derived from hyperthermophilic strain Anoxybacillus flavithermus whose growing environment was hot spring sludge (Li et al. 2011). The L-AIase with the lowest optimum

Fig. 4.2 The optimal pH and temperature of L-AIases from different microorganisms

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H. Wu and W. Xu

temperature of 15  C was derived from psychrotolerant strain Shewanella sp. whose growing environment was wooden pier located in a brackish estuary (Rhimi et al. 2011). It can be seen from Table 4.1 that most of the L-AIases coming from thermophilic and hyperthermophilic microorganisms possess a higher optimum temperature of over 60  C. At the same time, these L-AIases with higher optimum temperatures show better thermostability. For instance, the half-life time of L-AIase from T. maritime was 185 min at 90  C in the presence of 5 mM Mn2+ (Lee et al. 2004). The half-life time of L-AIase from T. neapolitana was 120 min at 90  C in the presence of 1 mM Co2+ and Mn2+ (Kim et al. 2002). Half-life time of B. stearothermophilus L-AIase was 60 min at 75  C without metallic ions. If adding 0.2 mM Co2+ or 1 mM Mn2+, its half-life time was increased to 110 min (Rhimi and Bejar 2006). However, the L-AIase from Mycobacterium smegmatis had a half-life time of 10 min at its optimal temperature of 45  C (Izumori et al. 1978). Interestingly, the mesophilic L-AIase from L. plantarum was perfectly stable after a 120-min heating at 70  C in the presence of 1 mM Mn2+ or 1 mM Mn2+ plus 0.5 mM Co2+ (Chouayekh et al. 2007). If not supplemented with metal ions or only added 0.5 mM Co2+, the enzyme was completely inactivated after 30 min. These results suggest that Mn2+ is involved in the enzyme stabilization at high temperatures besides its role in the catalytic mechanism, whereas the Co2+ seems to be essentially implicated in the isomerization reaction.

4.2.3

pH

pH is another non-ignorable parameter to deeply understand the enzymatic properties. As we all know, the charged state and conformation of the enzyme are affected by various pH environments, thereby influencing the activity of the enzyme. As stated above, a slightly acidic environment is favored for the production of D-tagatose in industry due to the Maillard reaction which can be inhibited in this condition. Currently, the identified L-AIases show a wide optimal pH range of 5.0–10.5 (Table 4.1). Nevertheless, most L-AIases exhibit alkaline pH properties. For example, L-AIase derived from thermophile strain Bacillus stearothermophilus displayed an optimal pH of 7.5–8.0 (Rhimi and Bejar 2006). Hyperthermophilic eubacterium T. maritime L-AIase exhibited an optimal pH of 7.5 (Lee et al. 2004). Thermophilic bacterium G. thermodenitrificans had an optimal pH of 8.5 (Kim and Oh 2005). Interestingly, psychrotolerant bacterium Pseudoalteromonas haloplanktis L-AIase showed maximal activity at pH 8.0 (Xu et al. 2016). On the other hand, it also indicates that the optimal pH of L-AIase is not directly related to temperature and its source. It is worth noting that among the L-AIase reported so far, A. flavithermus-derived ones have the highest optimum temperature (95  C) and the highest pH (9.5–10.5) (Table 4.1). The recombinant Arthrobacter sp. L-AIase was optimally active at a broad pH range of 5–9 (Wanarska and Kur 2012). This characteristic makes the enzyme have good adaptability in the process of producing D-tagatose and also makes it have better advantages compared with other L-AIases.

4 L-Arabinose Isomerase: Sources, Biochemical Properties, and Its Use to Produce. . .

71

The stability of pH is also an evaluation index for the performance of L-AIase, which is of great significance for ensuring long-term biotransformation in industry. For example, psychrotolerant bacterium Pseudoalteromonas haloplanktis L-AIase was stable at the pH range of 7.0–8.5, and 70–90% of the residual activity could be reserved after incubation for 10 h (Xu et al. 2016). However, this enzyme showed low optimal temperature of 40  C. The L. plantarum NC8 L-AIase retained 89% of its activity after incubation for 24 h at pH 5.0 (Chouayekh et al. 2007). The A. cellulolyticus L-AIase retained 80% of its activity at pH 6.0 for 24 h (Cheng et al. 2010). Nonetheless, these L-AIases showed optimal alkaline pH properties. These are not the expected enzymes with high temperature and weak acid characteristics in the industry. Therefore, molecular modification of L-AIases must be carried out in the subsequent research to make the mutant enzymes meet the requirements of the industry.

4.2.4

Metal Ion

Metal ions play an important role in the conversion of enzymes to produce rare sugars. They often play a role in promoting catalytic activity and enhancing the stability of enzyme structure. Aldose-ketose isomerization is a simple but mechanistically ambiguous reaction which involves the formal transfer of H2 from C-2 and O-2 to C-1 and O-1 of an R-hydroxy aldehyde to form the corresponding R-hydroxy ketone (Nagorski and Richard 2001). Researches revealed that all aldose-ketose isomerases acting on sugars without phosphate groups use a metal ion in catalysis (Nagorski and Richard 2001; Manjasetty and Chance 2006). Without exception, as an aldose-ketose isomerase, the activity or structural stability of L-AIase also requires the participation of metal ions. As shown in Table 4.1, metal ions Co2+, Mn2+, Fe2+, Mg2+, and Ni2+ have been identified as an optimal cofactor for the activity of L-AIases. For example, Mg2+ was proved to be the optimal metal ion for enhancing the activity of L-AIase from Clostridium hylemonae by 20% when compared with the control group, while there was no significant effect for the addition of Ni2+, Zn2+ , Ca2+, and Cu2+ (Nguyen et al. 2018). Lee et al. reported that the activity of T. maritima L-AIase which was dialyzed against 10 mM EDTA in advance could be recovered by the adding of Mn2+ or Co2+, whereas other metal ions such as Mg2+, Ca2+, Fe2+, and Ni2+ were poor activators (Lee et al. 2004). On the contrary, the activity of L-AIase from A. flavithermus was inhibited by Zn2+, Fe3+, Mg2+, Co2+, Cu2+, and Mn2+, but increased by Ni2+ (Li et al. 2011). However, Rhimi et al. found that the L-AIase from B. stearothermophilus was not a strictly metal ion-dependent enzyme, because the EDTA-treated enzyme showed the isomerization activity at 65  C (Rhimi and Bejar 2006). Interestingly, the effect of metal ions on the activity of B. stearothermophilus L-AIase was related to temperature. Although addition of Co2+ and Mn2+ had no effect on the activity of EDTA-treated enzyme at 65  C, its activity indeed increased to be 140% of its control (presence of

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Mn2+ and Co2+) at 80  C, indicating that metal ions might be implicated to the stabilization of the enzyme at higher temperatures over 65  C. The phenomenon that metal ions can help to increase the structure stability of L-AIase has been verified in other study, especially for the L-AIase derived from hyperthermophilic and thermophilic strains. For example, the activity of L-AIase derived from hyperthermophile T. neapolitana strain lost over 50% after incubation at 80  C for 2 h without the presence of metal ions. If it was incubated in the presence of Co2+ or Mn2+, the enzyme activity could retain 80% of its original activity (Kim et al. 2002). The L-AIase from hyperthermophilic eubacterium T. maritima was unstable at 90  C in the absence of metal ions. If the T. maritima L-AIase solution was added 5 mM Mn2+ or 1 mM Co2+, its thermostability was greatly increased and that its activity was not loss after incubating with the presence of these metal ions at 80  C for 240 min (Lee et al. 2004).

4.2.5

Substrate Specificity and Kinetic Parameters

L-AIase can catalyze the isomerization reaction between L-arabinose and L-ribulose, D-galactose, and D-tagatose. As shown in Table 4.1, almost all the reported L-AIase displayed the substrate specificity for L-arabinose and D-galactose and that these L-AIases had higher specificity toward L-arabinose than D-galactose. However, the L-AIase from A. hesperidum (Fan et al. 2014) and A. flavithermus (Li et al. 2011) represented a much higher affinity toward D-galactose than L-arabinose, which was different from many other reported L-AIases. Specially, L-AIase from B. subtilis (Kim et al. 2010) and P. haloplanktis (Xu et al. 2016) expressed no detectable activity when using D-galactose as substrate. Molecular docking study indicated that these two L-AIases could only immobilize and recognize L-arabinose as substrate. Besides, some L-AIases also showed slightly activity to other substrates. For example, the L-AIase from G. thermodenitrificans exhibited weak activity to D-allose, D-ribulose, D-fucose, D-xylose, and D-gulose, and no activity to D-glucose, D-mannose, and L-fucose substrate (Kim and Oh 2005). Similarly, the A. flavithermus L-AIase showed activity toward D-xylose (Li et al. 2011). Kinetic parameters, such as kcat/Km, are key index used for evaluating the catalytic efficiency of enzyme. The value of kcat/Km has important reference value for the industrial application of L-AIase. As shown in Table 4.1, the highest kcat/Km value of 221.4 min 1 mM 1 toward D-galactose was found the L-AIase from C. hylemonae (Nguyen et al. 2018). On the other hand, the B. subtilis L-AIase displayed the highest kcat/Km value of 121 min 1 mM 1 toward L-arabinose, though this enzyme can’t catalyze the D-galactose substrate (Kim et al. 2010).

4 L-Arabinose Isomerase: Sources, Biochemical Properties, and Its Use to Produce. . .

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73

Molecular Modification of L-Arabinose Isomerase Structure of L-AIase

Analyzing the structure of a protein helps people understand its catalytic mechanism and functional properties. As early as 1977, scientists had investigated the structure of L-AIase by using electron microscopes techniques (Wallace et al. 1978). However, it was not until 30 years later that the first crystal structure of L-AIase from E. coli was resolved with PDB number of 2AJT (Fig. 4.3) (Manjasetty and Chance 2006). The crystal structure shows that the enzyme is a hexamer with three domains including C-terminal, N-terminal, and a central domain. The monomer structure is composed of 500 amino acids and includes 16 β-strands and 17 α-helices. Besides, multiple sequence alignment of L-AIase from E. coli with other thermophilic and hyperthermophilic counterparts found some highly conserved amino acids and secondary structure elements in these domains, such as M351, I373, Y335, and H449 in C-terminal; H128, Q125, Y19, L18, and Q16 in N-terminal; and M185 and F279 in central domain. These amino acid residues play an important part in regulating the entrance of substrate into the active site and the releasing process of product. In addition, the E306 and E333 are identified as key amino acids which are responsible for the catalysis, and H350 and H450 are the key binding sites for the metal ions to stabilize the conformation of the enzyme. Similar to most of other sugar isomerases, such as L-fucose isomerase (Seemann and Schulz 1997) and D-lyxose isomerase (van Staalduinen et al. 2010), the catalytic mechanism of L-AIase catalyzing the isomerization of D-galactose to D-tagatose is the enediol mechanism (Manjasetty and Chance 2006). E306 and E333 of E. coli L-AIase might be involved in the similar proton transfers. Besides, the structure of L-AIase from Geobacillus kaustophilus was also resolved including its apo form (PDB 4R1O), holo form (4R1P), and holo form with L-arabitol (4R1Q), which showed in hexamer form (Choi et al. 2016). However, there are no available crystal structures of other multimeric L-AIases in the PDB database. As seen from Table 4.1, the L-AIase from Lactobacillus brevis exists as a monomer in solution with a molecular weight

Fig. 4.3 The crystal structure of L-AIase from different microorganisms of E. coli (A, PDB 2AJT), L. fermentum (B, PDB 4LQL), G. kaustophilus (C, PDB 4R1O)

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of 60.1 kDa (Du et al. 2019). The molecular mass of Bacillus licheniformis L-AIase was estimated to be 53 kDa by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and 113 kDa by gel filtration chromatography, indicating a homodimer form (Prabhu et al. 2008). Interestingly, the tetramer forms of L-AIases in solution were also identified from the strain of Thermoanaerobacter mathranii (Jorgensen et al. 2004), Thermotoga maritima (Lee et al. 2004), Geobacillus thermodenitrificans (Kim and Oh 2005), and Bacillus stearothermophilus (Rhimi and Bejar 2006). These studies showed that the structure of L-AIase has a certain diversity and difference.

4.3.2

Lowing the Optimal pH

As we all know, a slightly acidic pH is beneficial for the production of functional rare sugars in industry. However, most reported L-AIases showed alkaline pH properties including E. coli L-AIase (pH 8.0) (Yoon et al. 2003), Tan. mathranii L-AIase (pH 8.0) (Jorgensen et al. 2004), and G. thermodenitrificans L-AIase (pH 8.5) (Kim and Oh 2005). Therefore, scientists have tried their best to conduct the molecular modification of L-AIases to lower the optimal pH through protein engineering via semirational design from the amino acid sequence comparison insights. Lee et al. conducted the homologous alignment of alkalophilic and acidophilic L-AIase and revealed the region-specific amino acid that would contribute to the pH dependence of activity and stability (Lee et al. 2012). They also suggested that altering the charged state of amino acids near catalytic sites and the net charge of a region of the protein might be an efficient strategy to modify the pH dependence of the activity and stability. Generally, the acid-base replacement is a recognized theory used to guide the molecular modification of changing the optimal pH of enzyme. Lee et al. has found that the residue amino acid Lys-269 in L-AIase from A. acidocaldarius plays an important role in the determination of the pH optima (Lee et al. 2005). When the Lys-269 was mutated to Glu-269 by site-directed mutagenesis, the pH optima were changed from 6.0 to 7.0. In addition, the replacement of Glu to Lys of L-AIase at 268 position from Bacillus halodurans resulted in the optimal pH changing from 8.0 to 7.0. In another study, although mutant H18T of L-AIase from Geobacillus stearothermophilus showed greater substrate specificity and better thermostability, its activity under acidic conditions was relatively low (Laksmi et al. 2020). Based on random mutagenesis using error-prone PCR closed to the binding area of H18T, a double H18T/Y234C mutant displayed 1.8-fold and 3-fold higher activity than H18T and wild-type L-AIase at pH 6.0. The increasing activity might be attributed to the change in the binding pocket area involving residue 234 and further identified the importance of this residue amino acid in improving the activity under acidic conditions.

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75

Increasing the Catalytic Activity Toward D-Galactose

As described above, it could be indicated that most currently reported L-AIases displayed a higher affinity to L-arabinose compared to D-galactose. A higher catalytic activity to D-galactose will be beneficial to enhance the production of D-tagatose through the isomerization of L-AIase. At present, the random mutagenesis techniques have been widely used in the improving catalytic activity of L-AIase toward D-galactose. Through an error-prone polymerase chain reaction using L-AIase gene from G. stearothermophilus, the three mutant L-AIase (M322V/ S393T/V408A) showed different biochemical characteristics when compared with wild-type enzyme. The maximum isomerization activity of mutant L-AIase was changed to pH 8.0, 65  C, and 1.0 mM Co2+, whereas the wild-type enzyme was pH 8.0, 60  C, and 1.0 mM Mn2+ (Kim et al. 2006). Using 10 mM D-galactose as substrate, the D-tagatose yield from D-galactose was 45% by mutated enzyme and 28.5% for wild-type enzyme after 10 h incubation at pH 8.0 and 60  C. Kim et al. conducted another challenging work about the molecular modification of L-AIase from G. thermodenitrificans in terms of its catalytic activity toward D-galactose (Kim et al. 2014). They claimed that the F280N variant of L-AIase was identified as a D-galactose isomerase. Firstly, they have performed the molecular docking study of D-galactose with double-site variant (C450S/N475K). Then the residues (M186, F280, I371) near to D-galactose O6 were identified as potential key amino acids involved in the substrate specificity. After characterizing the site-directed mutagenesis of the three residue variants, the triple-site (F280N) variant enzyme displayed 2.1-fold kcat/Km value for D-galactose than L-arabinose, while the double-site variant (C450S/N475K) had 43.9-fold higher kcat/Km value for L-arabinose than for D-galactose. Molecular dynamic simulation study revealed that the increased activity toward D-galactose for triple-site variant might be due to the lower binding energy when compared with the double-site variant. Recently, another study found that the mutant L-AIase (H18T) from G. stearothermophilus exhibited increased activity for D-galactose when compared with wild-type enzyme. Using D-galactose as substrate, the substrate specificity for D-galactose was increased by 45.4% after replacing histidine with threonine at 18 position (Laksmi et al. 2018).

4.4

Brief Description of D-Tagatose

D-Tagatose is an epimer of D-fructose at the C-4 position and the isomer of D-galactose. The chemical formula and molecular weight are C6H1206 and 180.16 g/mol, respectively. Pure D-tagatose is a white crystalline substance with odorless smell. Besides, this rare sugar has high solubility in water compared with ethanol. At the temperature 21  C, 58% (w/w) of D-tagatose can dissolve in water, whereas only 0.2 g of it can dissolve in 1 L ethanol. In the 1% (w/v) aqueous solution, the specific optical rotation of D-tagatose is 5 . Its calorie is 1.5 kcal g 1. The melting

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temperature and glass transition temperature are 134  C and 15  C, respectively. It is very stable under the environment of pH 2–7. This sugar can be easily prone to happen the Maillard reaction at high temperature, while at low temperature, the caramelization reaction is also occurred (Kim 2004; Oh 2007). D-Tagatose can be metabolized by many microorganisms because of the presence of D-tagatose-6-phospate pathway in it which is a branch of galactose metabolism (Rosey et al. 1991). The tagatose-6-phosphate pathway bypasses fructose 1,6-diphosphate, which is a regulatory intermediate for central carbon flux and growth rate (Kim 2004; Bond et al. 1998). However, there is no D-tagatose-6phosphate pathway in higher animals including humans so that the D-tagatose can’t be broken down spontaneously. Orally adapted rat experiment showed that 68% of D-tagatose was decomposed to CO2 though it is not metabolized. During this process, the intestinal microbial flora may play a crucial role (Laerke et al. 2000).

4.5

Physiological Function and Benefits

The low-calorie value makes D-tagatose a better substitute food for sucrose. Currently, this rare sugar has been approved as a “generally recognized as safe” (GRAS) material for human consumption (Levin 2002). With the deepening of research, many important physiological functions and health benefits of D-tagatose have gradually been understood by the public. D-Tagatose has been widely used in food fields of fruit juices, beverages, chewing gum, and pharmaceuticals. As a prebiotic, D-tagatose can improve the intestinal microbial flora because the probiotic flora in the intestines, such as lactic acid bacteria, can use D-tagatose to reduce pH and increase the concentration of ATP, which is conducive to the growth of probiotics and inhibits the growth of potential pathogenic bacteria such as E. coli (Laerke and Jensen 1999). In addition, D-tagatose in the intestine can be fermented to produce short-chain fatty acids such as propionic acid, butyric acid, and valeric acid, which can promote the proliferation of lactobacilli, improve the immunity of the intestinal mucosa, and prevent colon cancer (Venema et al. 2005; Laerke et al. 2000). D-Tagatose can also increase the content of high-density lipoprotein (HDL), and HDL is a protective factor for coronary heart disease, which can transport excess cholesterol in the surrounding tissues back to the liver and convert it into bile acid or direct excretion from the intestines through bile, thereby reducing the incidence of coronary heart disease (Lu et al. 2008).

4.6

Production of D-Tagatose Using L-Arabinose Isomerase

As an isomer of D-galactose, D-tagatose has a low calorie and can be used as a sugarsubstitute bulking agent in food. Its higher efficient production and preparation has attracted more and more attentions from the researchers. Immobilized cell or enzyme

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techniques are mature methods being applied in the production of sugar in the longterm transformation in industry, and these strategies have the advantages of improving the utilization of enzymes and reducing the production costs. Therefore, most of the study regarding the D-tagatose production has focused on the immobilized method (Fig. 4.4, Table 4.2). By immobilizing the L-AIase from E. coli through covalent binding to agarose, 104.1 g L 1 of D-tagatose was produced from 500 g L 1 of D-galactose with a conversion rate of 20.82% and productivity of 2.17 g L 1 h 1 under the condition of pH 7.0 and 30  C (Kim et al. 2001). In another study, the E. coli cells containing L-AIase from Tan. mathranii were immobilized in glutaraldehyde and polyethylenimine material through cross-linking interaction (Jorgensen et al. 2004). Finally, 126 g L 1 of D-tagatose was obtained from 300 g L 1 of D-galactose in 48 h with a conversion rate of 42% and productivity of 2.63 g L 1 h 1 under the condition of pH 6.9 and 65  C. Besides, chitopearl bead was also employed as material to immobilize L-AIase from Geobacillus stearothermophilus to convert D-galactose to D-tagatose because this material is stable at high temperature (over 60  C) (Cheon et al. 2008). The chitopearl beads containing L-AIase expressed in Bacillus subtilis could produce 4.3 g L 1 of D-tagatose from 10 g L 1 of D-galactose about 62 h with productivity of 0.07 g L 1 h 1 at 60  C and pH 8.0. It is worth mentioned that the L-AIase Gali 52 from G. stearothermophilus which was immobilized in alginate displayed the highest conversion rate of 58.2% from 100 g L 1 of D-galactose at 60  C and pH 8.0 to the best of our knowledge (Kim et al. 2003a). If optimizing the bead size, L/D (length/diameter) of reactor, dilution rate, total loaded enzyme amount, and substrate concentration, the L-AIase which was immobilized in alginate from G. stearothermophilus could catalyze 300 g L 1 of

Immobilized L-Alase

or

or Free L-AIase

D-galactose

D-tagatose Immobilized cells containing L-AIase

or cells

L-arabinose isomerase

agarose alginate beads chitopearl beads Glutaraldehyde and polyethylenimine

Fig. 4.4 The current strategies for the production of D-tagatose from D-galactose

300

Free L-AIase

Immobilized L-AIase in in calcium alginate

Thermoanaerobacter mathranii

Geobacillus stearothermophilus

Thermotoga neapolitana

Immobilized E. coli cells containing Gali 152 L-AIase mutant in sodium alginate Immobilized Escherichia coli cells in calcium alginate Beads Immobilized L-AIase in chitopearl bead

Immobilized E. coli cells containing L-AIase in glutaraldehyde and polyethylenimine Free L-AIase

100

18

10

180

300

1

300

100

Strategy Immobilized L-AIase using covalent binding to agarose Immobilized Gali152 L-AIase using alginate beads Free Gali152 L-AIase

Geobacillus stearothermophilus

Thermus sp. IM6501

Tan. mathranii

Geobacillus stearothermophilus Geobacillus stearothermophilus Tan. mathranii

Source Escherichia coli

Substrate concentration (g L 1) 500

pH 7.5, 75  C, 5 mM Mn2+

pH 8.0, 60  C, 62 h

70  C, 12 h

pH 8.0, 60  C, 5 mM Mn2+, 72 h pH 7.0, 70  C, Mn2+

pH 6.9, 65  C, 48 h,

Reaction condition pH 7.0, 30  C, 48 h pH 8.0, 60  C, 90 h pH 8.0, 60  C, 16 h pH 6.9, 65  C, 5 mM Mn2+

7.9

4.3

49

59

0.54

43.9

43

27.22

19.5

54

42

25

75

126

30.6

58.2

Conversion (%) 20.82

30.6

58.2

D-Tagatose (g L 1) 104.1

1.9

0.07

4.08

2.9

0.0075

2.63

–

1.91

0.65

Productivity (g L 1 h 1) 2.17

Table 4.2 Production of D-tagatose from D-galactose using L-arabinose isomerase from different substrates under various reaction conditions

Hong et al. (2007) Cheon et al. (2008) Liang et al. (2012)

Jung et al. (2005)

Reference Kim et al. (2001) Kim et al. (2003a) Kim et al. (2003a) Jorgensen et al. (2004) Jorgensen et al. (2004) Kim et al. (2003b)

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D-galactose to 145 g L 1 of D-tagatose and with the highest productivity of 54 g L 1 h 1 to date (Ryu et al. 2003). On the other hand, immobilized cells for the production of D-tagatose are another alternative choice, because the cells containing enzyme used for catalysts can help maintain the stability of intracellular enzymes and prevent damage to the enzymes by external environmental factors as much as possible (Wu et al. 2020). For example, the immobilized E. coli cells in sodium alginate containing L-AIase from G. stearothermophilus could produce an average of 59 g L 1 of D-tagatose from 300 g L 1 of D-galactose with a conversion rate of 19.5% and productivity of 2.9 g L 1 h 1 under the optimum conditions of 70  C and pH 7.0 and presence of Mn2+ (Jung et al. 2005). Coincidentally, if immobilizing the E. coli cells containing the L-AIase from Thermotoga neapolitana using calcium alginate beads, the cell reactor could produce 49 g L 1 of D-tagatose from 180 g L 1 of D-galactose within 12 h at 70  C in a continuous recycling mode (Hong et al. 2007). However, when using the same material to immobilize free L-AIase from Thermoanaerobacter mathranii, 7.9 g L 1 of D-tagatose was produced from 18 g L 1 of D-galactose under the optimal condition of pH 7.5, 75  C, and 5 mM Mn2+ (Liang et al. 2012). In addition to the abovementioned immobilization methods, the use of free enzymatic conversion to produce D-tagatose has also been investigated. The free L-AIase Gali 152 from G. stearothermophilus converted D-galactose into D-tagatose at a rate of 1.91 g L 1 h 1 and conversion rate of 30.6% at pH 8.0 and 60  C (Kim et al. 2003a). 75 g L 1 of D-tagatose was obtained from 300 g L 1 of D-galactose after using the free L-AIase from Tan. mathranii at pH 6.9, 65  C and 5 mM Mn2+, corresponding to 25% conversion (Jorgensen et al. 2004). Furthermore, using 1.0 g L 1 of D-galactose as substrate, the purified L-AIase from Thermus sp. could give a 54% conversion yield of D-galactose to D-tagatose at pH 8.0, 60  C for 3 days (Kim et al. 2003b).

4.7

Conclusion

The large-scale production of rare sugar is of great significance as broad application and huge benefits for people’s health are revealed. In this chapter, we have introduced the L-AIase which is an important aldose-ketose isomerase in the sugar industry. Because of the ability of converting the D-galactose to D-tagatose, L-AIase is considered to be a promising enzyme used for the production of rare sugar D-tagatose. The sources, biochemical properties about optimal temperature, pH, and metal ions are systematically summarized from the currently reported articles. Besides, the substrate specificity, kinetic parameters, and crystal structure of L-AIase are also presented. Finally, production of D-tagatose using different strategies such as immobilized enzymes, cells, or free enzymes is compared and analyzed. Especially in view of the deficiencies of the existing L-AIase’ enzymatic properties, molecular modifications have been carried out, such as lowering the optimal pH and increasing the catalytic activity toward D-galactose. Of course,

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more and more L-AIases with higher catalytic activity should be screened in the future’s work, and the molecular modification to the currently existing L-AIases should also be conducted based on the solved crystal structure.

References Adachi S (1958) Formation of lactulose and tagatose from lactose in strongly heated milk. Nature 181(4612):840–841 Ali MB, Mezghani M, Bejar S (1999) A thermostable α-amylase producing maltohexaose from a new isolated Bacillus sp. US100: study of activity and molecular cloning of the corresponding gene. Enzym Microb Technol 24(8):584–589. https://doi.org/10.1016/S0141-0229(98)00165-3 Belkin S, Wirsen CO, Jannasch HW (1986) A new sulfur-reducing, extremely thermophilic eubacterium from a submarine thermal vent. Appl Environ Microbiol 51(6):1180–1185. https://doi.org/10.1128/AEM.51.6.1180-1185.1986 Bond DR, Tsai BM, Russell JB (1998) The diversion of lactose carbon through the tagatose pathway reduces the intracellular fructose 1,6-bisphosphate and growth rate of Streptococcus bovis. Appl Microbiol Biotechnol 49(5):600–605 Chen Z, Chen J, Zhang W, Zhang T, Guang C, Mu W (2018) Recent research on the physiological functions, applications, and biotechnological production of d-allose. Appl Microbiol Biotechnol 102(10):4269–4278. https://doi.org/10.1007/s00253-018-8916-6 Cheng L, Mu W, Zhang T, Jiang B (2010) An L-arabinose isomerase from Acidothermus cellulolytics ATCC 43068: cloning, expression, purification, and characterization. Appl Microbiol Biotechnol 86(4):1089–1097. https://doi.org/10.1007/s00253-009-2322-z Cheon J, Kim SB, Park SW, Han JK, Kim P (2008) Comparative analysis of tagatose productivity of immobilized L-arabinose isomerase expressed in Escherichia coli and Bacillus subtilis. Food Sci Biotechnol 17:655–658 Choi JM, Lee YJ, Cao TP, Shin SM, Park MK, Lee H, Luccio ED, Kim SB, Lee SJ, Lee SJ (2016) Structure of the thermophilic l-Arabinose isomerase from Geobacillus kaustophilus reveals metal-mediated intersubunit interactions for activity and thermostability. Arch Biochem Biophys 596:51–62 Chouayekh H, Bejar W, Rhimi M, Jelleli K, Mseddi M, Bejar S (2007) Characterization of an L-arabinose isomerase from the Lactobacillus plantarum NC8 strain showing pronounced stability at acidic pH. FEMS Microbiol Lett 277(2):260–267 Donner T, Magder LS, Zarbalian K (2010) Dietary supplementation with d-tagatose in subjects with type 2 diabetes leads to weight loss and raises high-density lipoprotein cholesterol. Nutr Res 30 (12):801–806 Du M, Zhao D, Cheng S, Sun D, Chen M, Gao Z, Zhang C (2019) Towards efficient enzymatic conversion of D-galactose to D-tagatose: purification and characterization of L-arabinose isomerase from Lactobacillus brevis. Bioprocess Biosyst Eng 42(1):107–116. https://doi.org/ 10.1007/s00449-018-2018-9 Fan C, Liu K, Zhang T, Zhou L, Xue D, Jiang B, Mu W (2014) Biochemical characterization of a thermostable L-arabinose isomerase from a thermoacidophilic bacterium, Alicyclobacillus hesperidum URH17-3-68. J Mol Catal B Enzym 102:120–126. https://doi.org/10.1016/j. molcatb.2014.02.001 Heath EC, Horecker BL, Smyrniotis PZ, Takagi YP (1958) Pentose fermentation by Lactobacillus plantarum. II. L-arabinose isomerase. J Biol Chem 231(2):1031–1037 Hong Y, Lee DW, Lee S, Choe E, Kim S, Lee Y, Cheigh C, Pyun Y (2007) Production of D-tagatose at high temperatures using immobilized Escherichia coli cells expressing L-arabinose isomerase from Thermotoga neapolitana. Biotechnol Lett 29(4):569–574

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Chapter 5

Various Enzymes for the Biotechnological Production of D-Allose Ziwei Chen, Wenli Zhang, and Wanmeng Mu

5.1

Introduction

In recent years, more and more people are suffering from chronic diseases, such as obesity, hyperglycemia, and hypertension, due to excessive intake of high-sugar foods. Therefore, exploiting new functional sweeteners to substitute traditional table sugar (sucrose) has become very urgent, and due to their distinct health benefits, rare sugars have caused the attention of researchers. D-Allose is a momentous rare monosaccharide and sparsely distributed in the natural environment. D-Allose has 80% sweetness of table sugar but exhibits ultralow energy and nontoxic characteristics (Mooradian et al. 2017). D-Allose has been confirmed to have multitudinous health benefits and physiological functions such as antitumor, anti-inflammatory, cryoprotective, anti-osteoporotic, anti-hypertensive, neuroprotective, and antioxidative properties and could also be used as immunosuppressant (Chen et al. 2018a). D-Allose is an ideal table sugar substitute and promises a huge application potential in different fields, including food engineering, clinical treatment, and health care. However, the limited amount of D-allose heavily hinders its application. It is unrealistic to extract large amounts of D-allose from natural resources, due to low efficiency, high cost, and environmental damage. On the other hand, the chemical

Z. Chen · W. Zhang (*) State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China e-mail: [email protected] W. Mu State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, Jiangsu, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_5

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synthesis of D-allose is very inefficient and has many disadvantages, including low conversion rate, toxic by-product formation, and environmental pollution. In 2006, an Izumoring strategy was developed by Prof. Izumori (Rare Sugar Research Centre, Kagawa University, Japan), which could effectively realize the biological conversion between all hexoses (Izumori 2006). Based on this strategy, D-allose could be directly synthesized from D-allulose, another expensive rare sugar (originally known as D-psicose), through one-step conversion using various aldoseketose isomerases, mainly including L-rhamnose isomerase (EC 5.3.1.14), galactose-6-phosphate isomerase (EC 5.3.1.26), glucose-6-phosphate isomerase, and ribose-5-phosphate isomerase (EC 5.3.1.6). In order to reduce production costs of D-allose, the substrate D-allulose can be efficiently produced from D-fructose, a renewable and low-cost feedstock, using ketose 3-epimerase (EC 5.1.3.31). In this chapter, the physicochemical properties, metabolism pathway, physiological functions, biotechnological production, and miscellaneous applications of D-allose are presented in detail.

5.2 5.2.1

Conspectus of D-Allose Physicochemical Properties

D-allose is a D-form aldohexose and structurally equivalent to the C-3 epimer of D-glucose with a molecular formula of C6H12O6. The molecular weight and melting temperature of D-allose are 180.16 g/mol and 128  C, respectively. D-Allose exhibits a high water solubility and is insoluble in ethanol. Highly purified D-allose behaves as nontoxic and tasteless white powder (Iga et al. 2010). In aqueous solutions, D-allose exists in the conformation of an α β-D-pyranose ring in 4C1 (Kozakai et al. 2015). However, the D-allose molecular exists in four types of ring structure in dimethyl sulfoxide solutions, including α-D-allose-1,4 furanose, αD-allose-1,5-pyranose, β-D-allose-1,4-furanose, and β-D-allose-1,5-pyranose, with the percentages of 3.5%, 5%, 14%, and 77.5%, respectively (Angyal 1994; Kpper and Freimund 2003).

5.2.2

Occurrence in Plants

D-Allose has been extracted from plants in very small amounts, including Protea rubropilosa beard (Perold et al. 1973), Mentzelia (Jensen et al. 1981), Veronica filiformis (Chari et al. 1981), potato leaves (Weckwerth et al. 2004), and the African shrub Protea rubropilosa (O’Neil et al. 2006). Additionally, D-allose has also been isolated from the antibacterial Indian seagrasses Halodule pinifolia (Kannan et al. 2012) with a yield of 3.67%. Recently, D-allose has been detected in Acalypha hispida leaves at a ratio of 4.45% (Sithara et al. 2017).

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Metabolism Pathway

D-Allose, as a single carbon source, can be consumed by Escherichia coli K-12. The utilization of D-allose in E. coli K-12 needs a minimum of two operons. One of the operons is alsI, which encodes the AlsI protein. Another operon, composed of six genes (alsRBACEK) that encode six proteins (AlsRBACEK), regulates D-allose metabolism, and it can be induced by D-allose and repressed by the alsR gene and catabolite (Poulsen and Ying-Ying 1999). A transport system composed of three proteins, including ATP-binding component (AlsA), D-allose-binding protein (AlsB), and transmembrane protein (AlsC), is responsible for the transport of D-allose through the cellular membrane into the cytoplasm. AlsB, a periplasmic protein, is responsible for binding D-allose into the cytoplasm, due to its affinity. In cytoplasm, D-allose is converted linearly and orderly into D-allose-6-phosphate, D-allulose-6-phosphate, and D-fructose-6-phosphate utilizing, hypothetically, D-allose kinase (AlsK), D-allose-6-phosphate isomerase (AlsI), and D-allulose-6phosphate 3-epimerase (AlsE), respectively. D-Fructose-6-phosphate enters the EMP pathway to participate in metabolism. Moreover, it was found that AlsE is essential, while AlsK is dispensable in the D-allose metabolism pathway (Kim et al. 1997). The genetic regulation and metabolism pathway were shown in Fig. 5.1.

5.2.4

Chemical Preparation

D-Allose was chemically prepared using ribose derived from nucleic acid as starting material, involving the following procedures: cyanohydrin reaction, recrystallization, reduction with sodium amalgam, and purification with alcohol (Phelps and Bates 1934; Baker et al. 1972). D-Allose could be synthesized by the reduction of 3-ketosucrose with Raney nickel, and the corresponding by-products were eliminated by bacterial fermentation (Bernaerts et al. 1963). Additionally, D-allose could also be produced by the C-3 epimerization reaction of D-glucose using molybdenum as a catalyst and then subjected to concentration, decolorization, deionization, and separation (Herber et al. 1995). However, these chemical synthesis methods involve complicated steps and have many disadvantages, including low transformation rate, low selectivity, by-product formation, and chemical pollution. Therefore, the enzymatic production of D-allose has been drawn much attention in recent years.

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Fig. 5.1 The genetic regulation (red dashed box) and metabolism pathway of D-allose in E. coli. AlsA ATP-binding component, AlsB D-allose-binding protein, AlsC transmembrane protein, AlsK D-allose kinase, AlsI D-allose-6-phosphate isomerase, AlsE D-allulose-6-phosphate 3-epimerase

5.3 5.3.1

Physiological and Healthy Functions Anticancer and Antitumor

D-Allose possesses prominent anticancer and antitumor characters, which has attracted the attention of many researchers in recent years. So far, D-allose has been verified to restrain the proliferation and metastasis cancer and tumor cells which occurs in different organs or tissues, including the ovary (Sui et al. 2005a), pancreas (Malm et al. 2015), liver (Yokohira et al. 2008; Yamaguchi et al. 2008), prostate (Jeong et al. 2011; Naha et al. 2009), oral cavity (Indo et al. 2014), uterus, and skin (Sui et al. 2005b). In 2008, the underlying molecular action mechanism of D-allose-inhibited cancer cell proliferation was revealed by the investigation of HuH-7 hepatocellular carcinoma cells treated with D-allose for 48 h. On the one hand, D-allose distinctly upregulated the overexpression of thioredoxin-interacting protein (TXNIP), which can induce G1 cell cycle arrest instead of apoptosis and is usually inhibited in tumor cells. On the other hand, D-allose significantly improves the level of p27kip1 protein,

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which is a pivotal regulator of G1/S cell cycle transition. After D-allose treatment, the stabilization and localization of p27kip1 protein was apparently observed, suggesting that D-allose suppresses cancer cells growth by distinct TXNIP induction and p27kip1 protein stabilization (Yamaguchi et al. 2008). Later on, Hoshikawa et al. verified this upregulation of TXNIP action mechanism by in vitro and in vivo experiments using head and neck cancer cell lines (Hoshikawa et al. 2011). More recently, Noguchi et al. have further studied the inhibition cancer cell mechanism of D-allose. The growth restriction deregulation of cancer cells demands a multitude of glucose as a major energy source which are transported into cells by glucose transporters (GLUTs). Western blot and real-time PCR results showed that D-allose treatment not only increases the expression of TXNIP protein but also suppresses the expression of GLUT1 protein in a dose-dependent way in three cancer cell lines (hepatocellular carcinoma, adenocarcinoma, and Caucasian breast neuroblastoma), suggesting that D-allose may inhibit cancer cells growth by inhibiting GLUT1 expression and glucose absorption (Noguchi et al. 2016). D-Allose which inhibits cell proliferation by competing with glucose at the mitochondrial respiratory chain to reduce ATP synthesis has been investigated in neuroblastoma Neuro2A cells (Ishihara et al. 2011). Additionally, D-allose combined with other anticarcinogens has also been found to display more efficacious anticancer and antitumor effects than single D-allose treatment. In vivo experiments showed that docetaxel (an effective antitumor agent) combined with D-allose exerted stronger antitumor activity than either agent alone (Indo et al. 2014). Studies in pancreatic cancer models suggest that D-allose showed a promising synergistic effect with platinum agents (Malm et al. 2015). D-Allose can be used as a fortifier to improve the effectiveness of radiotherapy and chemotherapy toward tumor cells and a protectant to reduce the radiation-induced side effects, such as hyperkeratosis, tumor necrosis factor-α immunostaining, and epidermal thickening (Hoshikawa et al. 2018). In a word, these preclinical studies suggest that D-allose promises a tremendous application potentiality in the clinical treatment of tumors.

5.3.2

Antioxidant Properties

In 2003, D-allose was first proven to hold antioxidative properties by scavenging reactive oxygen species (ROS) and inhibiting the generation of ROS from stimulated neutrophils, suggesting that D-allose can ameliorate the negative effects of liver transplantation and ischemia/reperfusion (I/R) injury (Murata et al. 2003). In rat experiments, subjected to transitory middle cerebral artery occlusion (MCAO), D-allose significantly weakens behavioral deficits and brain damage and induces neuroprotection in focal cerebral ischemia by inhibiting ROS production (Nakamura et al. 2011). Additionally, the antioxidant action mechanism of D-allose was also partly investigated. In the cells, D-glucose induces ATP synthesis and promotes the generation of ROS. However, D-allose can inhibit ROS formation by competing with D-glucose in the mitochondria (Ishihara et al. 2011).

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Anti-inflammatory Effects

Based on previous studies that D-allose attenuates I/R injury by antioxidant effects, Gao et al. hypothesized that D-allose has the neuroprotection against cerebral I/R injury, possibly due to its anti-inflammatory properties (Gao et al. 2011). Subsequently, this hypothesis was verified in rat experiments, and the present results showed that D-allose has distinct anti-inflammatory effects and significantly contributes to beneficial effects (Gao et al. 2013). Recently, the effects of D-allose on the inflammatory reaction have been investigated in the gerbil after transient forebrain ischemia, suggesting that D-allose decreases brain damage by inhibiting inflammation response and oxidative stress (Shinohara et al. 2016). Mice fed the high-fat diet containing D-allose experiment showed that D-allose can ameliorate nonalcoholic steatohepatitis disease by inhibiting inflammation (Yamamoto et al. 2017). The potential anti-inflammatory mechanism of D-allose by inducing upregulation of peroxisome proliferator-activated receptor γ (PPARγ) was also indicated (Huang et al. 2016).

5.3.4

Other Health Benefits

Besides these physiological functions described above, other salutary functions of D-allose have also been studied. D-Allose was confirmed to be a potent inhibitor for the development of salt-induced high blood pressure, synchronously accompanied with the attenuation of superoxide production (Kimura et al. 2005). D-Allose exhibits beneficial cryoprotective effects on the cells of various tissues and organs, which is similar to that of trehalose (an acknowledged cryoprotectant) (Sui et al. 2007). D-Allose, as an immunosuppressant, has been proven effective to suppress immunological response in allogenic orthotopic liver transplantation in rats and has no detectable side effects (Hossain et al. 2000; Tanaka and Sakamoto 2011). Altogether, D-allose promises a great application potential in food, clinical medicine, and health-care fields. All conspicuous physiological functions and healthy benefits of D-allose are shown in Fig. 5.2.

5.4 5.4.1

Applications Application in the Food Industry

D-Allose is noncaloric sweetener but has 80% of the sweetness of table sugar (Mooradian et al. 2017). The acute and sub-chronic toxicity experiments of rats indicate that D-allose is a nontoxic monosaccharide and shows a different nutritive peculiarity from other monosaccharides (e.g., D-glucose and D-fructose) (Iga et al.

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Fig. 5.2 Recent researches on the distinct physiological functions of D-allose, including anticancer, cryoprotective, antiosteoporotic, antioxidative, neuroprotective, antihypertensive, antiinflammatory, antihypertensive, and immunosuppressant effects

2010). D-Allose can be used as food additive to substitute table sugar, which not only provides high sweetness but also is conducive to losing weight. Additionally, D-allose is a reducing sugar, and relevant studies have shown that compared with D-fructose or D-glucose, D-allose shows a faster reaction rate and higher crosslinking efficiency with α-lactalbumin in Maillard reactions, suggesting that D-allose may improve the color and flavor in during food processing (Sun et al. 2006). Its salutary physiological, chemical properties, and safety feature allows that D-allose has a enormous application prospect in the food industry.

5.4.2

Application in Clinical Medicine

Due to its versatile physiological properties, D-allose can be used as a potent therapeutic agent for the treatment of different diseases, such as cancer and tumors, obesity, hypertension, inflammation, and apoplexia (Gao et al. 2013). Due to its cryoprotective and immunosuppressant effects (Kashiwagi et al. 2016; Sui et al. 2007), the application of D-allose in surgery and organ transplantation to ameliorate tissue injury and improve the success rate of operation has been patented (US Patent No. 5620960, 1997). Due to its anti-inflammatory effects, D-allose can attenuate nephrotoxicity induced by cisplatin (an antineoplastic agent) by inhibiting serum TNF-alpha production (Miyawaki et al. 2012). Additionally, D-allose can be also used as an antioxidant to treat different diseases caused by oxidative stress (Nakamura et al. 2011; Ishihara et al. 2011).

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Application in Health Care

D-Allose can be potentially employed as a supplement ingredient in health products to prevent osteoporosis (Yamada et al. 2012). D-Allose has been reported to significantly enhance the action of metronidazole on trichomonad parasites Tritrichomonas foetus, suggesting that D-allose can decrease the drug dosage of metronidazole and prevent the development of drug resistance (Harada et al. 2012). The growth inhibition of D-allose on nematodes Caenorhabditis elegans has also been reported (Sakoguchi et al. 2016). Additionally, D-allose can also be potentially applied in crop disease prevention. The treatment of D-allose has been proven to be valid for reducing the development of rice diseases by triggering self-defense with ROS generation (Kano et al. 2013).

5.5 5.5.1

Various Enzymes for D-Allose Production L-Rhamnose Isomerase

L-Rhamnose isomerase (L-RIase, EC 5.3.1.14) is an aldose-ketose isomerase and plays the most important role in the biological production of D-allose. Due to its wide substrate specificity, L-RIase can catalyze the reversible isomerization not only between L-rhamnose and L-rhamnulose but also between D-allose and D-allulose (Xu et al. 2016). To date, D-allose-producing L-RIases have been characterized in different strains, including Pseudomonas stutzeri (Leang et al. 2004), Bacillus pallidus Y25 (Poonperm et al. 2007), Thermoanaerobacterium saccharolyticum NTOU1 (Lin et al. 2010), Thermotoga maritima ATCC 43589 (Park et al. 2010), Caldicellulosiruptor saccharolyticus ATCC 43494 (Lin et al. 2011), Mesorhizobium loti (Takata et al. 2011), Dictyoglomus turgidum DSMZ 6724 (Kim et al. 2013), Bacillus subtilis WB600 (Bai et al. 2015), Thermobacillus composti KWC4 (Xu et al. 2017), Clostridium stercorarium ATCC 35414 (Seo et al. 2017), and Caldicellulosiruptor obsidiansis OB47 (Chen et al. 2018b). These characterized L-RIases show a metal-dependent attribute, and their catalytic activities can be remarkably activated by Co2+ or Mn2+. L-RIases display prominent thermal adaptability and have high optimal reaction temperatures ranging from 60 to 85  C. Their optimal pH is at neutral or in an alkaline range of 7.5–9.0 (Xu et al. 2016). The majority of characterized L-RIases have a prominent thermal stability, which can improve enzyme utilization and is conducive to the consecutive production of D-allose. However, an excessively high reaction temperature or alkaline conditions will result in the Maillard reaction and unwanted by-product formation, which are disadvantages for the isolation and purification of D-allose. Thus, the reaction conditions, including pH, temperature, and thermal stability, should be considered together in the actual production of D-allose. Additionally, the kinetic parameters of different L-RIases have been largely investigated toward

5 Various Enzymes for the Biotechnological Production of D-Allose

93

D-allulose but not toward D-allose. The enzymatic properties of L-RIases are summarized in Table 5.1.

5.5.2

D-Ribose-5-Phosphate Isomerase

D-Ribose-5-phosphate isomerase (RPIase, EC 5.3.1.6) is a ubiquitous intracellular enzyme and plays an indispensable role in monosaccharide anabolism and catabolism. It can catalyze the interconversion of D-ribulose 5-phosphate and D-ribose 5-phosphate (Zhang et al. 2003). Previously, RPIase has been only found to participate in D-allose metabolism (Kim et al. 1997). Until 2007, the RPIase from Clostridium thermocellum was first used for the biological production of D-allose (Park et al. 2007a). Subsequently, three RPIases were characterized from various microorganisms, including Thermotoga lettingae TMO (Feng et al. 2013), Clostridium difficile ATCC BAA-1382D-5, and Thermotoga maritima ATCC 43589D-5 (Yeom et al. 2010). The RPIase from T. maritima ATCC 43589D-5 displays a relatively high catalytic activity (12 U/mg) toward D-allulose and a salient thermal stability (Half-life: 195 h, 75  C). Thus, this RPIase is preferential candidate for the application in the industrial production of D-allose.

5.5.3

Other D-Allose-Producing Enzymes

D-Galactose-6-phosphate isomerase (GaPIase, EC 5.3.1.26) is an essential enzyme in the metabolism pathway of D-tagatose and can isomerize D-tagatose-6 phosphate to galactose-6-phosphate (vice versa). However, only one GaPIase from Lactococcus lactis has been characterized to produce D-allose from D-allulose so far (Park et al. 2007b). The GaPIase from L. lactis exhibits relatively low optimum temperature (30  C) and specific activity toward D-allose (1.8 U/mg). Particularly, glucose-6-phosphate isomerase (GlPIase, EC 5.3.1.9) from Pyrococcus furiosus displays the highest catalytic activity (324 U/mg), optimal temperature (95  C), and thermal stability (half-life: 7.9 h, 95  C) toward D-allulose, compared to other reported D-allose-producing enzymes, which promise an enormous application potentiality for D-allose production (Yoon et al. 2009). Additionally, the mannose6-phosphate isomerase (MPI, EC 5.3.1.8) from Bacillus subtilis ATCC 23857 can also convert D-allulose to D-allose, due to its extensive substrate specificity, but displays an ultralow catalytic activity (Yeom et al. 2009). The properties of various D-allose-producing enzymes are compared in Table 5.1.

Enzymes L-RIase 8

7 7 8

85

65

70 75

60

90

75

85

65

60

T. composti KWC4

B. subtilis WB600 D. turgidum DSMZ 6724

M. loti

C. saccharolyticus ATCC 43494 T. saccharolyticum NTOU1 T. maritima

B. pallidus Y25

P. stutzeri

9

7

9

8 8

7.5

pH 7

Temp. ( C) 75

Strains C. stercorarium ATCC 35414 C. obsidiansis OB47

42

41.8

NR

121

14.3

7.11

2500

34.5

NR

33.9

68.1

1.33

0.74 81

2.46a

70.4a 5.98 61.5

11.25

kcat (s 1) 36.3a

25.8

Km (mM) 17.2

Table 5.1 Properties of various D-allose-producing enzymes

59.5

0.825

NR

0.28

4.77

0.19

0.12 1.3

0.035a

0.44

kcat/Km (Mm s 1) 2.11a

1

7.5

2.58

1.1a

5.7

21

3.03

NR NR

1.7a

13.7

Specificity activities (U/mg) 4.5a

NR

Half-life (h) 22.8 (65  C) 3.30 (85  C) 3.65 (65  C) 6.0 (65  C) 12.7 (80  C) >1 h (50  C) ~1 h (90  C) >2 h (75  C) 773 h (75  C) 1 h (65  C)

Poonperm et al. (2007) Leang et al. (2004)

Park et al. (2010)

Lin et al. (2010)

Takata et al. (2011) Lin et al. (2011)

Bai et al. (2015) Kim et al. (2013)

Chen et al. (2018b) Xu et al. (2017)

Reference Seo et al. (2017)

94 Z. Chen et al.

65 40 70

75 30 95

C. thermocellum C. difficile T. maritima

T. lettingae TMO L. lactis P. furiosus

8 7 7

7.5 7.5 8

2347 200 140 0.116a 0.0448 338.2

53 460 130 64a 58 214

0.2a 1.8 324a

1.8  10 7.7  10 0.63 4

3a

1.9a 45 12a

44 0.4 1.1

4.7 (65  C) 53 (55  C) 195 (75  C) 3.3 (75  C) NR 7.9 (95  C) Feng et al. (2013) Park et al. (2007b) Yoon et al. (2009)

Park et al. (2007a) Yeom et al. (2010) Yeom et al. (2010)

Substrate on D-allulose, otherwise on D-allose; NR not reported, L-RIase (L-rhamnose isomerase), RPIase (D-ribose-phosphate isomerase), GaPIase (D-galatcose-6-phosphate isomerase), GlPIase (glucose-6-phosphate isomerase)

a

GaPIase GlPIase

RPIase

5 Various Enzymes for the Biotechnological Production of D-Allose 95

96

5.6

Z. Chen et al.

Biological Production of D-Allose

According to Izumoring strategy, the production of D-allose requires D-allulose as its synthetic precursor of, which would result in a high production cost. To avoid this problem, an economical synthetic route using low-cost D-glucose or D-fructose as starting materials is preferentially selected to produce D-allose by two-step enzymatic reactions, involving ketose 3-epimerase and aldose-ketose isomerase (Fig. 5.3). In the first step, the rare sugar D-allulose is obtained from D-fructose using D-allulose 3-epimerase (DAEase) or D-tagatose 3-epimerase (DTEase) (Li et al. 2019; Zhang et al. 2016). Alternatively, D-glucose can be also used as starting material with additional D-glucose isomerase (GIase) in this step (Lim and Oh 2011). In the second step, D-allulose is isomerized into D-allose by L-RIase or other D-allose-producing enzymes. The final D-allose product is isolated and purified from the reaction mixture containing reaction feedstocks by a moving-bed chromatograph system and crystallization (Morimoto et al. 2006). The recombinant L-RIase from P. stutzeri, immobilized by cross-linking with glutaraldehyde, can convert 100 g D-allulose into 25 g D-allose, simultaneously accompanied with 8% undesired by-product D-altrose. This cross-linked L-RIase displays an effective catalytic half-life of approximately 2 months after repeated usages (Menavuvu et al. 2006). The D-allulose substrate was largely obtained from an inexpensive D-fructose by immobilized DTEase (Menavuvu et al. 2006). Additionally, 20,000 units of P. stutzeri L-RIase are immobilized on BCW-2510 chitopearl beads and convert D-allulose into D-allose with an approximate conversion rate of 30%, without significant reduction in the catalytic activity after 17 days. This large-scale production of D-allose was carried out in a continuous bioreactor using a simulated moving bed chromatograph separation system. After continuous separation, the D-allose solution was concentrated to approximately 50% and gradually crystallized at room temperature. Finally, 1.65 kg D-allose crystals with a purity of 100% were obtained, and the productivity of D-allose crystals from the D-allulose substrate was calculated to be approximately 10% (Morimoto et al. 2006). The L-RIase from B. pallidus Y25 synthesizes D-allose from D-allulose with an

Fig. 5.3 Enzymatic route for the conversion of D-glucose or D-fructose to D-allose using various enzymes, including D-glucose isomerase (GIase), D-tagatose 3-epimerase (DTEase), D-allulose 3-epimerase (DAEase), L-rhamnose isomerase (L-RIase), D-ribose-5-phosphate isomerase (RPIase), D-galactose-6-phosphate isomerase (GaPIase), and glucose-6-phosphate isomerase (GlPIase)

5 Various Enzymes for the Biotechnological Production of D-Allose

97

equilibrium rate of 35:65 after 48 h, without any by-product contaminations (Poonperm et al. 2007). A thermostable L-RIase from T. saccharolyticum NTOU1 catalyzes the isomerization of 100 mM D-allulose to D-allose, approaching equilibrium after 24 h, with a transformation rate of 29% and without any detectable by-products (Lin et al. 2010). Another thermostable L-RIase from C. saccharolyticus ATCC 43494 produces D-allose from 50 mM D-allulose with a transformation rate of 33% lacking of any by-products, and the reaction was reaching equilibrium after 0.5 h (Lin et al. 2011). Compared with other D-alloseproducing enzymes, the L-RIase from B. subtilis WB600 producing D-allose from D-allulose displays the highest transformation rate of 37.5% without any by-product formation when the reaction reaches equilibrium after 48 h (Bai et al. 2015). Recently, two novel thermostable L-RIases from T. composti KWC4 (Xu et al. 2017) and C. stercorarium ATCC 35414 (Seo et al. 2017) have been recombinantly expressed to produce 23 g/L and 199 g/L D-allose from D-allulose with transformation rates of 23% and 33%, respectively. More recently, D-allose has been effectively produced from low-cost substrate D-fructose (500 g/L) by a one-pot enzymatic process using the DAEase from Ruminococcus sp. and the L-RIase from Bacillus subtilis 168, which are immobilized on anion exchange resin D301 and amino Resin LX-1000EA, respectively (Can et al. 2020). After 5 h, the reaction was reached equilibrium state, and the mass ratios of D-allose, D-allulose, and D-fructose were determined to be 10%, 24%, and 66%, respectively (Can et al. 2020). This one-pot enzymatic process, in which D-allose is produced from renewable D-fructose using immobilized L-RIase combined with DAEase, is much simplified and low cost and thus displays greater application potential in the industrial production of D-allose. To sum up, L-RIases are paramount D-allose-producing enzymes, and majority of them are outstanding candidate for industrial production. The RPIase from C. thermocellum converts 500 g/L D-allulose into 165 g/L D-allose after 6 h reaction without detectable by-products (Park et al. 2007a). Improving catalytic activity and conversion rate of C. thermocellum RPIase toward D-allulose by through site-directed mutagenesis has also been investigated. A dozen active-site residues of C. thermocellum RPIase, which were confirmed by molecular modeling according to resolved crystal structure, were, respectively, replaced with Ala, and only the mutant R132A showed an improved catalytic activity toward D-allulose. When the active-site R132 was substituted with Glu, Ala, Asp, Ile, Gln or Lys, the mutant R132E displayed the highest activity and increased the conversion rate of D-allulose to D-allose by 7% compared to wild-type enzyme (Yeom et al. 2011). Recently, a one-pot enzymatic reaction, using a two-enzyme system composed of the RPIase from C. thermocellum and the DAEase from Flavonifractor plautii, has also been employed to produce 79 g/L D-allose from 600 g/L D-fructose with a transformation yield of 13% after 2 h (Lee et al. 2018). The RPIase from T. lettingae TMO produces approximately 32% D-allose from 100 mM D-allulose without detectable by-products when the conversion was approaching equilibrium after 4 h (Feng et al. 2013). Additionally, 2 g of dry E. coli cells harboring recombinant T. lettingae TMO RPIase can produce 28 g/L D-allose from 100 g/L D-allulose in a 1 L reaction system after 24 h (Feng et al. 2013). These RPIases can

98

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potently convert D-allulose into D-allose without any detectable by-products, suggesting that RPIases are ideal biocatalysts to apply in the large-scale production of D-allose. The GaPIase from Lactococcus lactis has the ability to convert 100 g/L D-allulose into 25 g/L D-allose but synchronously produce a fair amount of D-altrose (13 g/L) as a by-product after 12 h, although the conversion between D-allose and D-allulose reaction was faster than the conversion between D-altrose and D-allulose (Park et al. 2007b). The GlPIase from P. furiosus can convert D-allulose into 32% D-allose and also produce a small proportion of D-altrose (2%) when the reaction was approaching equilibrium state after 12 h (Yoon et al. 2009). Although the GlPIase from P. furiosus shows an excellent enzymatic property, such as high catalytic activity and prominent thermal stability, the formation of concomitant D-altrose by-product significantly hinders its application in industrial production of D-allose. The biological production of D-allose using various D-allose-producing enzymes is summarized in Table 5.2.

5.7

Conclusions and Future Perspectives

D-Allose displays multifarious physiological functions and biological activity, especially in anticancer effects. A detailed understanding on D-allose antitumor mechanism can facilitate its applications in clinical treatment. However, its whole antitumor mechanism has not been totally revealed so far, and thus more experiments are imminently needed to investigate the detailed anticancer mechanism of D-allose. Not only that, the elaborate mechanisms of other health benefits of D-allose should be also further investigated to expand its application prospect. Although these beneficial physiological functions of D-allose have been extensively investigated, they are only verified by animal experiments rather than clinical experiments in humans, and its metabolism pathway, health benefits, and safety on the human body have not been explored yet. Therefore, it is indispensable to carry out clinical tests in humans to form guide instructions in the practical applications, such as metabolic pathways, absorptive dosage, positive therapeutic effects, side effects, and food safety. To date, all of characterized D-allose-producing enzymes display optimal pH values within a range from neutral to slightly alkaline, which easily lead to the Maillard reaction at a high temperature. Additionally, the majority of them possess metal-dependent attributes, which are not conducive to the isolation and purification of D-allose and may result in health risk in the food application. Therefore, to meet the requirement of industrial application, the D-allose-producing enzymes holding acidic optimal pH and metal-independent properties should be exploited by novel enzyme scanning and rational design on basis of their resolved crystal structures. The immobilization of D-allose-producing enzymes can improve the repeated uses, and thus it is necessary to develop new immobilization materials. Contemplating the future, the directed evolution combined with high-throughput screening is

Free

0.18

0.058

25

28

5.76

NR

165

150

25

0.63

6.12

23 3.75 2.97

Product (g L 1) 199

32

25

28

32

32

33

30

25

35

29

23 37.5 33

Conversion ratio (%) 33

0.005

2.1

1.21

1.44

NR

27.5

0.37

2.1

0.013

1

1.53 0.17 5.9

Productivity (g L 1 h 1) 79.6

2

13

0

0

0

0

1.3

8

0

0

0 0 0

By-product ratio 0

Poonperm et al. (2007) Menavuvu et al. (2006) Morimoto et al. (2006) Park et al. (2007a) Yeom et al. (2011) Feng et al. (2013) Feng et al. (2013) Park et al. (2007b) Yoon et al. (2009)

(Lin et al. 2010)

Xu et al. (2017) Bai et al. (2015) Lin et al. (2011)

Reference Seo et al. (2017)

NR not reported, L-RIase (L-rhamnose isomerase), RPIase (D-ribose-phosphate isomerase), GaPIase (D-galatctose-6-phosphate isomerase), GlPIase (glucose6-phosphate isomerase)

P. furiosus

GlPIase

100

100

Cells

Free

18

Free

L. lactis

NR

500

Immobilized

Free

100

Immobilized

P. stutzeri

C. thermocellum R123E mutant T. lettingae TMO

1.8

Free

500

18

Free

Free

100 10 9

Free Free Free

C. thermocellum

Substrate (g L 1) 600

Biocatalyst Free

Strains C. stercorarium ATCC 35414 T. composti KWC4 B. subtilis WB600 C. saccharolyticus ATCC 43494 T. saccharolyticum NTOU1 B. pallidus Y25

GaPIase

RPIase

Enzymes L-RIase

Table 5.2 Biological production of D-allose from D-allulose using different enzymes

5 Various Enzymes for the Biotechnological Production of D-Allose 99

100

Z. Chen et al.

considered to be an effective approach to improve the catalytic activity and conversion rate toward D-allose. Additionally, the food-grade hosts, including Lactococcus lactis, Bacillus subtilis, and Saccharomyces cerevisiae, can be used for the recombinant expression of D-allose-producing enzymes to avoid the food safety issues resulted from pathogenic bacterium.

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Huang T, Gao D, Hei Y, Zhang X, Chen X, Fei Z (2016) D-allose protects the blood brain barrier through PPARγ-mediated anti-inflammatory pathway in the mice model of ischemia reperfusion injury. Brain Res 1642:478–486 Iga Y, Nakamichi K, Shirai Y (2010) Acute and sub-chronic toxicity of D-allose in rats. Biosci Biotechnol Biochem 74(7):1476–1478 Indo K, Hoshikawa H, Kamitori K, Yamaguchi F, Mori N (2014) Effects of D-allose in combination with docetaxel in human head and neck cancer cells. Int J Oncol 45(5):2044 Ishihara Y, Katayama K, Sakabe M, Kitamura M, Aizawa M, Takara M, Itoh K (2011) Antioxidant properties of rare sugar D-allose: effects on mitochondrial reactive oxygen species production in Neuro2A cells. J Biosci Bioeng 112(6):638–642 Izumori K (2006) Izumoring: a strategy for bioproduction of all hexoses. J Biotechnol 124 (4):717–722 Jensen SR, Mikkelsen CB, Nielsen BJ (1981) Iridoid mono-and di-glycosides in Mentzelia. Phytochemistry 20(1):71–83 Jeong RU, Lim S, Kim MO, Moon MH (2011) Effect of D-allose on prostate cancer cell lines: phospholipid profiling by nanoflow liquid chromatography-tandem mass spectrometry. Anal Bioanal Chem 401(2):689–698 Kannan RRR, Arumugam R, Anantharaman P (2012) Chemical composition and antibacterial activity of Indian seagrasses against urinary tract pathogens. Food Chem 135(4):2470–2473 Kano A, Fukumoto T, Ohtani K, Yoshihara A, Ohara T, Tajima S, Izumori K, Tanaka K, Ohkouchi T, Ishida YJ (2013) The rare sugar d-allose acts as a triggering molecule of rice defence via ROS generation. J Exp Bot 64(16):4939–4951 Kashiwagi H, Asano E, Noguchi C, Sui L, Hossain A, Akamoto S, Okano K, Tokuda M, Suzuki YJ (2016) Beneficial effect of D-allose for isolated islet culture prior to islet transplantation. JoH-BPS 23(1):37–42 Kim C, Song S, Park C (1997) The D-allose operon of Escherichia coli K-12. J Bacteriol 179 (24):7631. https://doi.org/10.1128/jb.179.24.7631-7637.1997 Kim YS, Shin KC, Lim YR, Oh DK (2013) Characterization of a recombinant L-rhamnose isomerase from Dictyoglomus turgidum and its application for L-rhamnulose production. Biotechnol Lett 35(2):259–264. https://doi.org/10.1007/s10529-012-1069-2 Kimura S, Zhang GX, Nishiyama A, Nagai Y, Nakagawa T, Miyanaka H, Fujisawa Y, Miyatake A, Nagai T, Tokuda M (2005) D-allose, an all-cis aldo-hexose, suppresses development of saltinduced hypertension in Dahl rats. J Hypertens 23(10):1887 Kozakai T, Fukada K, Kuwatori R, Ishii T, Senoo T, Izumori K (2015) Aqueous phase behavior of the rare monosaccharide D-allose and X-ray crystallographic analysis of D-allose dihydrate. Bull Chem Soc Jpn 88(3):465–470 Kpper S, Freimund S (2003) The Composition of keto aldoses in aqueous solution as determined by NMR spectroscopy. Helv Chim Acta 86(3):827–843 Leang K, Takada G, Fukai Y, Morimoto K, Granstrom TB, Izumori K (2004) Novel reactions of L-rhamnose isomerase from Pseudomonas stutzeri and its relation with D-xylose isomerase via substrate specificity. Biochim Biophys Acta 1674(1):68–77. https://doi.org/10.1016/j.bbagen. 2004.06.003 Lee T-E, Shin K-C, Oh D-K (2018) Biotransformation of fructose to allose by a one-pot reaction using Flavonifractor plautii D-allulose 3-epimerase and Clostridium thermocellum ribose 5-phosphate isomerase. J Microbiol Biotechnol 28(3):418–424 Li S, Chen Z, Zhang W, Guang C, Mu W (2019) Characterization of a D-tagatose 3-epimerase from Caballeronia fortuita and its application in rare sugar production. Int J Biol Macromol 138:536–545. https://doi.org/10.1016/j.ijbiomac.2019.07.112 Lim YR, Oh DK (2011) Microbial metabolism and biotechnological production of D-allose. Appl Microbiol Biotechnol 91(2):229–235 Lin CJ, Tseng WC, Fang TY (2011) Characterization of a thermophilic L-rhamnose isomerase from Caldicellulosiruptor saccharolyticus ATCC 43494. J Agric Food Chem 59(16):8702–8708. https://doi.org/10.1021/jf201428b

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Chapter 6

Characteristics of Cellobiose 2-Epimerase and Its Application in Enzymatic Production of Lactulose and Epilactose Qiuming Chen, Yaqin Xiao, and Yanchang Wu

6.1

Introduction

In 1967, Tyler et al. first found cellobiose 2-epimerase (CE, EC 5.1.3.11) in Ruminococcus albus ATCC 27210, an anaerobic ruminal bacterium, named for its ability to catalyze the epimerization of cellobiose to produce 4-O-β-Dglucopyranosyl-D-mannose (Tyler and Leatherwood 1967). However, cellobiose is not the optimal substrate for the catalytic reaction of CE. β-1,4-Linked mannobiose was reported to be the optimal substrate for CE (Park et al. 2013), which suggests that CE should be renamed as mannobiose 2-epimerase. CE can catalyze the interconversions of monosaccharides, disaccharides, and trisaccharides, which is the only identified epimerase currently known to work on unsubstituted disaccharides (Van Overtveldt et al. 2015). CE belongs to N-acyl-D-glucosamine 2-epimerase (AGE, EC 5.1.3.8) superfamily and has the same basic skeleton structure as the α6/α6-barrel in AGE, but none of them showed any AGE activities (Chen et al. 2018b). The CE from Ruminococcus albus can be used to catalyze the reversible epimerization of lactose to epilactose (Ito et al. 2008), which was the first biosynthetic pathway of epilactose. This discovery has drawn people’s attention to CE and greatly promoted the research on CE, because it is very difficult to obtain epilactose by chemical synthesis. Since then, more and more CEs from different microorganisms have been cloned and expressed. To meet possible requirements for industrial application, it is necessary to identify novel CEs from different microorganisms with different substrates preferences or CEs that can work at extreme conditions. At present, more than 20 types of CEs have been cloned and identified, as shown in the Table 6.1. In 2012, Kim et al. first reported that CE from Caldicellulosiruptor saccharolyticus can catalyze the isomerization of lactose to generate lactulose, which is also the first trial of enzymatic synthesis of lactulose Q. Chen (*) · Y. Xiao · Y. Wu State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_6

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Table 6.1 Summary of the reported CEs Name RhmaCE Dith-CE Casa-CE CaobCE Ditu-CE Thsa-CE Spth-CE Busp-CE Dyfe-CE Bafr-CE HeauCE Spli-CE Fiba-CE Roin-CE DygaCE Cele-CE CevuCE Euce-CE Fljo-CE Pehe-CE Sade-CE Tetu-CE Rual-CE

Strain for enzyme source Rhodothermus marinus JCM9785

GenBank No. BAK61777.1

Reference Ojima et al. (2011)

Dictyoglomus thermophilum Caldicellulosiruptor saccharolyticus Caldicellulosiruptor obsidiansis

ACI19378.1 WP_011915904.1 WP_013291422.1

Xiao et al. (2019b) Park et al. (2011) Chen et al. (2017)

Dictyoglomus turgidum Thermoanaerobacterium saccharolyticum Spirochaeta thermophila Butyrivibrio sp. AE2015 Dyadobacter fermentans Bacteroides fragilis Herpetosiphon aurantiacus

YP_002352551.1 AFK87331.1

Kim et al. (2012) Chen et al. (2015)

WP_013312912.1 WP_022776249 WP_015811450.1 BAH23773.1 ABX03535.1

Park et al. (2013) Kuschel et al. (2017a) Ojima et al. (2013) Senoura et al. (2009) Ojima et al. (2013)

Spirosoma linguale Firmicutes bacterium Roseburia intestinalis Dysgonomonas gadei

ADB36335.1 WP_02235242 RHN04075.1 WP_006797784.1

Cellulosilyticum lentocellum

WP_013658329.1

Cellvibrio vulgaris

–

Ojima et al. (2013) Kuschel et al. (2017a) Chen et al. (2020b) Krewinkel et al. (2015) Krewinkel et al. (2015) Saburi et al. (2015)

Eubacterium cellulosolvens Flavobacterium johnsoniae Pedobacter heparinus Saccharophagus degradans Teredinibacter turnerae Ruminococcus albus ATCC 27210

BAG68451.1 WP_012026921.1 WP_015809782.1 WP_011466993.1 WP_015818822.1 BAF81108.1

Taguchi et al. (2008) Ojima et al. (2013) Ojima et al. (2013) Ojima et al. (2013) Ojima et al. (2013) Ito et al. (2007)

using sole substrate source (Kim and Oh 2012). With such excellent catalytic function, CE shows a great potential for industrial application. Lactulose is a synthetic oligosaccharide with various excellent physiological functions. Although lactulose does not exist in nature, it has been studied extensively for a long time. Lactulose is a ketose isomer of lactose. In 1930 (Montgomery and Hudson 1930), lactulose was prepared from lactose in weak calcium hydroxide solution by the Lobry de Bruyn rearrangement. Lactulose crystal was obtained, and the physical properties were unraveled. About 30 years later, the excellent physiological function of lactulose was discovered and began to be used in food and medical treatment. Human small-intestinal lactase cannot hydrolyze lactulose (Dahlqvist and Gryboski 1965), so it doesn’t increase blood-reducing substances after eating. Lactulose can be used as a substitute for sucrose to help the people who

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suffers from obesity or type 2 diabetes, but it is more expensive. In 1957, lactulose had already been used in infant-feeding formulas. Lactulose can promote the development of intestinal bacteria, like Lactobacillus bifidus (Petuely 1957) Longcontinued heating of infant formulas results in considerable lactulose formation (Bernhart et al. 1965). Gerritsen et al. (Gerritsen et al. 1964) reported that the infant vomiting due to intestinal β-galactosidase activity deficiency can be cured by liquid milk containing two-thirds lactose and one-third lactulose in the total carbohydrate. Lactulose is widely used in the treatment of chronic constipation (Attar et al. 1999); compared with drastic laxatives, it has a gentle effect, without any toxic and systemic action forming (Wesselius-De Casparis et al. 1968). Lactulose (daily dose is 20.1 g) was proved to be safe for the treatment of chronic constipation in geriatric long-stay patients (Kinnunen et al. 1993). From infant to geriatric, lactulose has been shown to be both safe and effective. Lactulose is also used in health products to improve constipation in pregnant women. In 1966, lactulose was used to control systemic encephalopathy symptoms with great success (Müller et al. 1966). Since then, a growing number of clinical studies have used lactulose to alleviate and treat hepatic encephalopathy, especially in mild patients. Epilactose is a disaccharide that has many excellent physiological functions and is found in very small amounts in heated milk (Mu et al. 2013). Epilactose can be produced by the epimerization of lactose through chemical reactions. The chemical synthesis of epilactose is very difficult, because the by-products are complex and it is hard to obtain pure epilactose. As a result, the price of pure epilactose is extremely high, which limits the development of the research on epilactose. Ito et al. was the first to discover that cellobiose 2-epimerase from Ruminococcus albus NE1 could catalyze the epimerization of lactose to produce epilactose without generating any by-products (Ito et al. 2008). With the in-depth study of epilactose, more and more physiological functions of epilactose have been discovered. Such as prebiotics (Watanabe et al. 2008), anti-dental caries, and calcium absorption (Nishimukai et al. 2008). Unlike lactulose, which has decades of research history and mature industrial production, epilactose has not yet been put on the market. In the fixed reaction system, the conversion rate of lactose epimerization catalyzed by CE would not exceed 30% at equilibrium without other external means, such as continuous removal of products in the reaction. And the separation and purification of epilactose in the lactose solution is not easy and cheap. Improving the yield of epilactose by enzymatic production from lactose and simplifying the purification process (Saburi et al. 2010) is still in the experimental stage. Meanwhile, studies on the application of epilactose in food, medicine, and health products are being carried out (Krewinkel et al. 2014).

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Property of Lactulose and Epilactose General Physical and Chemical Properties

Lactulose (4-O-β-D-galactopyranosyl-D-fructose) is a polyhydroxy compound, which can be divided into one molecule of D-fructose and one molecule of D-galactose by breaking down the β-1,4-glucoside bond. Epilactose (4-O-β-Dgalactopyranosyl-D-mannose) is an epimer of lactose at the C-2 atom of the glucose moiety, which can be divided into one molecule of D-mannose and one molecule of D-galactose by breaking down the β-1,4-glucoside bond. The molecular formula and molecular weight of lactulose and epilactose are both C12H22O11 and 342.3 g mol 1. The structure formula of lactose, lactulose, and epilactose is shown in Fig. 6.1. Lactulose can exist in two different forms. It could be a clear viscous liquid with pale yellow or a stable white powdery solid with melting temperature at 169  C. Lactulose is a ketose isomer of lactose with a lower reducibility. Majid et al. heated milk ultrafiltration permeate with eggshell at 97  C to get lactulose (Nooshkam and Madadlou 2016). They approved that lactose is easier to cause mallard reaction than lactulose-rich product (about 70% lactulose content to total sugar). This is good news for industrial production with high temperature. Lactulose is easily soluble in the water, and the water solubility can reach 792 g L 1. Lactulose can be a sweetener that tastes cool, but the sweetness is about 0.6–0.8 of sucrose (Schumann 2002). Epilactose is a reducing oligosaccharide. Epilactose powder has a higher melting temperature than lactulose, but it has a lower water solubility of only 50 g L 1.

6.2.2

Physiological Effects of Lactulose and Epilactose

As sweetness, lactulose and epilactose do not cause tooth decay and dental plaque, because they won’t be used by the bacteria in mouth. Lactulose and epilactose are typical prebiotic which can resist digestion by gastric acid and the small intestine of the host. Taking them will not cause blood sugar to rise, which is very suitable for

Fig. 6.1 The molecular structures of lactose, lactulose, and epilactose

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Table 6.2 Comparison of properties and physiological effects of lactulose and epilactose CAS No. Molecular formula Molecular weight (g Mol 1) molomo111111111) (g/Mol) Melting temperature ( C) Water solubility (g L 1) Physiological effects

Reference

Lactulose 4618-18-2 C12H22O11 342.3

Epilactose 50468-56-9 C12H22O11 342.3

169

208

792

50

Prebiotic; preventing obesity, constipation, hepatic encephalopathy, salmonellosis, and colon cancer

Prebiotic; preventing obesity, metabolic disorders, arteriosclerosis, and colon cancer; promoting Ca absorption Watanabe et al. (2008), Mu et al. (2013) and Suzuki et al. (2010)

Chen et al. (2018b), Schumann (2002) and Panesar and Kumari (2011)

diabetics. Lactulose and epilactose both have positive effects on the large intestine, which can be used by Bifidobacterium and lactic acid bacteria to promote growth and fermentation (Moreno et al. 2014). Short-chain fatty acids (SCFAs) are the major metabolites produced in the fermentation. SCFAs are a series of mixtures, mainly composed of acetate, propionate, and butyrate (Shi et al. 2016). SCFAs play an important role in regulating colon function, which benefit the health of the host. SCFAs can be effectively absorbed by the epithelial cells that line the colonic lumen and provide energy for them. They also have functions of maintaining water and electrolyte balance in the intestine, protecting intestinal barrier, increasing the absorption of minerals, anti-inflammation (Puertollano et al. 2014) and modulating oxidative stress (Hamer et al. 2009). The proliferation of beneficial bacteria in the intestine can inhibit the survival of salmonella (Schumann 2002). SCFAs can also inhibit the proliferation, differentiation, and apoptosis of tumor cells, which are considered as a potential chemical defense agent in the adenoma-carcinoma sequence of the colorectum (Scheppach and Weiler 2004). SCFAs also have effect on treating metabolic diseases and inflammatory bowel disease (Huda-Faujan et al. 2010). Since SCFAs are the metabolic product of lactulose and epilactose fermented by Bifidobacterium and lactic acid bacteria in the colon, taking lactulose or epilactose has these benefits above. The comparison of properties and physiological effects of lactulose and epilactose are listed in Table 6.2. Intestinal microecology plays an important role in human health. Lactulose can be used not only to treat constipation (Cardelle-Cobas et al. 2011) and some intestinal diseases (Talley et al. 2011) but also to prevent and treat hepatic encephalopathy. Lactulose has been used as the first-line treatment for clinically significant and minor hepatic encephalopathy for decades (Gluud et al. 2013). Clinical trials show that lactulose can indeed lower the recurrence rate of hepatic encephalopathy (Courson

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et al. 2016). High levels of colonic secondary bile acids are thought to promote the development and progression of colon cancer (Farhana et al. 2016; Gadaleta et al. 2017). Lactulose can reduce the absorption of colonic secondary bile acids by acidifying colonic environment and rapid intestinal transport (van Berge Henegouwen et al. 1987). During long-term oral lactulose feeding, diarrhea and flatulence are the main adverse events. Lactulose can be given intravenously to induce systemic immune response that lowers endotoxin and tumor necrosis factor-α (TNF-α) (Greve et al. 1990). High levels of TNF-α are thought to promote necrosis of hepatocytes and liver inflammation. Research indicate that oral epilactose feeding reduces the sequelae of bone loss in rats after gastrectomy (Nishimukai et al. 2008). Epilactose can stimulate paracellular Ca absorption through the induction of myosin regulatory light chains phosphorylation via myosin light chain kinase- and Rho-associated kinase-dependent mechanisms (Suzuki et al. 2010). Ingestion of epilactose also increases the weight of the cecum wall and decreases the total cholesterol and low-density lipoprotein cholesterol in the plasma to reduce the risk of arteriosclerosis (Nishimukai et al. 2008). Uncoupling protein 1 is involved in the energy dissipation in skeletal muscle and brown adipose tissue, leading to increase energy consumption and decrease the risk of metabolic disorders. Epilactose can promote the expression of Uncoupling protein 1 (Murakami et al. 2015) to prevent obesity.

6.3

Property of Cellobiose 2-Epimerase

The specific activity of CEs from different microorganisms on the same substrate is obviously different, but the sequence of specific activity is basically the same when they catalyze different substrates. CE primarily catalyzes the epimerization and isomerization of disaccharides. Among the substrate specificity tested, the specific activity of CE was highest for mannobiose (Jaito et al. 2014; Senoura et al. 2011), followed by cellobiose and lactose. CE can also catalyze the interconversion of trisaccharides, which activity was much lower than disaccharides. The order of specific activity is mannotriose > maltotriose > cellotriose. CE can work on monosaccharides (Park et al. 2011), but the specific activities are extremely low, which are difficult to apply to actual industrial production. Substrates and products that can be catalyzed by cellobiose 2-epimerase are listed in Table 6.3. Temperature plays a very important role in catalytic function of CE, because temperature not only affects the enzyme activity of CE but also reflects the function of CE to some extent. The optimal temperatures of CEs are listed in Table 6.4. In previous research, all the CEs can catalyze epimerization, but only the part of them from thermophilic microorganisms can catalyze isomerization significantly, such as Dith-CE (Xiao et al. 2019b), Casa-CE (Kim and Oh 2012), Caob-CE (Chen et al. 2017), Ditu-CE (Kim et al. 2012), and Spth-CE (Park et al. 2013). The optimal temperatures for these CEs are high, and the production of lactulose can be detected within a few hours of reaction using lactose as the single substrate. The catalytic

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Table 6.3 Substrates and products that can be catalyzed by cellobiose 2-epimerase Substrate Monosaccharide D-Mannose

Product

Reference

D-Glucose, D-fructose

D-Glucose

D-Mannose

D-Xylose

D-Lyxose

D-Lyxose

D-Xylose

D-Fructose

D-Glucose, D-mannose

Park et al. (2013) Park et al. (2013) Park et al. (2011) Park et al. (2011) Park et al. (2011)

Disaccharide 4-O-α-D-GlucopyranosylD-mannose Maltose

Maltose 4-O-α-D-Glucopyranosyl-D-mannose

Epilactose

Lactose, lactulose

Lactose

Epilactose, lactulose

Lactulose

Epilactose, lactose

4-O-β-D-GlucopyranosylD-mannose Cellobiose

Cellobiose

Mannobiose

4-O-β-D-Mannopyranosyl-D-glucose

4-O-β-D-MannopyranosylD-glucose Trisaccharide Maltotriose

Mannobiose

Cellotriose Mannotriose

4-O-β-D-Glucopyranosyl-D- mannose

4-O-α-D-Glucopyranosyl-O-α-Dglucopyranosyl-D-mannose 4-O-β-D-Glucopyranosyl-O-β-Dglucopyranosyl-D-mannose 4-O-β-D-Mannopyranosyl-O-β-Dmannopyranosyl-D-glucose

Park et al. (2013) Park et al. (2013) Park et al. (2013) Park et al. (2013) Kim and Oh (2012) Park et al. (2013) Park et al. (2013) Park et al. (2013) Park et al. (2013) Park et al. (2013) Park et al. (2013) Park et al. (2013)

function of some CEs from mesophilic sources was investigated at a low reaction temperature in the extended reaction times from 8 to 21 days (Kuschel et al. 2017b). Rual-CE with the lowest optimal temperature could catalyze the isomerization of lactose to produce a small amount of lactulose. Other mesophilic CEs (Table 6.4) have also been shown to exhibit isomerization activities. Therefore, it is proposed that all CEs can catalyze both the epimerization and the isomerization reactions, but

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Table 6.4 Enzymatic properties of the reported CEs Name Rhma-CE Dith-CE Casa-CE Caob-CE Ditu-CE Thsa-CE Spth-CE Busp-CE Dyfe-CE Bafr-CE Heau-CE Spli-CE Fiba-CE Roin-CE Dyga-CE Cele-CE Cevu-CE Euce-CE Fljo-CE Pehe-CE Sade-CE Tetu-CE Rual-CE

Topt ( C) (Epimerization) 80 75 75 70 70 65 60 60 50 45 45 45 45 45 40 40 40 35 35 35 35 35 30

pHopt 6.3 7.0 7.5 7.5 7.0 7.0 7.0 7.0 7.7 7.5 7.3 7.7 7.5 7.0 7.5 8.0 8.0 7.5 8.4 6.3 7.7 8.8 7.5

Isomerization + ++ ++ ++ ++ NR + + + + NR NR + NR NR NR NR NR + NR NR NR +

Reference Ojima et al. (2011) Xiao et al. (2019b) Kim and Oh (2012) Chen et al. (2017) Kim et al. (2012) Chen et al. (2015) Park et al. (2013) Kuschel et al. (2017b) Kuschel et al. (2017b) Kuschel et al. (2017b) Ojima et al. (2013) Ojima et al. (2013) Kuschel et al. (2017b) Chen et al. (2020b) Krewinkel et al. (2015) Krewinkel et al. (2015) Saburi et al. (2015) Taguchi et al. (2008) Kuschel et al. (2017b) Ojima et al. (2013) Ojima et al. (2013) Ojima et al. (2013) Kuschel et al. (2017b)

++: high rate of isomerization reaction. +: low rate of isomerization reaction NR: not reported

the rate of CEs from mesophilic microorganisms which catalyzed isomerization was much lower than those from thermophilic microorganisms. CEs belong to the N-acetylglucosamine 2-epimerases superfamily, including several carbohydrate epi- and isomerases, e.g., N-acetylglucosamine 2-epimerases (AGE EC 5.1.3.8) and D-mannose isomerases (MI, EC 5.3.1.7). Although the amino acid sequences of most characterized CEs show only weak sequence identity with various MIs and AGEs (< 25%)(Chen et al. 2018b), the crystal structure from Rhodothermus marinus CE (Rhma-CE) deciphers that the three-dimensional structure has the same basic scaffold as the α6/α6-barrel of the catalytic domains of porcine kidney AGE (Pk-AGE) and Salmonella enterica MI (Saen-MI) (Fig. 6.2a). This barrel, composed of 12 α-helices, is characteristic for the six-hairpin enzyme superfamily in the SCOP database (scop.berkeley.edu) and the N-acylglucosamine 2-epimerase (GlcNAc 2-epimerase) family in the Pfam database (pfam.xfam.org). It seems that this common structure is suitable for the epimerization or isomerization of carbohydrates with mannosyl residues. Although the catalytic mechanism of this superfamily is still ambiguous, the binding pockets of CE, AGE, and MI are similarly located in the six inner helices

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Fig. 6.2 Crystal structure analysis. Three-dimensional macromolecular structures of Rhma-CE (PDB ID 3WKF), Pk-AGE (PDB ID 1FP3), and Saen-MI (PDB ID 2ZBL) (a) and their substrate binding pockets, highlighting the amino acid residues lining the pocket. (b) The substrate epilactose from Rhma-CE is displayed as sticks, and the C1 and C2 atoms of the reducing sugar end are labeled

of the α6/α6-barrel (Fig. 6.2b), indicating that the catalytic mechanisms of these three types of enzymes are probably similar. Two histidine residues in the binding pockets (His259 and His390 in Rhma-CE; Fig. 6.2b, yellow) are conserved in all the enzymes of this superfamily. They were postulated to proceed as general acid/base catalysts in deprotonation/reprotonation reactions for producing the epimerized product (Fujiwara et al. 2013, 2014). A third histidine residue (His200 in Rhma-CE, Fig. 6.2b, yellow), not conserved in the superfamily, was found to be required for the epimerization reaction catalyzed by CEs. It is in proximity to the C1 and C2 at the reducing end of the unmodified sugar (Fig. 6.2b).

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Enzymatic Production of Epilactose and Lactulose Enzymatic Production of Epilactose

At present, there is only one pathway for the biosynthesis of epilactose, which is obtained by using CE to catalyze the epimerization of lactose. The biotransformation method has the advantages of low cost, simple operation, and short time required for the reaction to reach equilibrium. Controlling the source of the enzyme and the amount of enzyme can achieve the effect without generating any by-products, which can simplify the purification steps (Xiao et al. 2019a). Up to now, CEs from 23 species have been cloned and identified to have epimerization. Since the behavior of epilactose is similar to lactose, the research emphasis focuses on the production and purification of epilactose and its applications in the food industry. Saburi et al. established a purification method for epilactose by four steps: crystallization, hydrolysis, digestion, and column chromatography. In following these steps, epilactose was recovered at 42.5% yield with 91.1% purity (Saburi et al. 2010). Sato et al. proved that immobilized CE can effectively reduce the amount of enzyme in the process of lactose processing (Sato et al. 2012). Kuschel et al. presented an HPLC protocol, which can separate epilactose from reaction mixture by ligand-exchange chromatography, and epilactose was recovered at 51% yield with 99% purity (Kuschel et al. 2016). Chen et al. constructed a food-grade enzymatic route for the production and purification of epilactose from lactose, which used Bacillus subtilis without antibiotic resistance genes as the enzyme expression host instead of E. coli. Epilactose with a purity >98% was separated from the reaction mixture by hydrolyzing lactose to monosaccharides, fermenting the yeast to consume the monosaccharides and cation-exchange chromatograph (Chen et al. 2018a). Some of the studies conducted experiments on CE to produce epilactose from milk under industrialized conditions to explore its industrial application value, which is listed in Table 6.5. Table 6.5 The production of epilactose from milk at 8  C Name DygaCE FljoCE PeheCE

pH 7.5

Lactose (g L 1) 48.3

Time (h) 10

Enzyme concentration (U mL 1) 1.30

Yield (g L 1) 29.9

7.0 7.0

42.5 45.5

24 24

0.48 0.84

33.6 30.5

Reference Krewinkel et al. (2015) Krewinkel et al. (2014) Krewinkel et al. (2014)

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6.4.2

115

Enzymatic Production of Lactulose

Enzymatic synthesis of lactulose is divided into two pathways. In the first path, lactulose can be produced from lactose and fructose with some commercial β-Dgalactohydrolase (β-galactosidase, EC 3.2.1.23) and β-glycosidase (EC 3.2.1.21). They can catalyze lactose hydrolysis (Neri et al. 2009; Petzelbauer et al. 2000) and transglycosidation (Hung and Lee 2002). In this reaction, lactose is hydrolyzed to produce galactose and glucose, and galactose is transferred to fructose via β-1,4glucoside bonds to form lactulose (Lee et al. 2004). The disadvantages are also very clear that several other oligosaccharides of different connection types are produced in the reaction, which increases the difficulty of purification and reduces the yield of lactulose. Most of the fructose on the market is obtained by enzymatic conversion using glucose as a substrate, which increases the cost. Song et al. found that lactulose can be produced from a single substrate, lactose, when β-galactosidase and βglycosidase were used together (Yoon-Seok et al. 2013). Table 6.6 lists some research on the production of lactulose through transglycosidation. Lactulose can also be obtained in one-step reaction with single substrate lactose and single by-product epilactose. CE catalyzes the isomerization of glucose on the lactose to fructose; conversion levels of lactulose reached 55–58% at equilibrium (Chen et al. 2017). But the isomerization generally requires longer reaction time, higher enzyme concentration, and higher reaction temperature than epimerization. Table 6.7 can reflect this rule, but its mechanism is still not clear. Research shows that borate can be used as cofactor to increase the yield of lactulose. Kim et al. used lactose and borate with the molar ratio of 1:1 to generate lactulose by CE from Caldicellulosiruptor saccharolyticus, which increases the conversion rate from 58% to 88% (Kim and Oh 2012; Park et al. 2013). This method is currently the highest conversion rate achieved by chemical and biological synthesis. However, borate makes the purification more difficult and may cause food safety problem. In 2015, Rentschler et al. first examined the application of CE for lactulose production in milk system at 50  C and 8  C; the result shows that CE was applicable in milk and low-temperature environment. Temperature has a significant effect on the time to reach the reaction equilibrium. The conversion of milk lactose resulted in a final yield of 57.7% lactulose at 50  C after 24 h and 56.7% lactulose at 8  C after 72 h (Rentschler et al. 2015). Since the isomerization activity of CE is slow, and current studies have shown that CE derived from thermophilic microorganisms naturally has a higher reaction rate. Therefore, many studies have started from finding new sources of CE with high thermal stability or molecular modification to improve the isomerization activity of CE. Shen et al. obtained a mutant G4-C5 after four sequential rounds of random mutagenesis and screening, which showed 2.8-fold increases in isomerization activity than wild type of Caldicellulosiruptor saccharolyticus (Shen et al. 2015). Xiao et al. verified the feasibility to discover novel thermostable enzymes by using molecular dynamics simulation as the preliminary computational filter. The CE obtained from Dictyoglomus thermophilum was one of the most thermostable CEs among all the reported CEs, with the highest epimerization

Condition Free Immobilized Free Immobilized Cell Immobilized

Strain for enzyme source Pyrococcus furiosus

Aspergillus oryzae

Kluyveromyces lactis Kluyveromyces lactis ATCC 8585 Streptomyces rubiginosus and Kluyveromyces lactis

Enzyme β-Glycosidase

β-Galactosidase

β-Glycosidase and βGalactosidase

Table 6.6 Production of lactulose through transglycosidation

200 400 200

200 200 --

Substrate (g L 1) Lactose Fructose 34.2 270 34.2 270 200 150 19.2 20 7.68

Yield of lactulose (g L 1) 16.3 16.7 65

Reference Mayer et al. (2004) Mayer et al. (2004) Adamczak et al. (2009) Song et al. (2013) Lee et al. (2004) Yoon-Seok et al. (2013)

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Table 6.7 Comparison of isomerization and epimerization

Name DithCE CaobCE ThsaCE

Substrate lactose (g L 1) 68.4

Enzyme concentration (U mL 1) 80

T ( C) 80

Lactulose Yield Time (g L 1) (h) 38.3 5

Epilactose Yield Time (g L 1) (min) 19.2 10

200

120

75

108

4

27.5

10

68.4

2.87

60

--

--

19.9

60

Reference Xiao et al. (2019b) Chen et al. (2017) Chen et al. (2015)

(160  6.5 U mg 1) and isomerization activities (3.52  0.23 U mg 1) (Xiao et al. 2019b). Molecular dynamics simulation can also be used to predict the thermostability changes of the mutated enzymes, which proved to be better than using staticstate information (Chen et al. 2020a).

6.5

Enzyme Engineering

According to different strategies, the molecular modification of enzymes can be divided into three categories: rational design, semi-rational design, and directed evolution. The rational design strategy is to carry out theoretical analysis on the sequence and structure information of the target protein, and site-directed mutagenesis is used to select certain sites or regions of the enzyme for modification. Park et al. (2017) analyzed the crystal structure information of the complex of Casa-CE and the substrate. They selected amino acid residues (Tyr114 and Asn184) that are more tightly bound to the mannosyl C2 atom of epilactose for site-directed mutagenesis to reduce the binding energy of epilactose. As a result, the epimeric activity of the Casa-CE single-point mutant Y114E was inhibited, while the isomeric activity increased accordingly. Within 2 h, the reaction using mutant Y114E can produce 86.9 g/L lactulose and 4.6 g/L epilactose at 200 g/L lactose. By introducing two amino acid residues in Casa-CE into Rhma-CE (S99M/Q371F), the reaction constant kcat of Rhma-CE toward glucose can be increased by two times. Directed evolution strategy is based on random mutation and high-throughput screening technology. Shen et al. (2016) used directed evolution strategy to improve the catalytic property of Casa-CE. A 5-point mutant enzyme G4-C5 was screened by this method. The structural isomerase activity of the mutant enzyme was increased by 2.8 times compared with wild-type Casa-CE, and the conversion rate of lactose to lactulose was increased from 56% to 76%. The lactulose production capacity of CE has been greatly improved.

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The concept of semi-rational design combines the ideas from the above two strategies. According to the related principles of protein, the mutation plan with better predicted effect is selected, the quality of the mutant library is improved, and the size of the mutant library is reduced. The target enzyme is obtained through experimental screening. A semi-rational design method was implemented to constructed mutants. The stability of a double-point mutant E161D/N365P obtained by combining single-point mutants has been greatly improved, and the half-life at 80  C has increased by four times (Shen et al. 2015).

6.6

Outlook and Perspectives

The balance of colonic microbiota is the key in the modulation of human immunity, metabolism, and endocrine activities. Prebiotics have demonstrated clear effects on colonic microbiota. Functional sugars can reduce the use of high-calorie sweet compounds such as sucrose, reducing the risk of diseases such as diabetes. Healthy diet strategies require a combination of sound nutritional evidence and technical tools with legal responsibility. Functional foods containing prebiotics have high value in this regard. Epilactose and lactulose are two prebiotics and functional disaccharide, which can be catalyzed from lactose by CE. The substrate lactose is a cheap disaccharide, naturally present in the milk of most mammals, and is the by-product of the dairy industry. Epilactose and lactulose are expensive and rely on artificial synthesis, which makes CE attractive for its irreplaceable functions. The number of publications about CE in the last decade reflects its broad prospects. The enzymatic reaction does not require additional substrates or lengthy steps and has the advantages of high efficiency and environmental protection. Biosynthesis will become the main trend in the production of functional food materials in the future. With the further study of CE, a comprehensive understanding of the relationship between the structure and function of CE and the mechanism of enzymatic catalysis, it is believed that the industry of lactose derivatives will take on a new look.

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Saburi W, Yamamoto T, Taguchi H, Hamada S, Matsui H (2010) Practical preparation of epilactose produced with cellobiose 2-epimerase from Ruminococcus albus NE1. Biosci Biotechnol Biochem 74(8):1736–1737. https://doi.org/10.1271/bbb.100353 Saburi W, Tanaka Y, Muto H, Inoue S, Odaka R, Nishimoto M, Kitaoka M, Mori H (2015) Functional reassignment of Cellvibrio vulgaris EpiA to cellobiose 2-epimerase and an evaluation of the biochemical functions of the 4-O-β-D-mannosyl-D-glucose phosphorylase-like protein, UnkA. Biosci Biotechnol Biochem 79(6):969–977. https://doi.org/10.1080/09168451. 2015.1012146 Sato H, Saburi W, Ojima T, Taguchi H, Mori H, Matsui H (2012) Immobilization of a thermostable cellobiose 2-epimerase from Rhodothermus marinus JCM9785 and continuous production of epilactose. Biosci Biotechnol Biochem 76(8):1584–1587. https://doi.org/10.1271/bbb.120284 Scheppach W, Weiler F (2004) The butyrate story: old wine in new bottles? Curr Opin Clin Nutr Metab Care 7(5):563–567 Schumann C (2002) Medical, nutritional and technological properties of lactulose. An update. Eur J Nutr 41(0):1–1. https://doi.org/10.1007/s00394-002-1103-6 Senoura T, Taguchi H, Ito S, Hamada S, Matsui H, Fukiya S, Yokota A, Watanabe J, Wasaki J, Ito S (2009) Identification of the cellobiose 2-epimerase gene in the genome of Bacteroides fragilis NCTC 9343. Biosci Biotechnol Biochem 73(2):400–406. https://doi.org/10.1271/bbb.80691 Senoura T, Ito S, Taguchi H, Higa M, Hamada S, Matsui H, Ozawa T, Jin S, Watanabe J, Wasaki J, Ito S (2011) New microbial mannan catabolic pathway that involves a novel mannosylglucose phosphorylase. Biochem Biophys Res Commun 408(4):701–706. https://doi.org/10.1016/j. bbrc.2011.04.095 Shen Q, Zhang Y, Yang R, Hua X, Zhang W, Zhao W (2015) Thermostability enhancement of cellobiose 2-epimerase from Caldicellulosiruptor saccharolyticus by site-directed mutagenesis. J Mol Catal B Enzym 120:158–164. https://doi.org/10.1016/j.molcatb.2015.07.007 Shen Q, Zhang Y, Yang R, Pan S, Dong J, Fan Y, Han L (2016) Enhancement of isomerization activity and lactulose production of cellobiose 2-epimerase from Caldicellulosiruptor saccharolyticus. Food Chem 207:60–67. https://doi.org/10.1016/j.foodchem.2016.02.067 Shi Y, Chen Q, Huang Y, Ni L, Liu J, Jiang J, Li N (2016) Function and clinical implications of short-chain fatty acids in patients with mixed refractory constipation. Colorectal Dis 18 (8):803–810. https://doi.org/10.1111/codi.13314 Song YS, Lee HU, Park C, Kim SW (2013) Batch and continuous synthesis of lactulose from whey lactose by immobilized β-galactosidase. Food Chem 136(2):689–694 Suzuki T, Nishimukai M, Takechi M, Taguchi H, Hamada S, Yokota A, Ito S, Hara H, Matsui H (2010) The nondigestible disaccharide epilactose increases paracellular Ca absorption via rho-associated kinase- and myosin light chain kinase-dependent mechanisms in rat small intestines. J Agric Food Chem 58(3):1927–1932. https://doi.org/10.1021/jf9035063 Taguchi H, Senoura T, Hamada S, Matsui H, Kobayashi Y, Watanabe J, Wasaki J, Ito S (2008) Cloning and sequencing of the gene for cellobiose 2-epimerase from a ruminal strain of Eubacterium cellulosolvens. FEMS Microbiol Lett 287(1):34–40. https://doi.org/10.1111/j. 1574-6968.2008.01281.x Talley NJ, Abreu MT, Achkar JP, Bernstein CN, Dubinsky MC, Hanauer SB, Kane SV, Sandborn WJ, Ullman TA, Moayyedi P, American College of Gastroenterology IBDTF (2011) An evidence-based systematic review on medical therapies for inflammatory bowel disease. Am J Gastroenterol 106(Suppl 1):S2–S25. https://doi.org/10.1038/ajg.2011.58; quiz S26 Tyler TR, Leatherwood JM (1967) Epimerization of disaccharides by enzyme preparations from Ruminococcus albus. Arch Biochem Biophys 119(1):363–367. https://doi.org/10.1016/00039861(67)90466-3 van Berge Henegouwen GP, Van der Werf SD, Ruben ATH (1987) Effect of long term lactulose ingestion on secondary bile salt metabolism in man: potential protective effect of lactulose in colonic carcinogenesis. Gut 28(6):675–680 Van Overtveldt S, Verhaeghe T, Joosten HJ, van den Bergh T, Beerens K, Desmet T (2015) A structural classification of carbohydrate epimerases: from mechanistic insights to practical

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applications. Biotechnol Adv 33(8):1814–1828. https://doi.org/10.1016/j.biotechadv.2015.10. 010 Watanabe J, Nishimukai M, Taguchi H, Senoura T, Hamada S, Matsui H, Yamamoto T, Wasaki J, Hara H, Ito S (2008) Prebiotic properties of epilactose. J Dairy Sci 91(12):4518–4526. https:// doi.org/10.3168/jds.2008-1367 Wesselius-De Casparis A, Braadbaart S, Bergh-Bohlken GE, Mimica M (1968) Treatment of chronic constipation with lactulose syrup: results of a double-blind study. Gut 9(1):84 Xiao Y, Chen Q, Guang C, Zhang W, Mu W (2019a) An overview on biological production of functional lactose derivatives. Appl Microbiol Biotechnol 103(9):3683–3691. https://doi.org/ 10.1007/s00253-019-09755-6 Xiao Y, Chen Q, Shakhnovich EI, Zhang W, Mu W (2019b) Simulation-guided enzyme discovery: a new microbial source of cellobiose 2-epimerase. Int J Biol Macromol 139:1002–1008. https:// doi.org/10.1016/j.ijbiomac.2019.08.075 Yoon-Seok S, Lee H-U, Park C, Kim S-W (2013) Optimization of lactulose synthesis from whey lactose by immobilized β-galactosidase and glucose isomerase. Carbohydr Res 369:1–5

Chapter 7

Enzymatic Production of Lactosucrose by Levansucrase, β-Fructofuranosidase, and β-Galactosidase Wei Xu, Wenli Zhang, and Hao Wu

7.1

Introduction

In nature, sucrose (O-α-D-glucopyranosyl-(1, 2)-β-D-fructofuranoside) and lactose (O-β-D-galactopyranosyl-(1,4)-D-glucopyranose) are two abundant and cheap disaccharides (Fewkes et al. 1971). For sucrose, it is also well-known as “table sugar” that is the most common sweetener consumed in our daily life. However, an excess ingestion of sucrose would definitely damage our teeth and increase our weight; even worse, this might result in some health risks, including type 2 diabetes and heart disease (Baker 2003). For the lactose, it is largely generated as a by-product in the dairy industry; but an inappropriate disposal of lactose would cause severe environmental pollution (Marwaha and Kennedy 1988). However, every coin has two sides. Both sucrose and lactose can serve as significant raw materials by biological transformation into other high value-added and functional derivatives. The sucrose functional derivatives primarily include its isomers, like trehalulose, turanose, leucrose, and isomaltulose, and its polymers like fructooligosaccharides (FOS), β-fructan, and α-glucan (Tian et al. 2019). Functional lactose derivatives include epilactose, lactulose, galactooligosaccharide (GOS), D-tagatose, lactobionic acid, and lactitol (Fig. 7.1) (Xiao et al. 2019). Notably, when sucrose and lactose are provided as substrates in the reaction system, a novel and bioactive trisaccharide, lactosucrose, could be synthesized. Lactosucrose is recognized as a kind of rare sugar since it does not occur largely in nature and is very hard to be chemically produced. The molecular formula and molecular weight of lactosucrose (CAS No. 87419-56-5) are C18H32O16 and 504.4 g/mol, respectively. The melting temperature of lactosucrose was about 181
C, and the specific rotation ([α]20 D) for lactosucrose was determined as

W. Xu · W. Zhang (*) · H. Wu State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China e-mail: [email protected]; [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_7

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Fig. 7.1 Functional derivatives of sucrose and lactose

+59
. Lactosucrose is stable at pH 4.5 and 120
C for 1 h, and its solubility in water is 3670 g/L at 25
C, which is much higher than that of sucrose (2000 g/L at 25
C). Lactosucrose has 30–60% of relative sweetness compared with sucrose; meanwhile, it has a high-quality taste similar to sucrose, which is distinguished remarkably from the other oligosaccharide sweeteners (Fujita et al. 2009). In 2005, lactosucrose was included in the list of foods for specific health uses (FOSHU) in Japan. Since then, lactosucrose has been widely applied as an important ingredient in the preparation of functional foods. The two major lactosucrose manufacturers in Japan are Ensuiko Sugar Refining Co. and Hayashibara Shoji, Inc., with the product name Nyuka-Origo and Newka-Oligo, respectively (Lígia Rodrigues and Torres 2005). It was reported that in Japan, the annual sales volume of lactosucrose is around several kilotons. By 2009, more than 30 types of food products containing lactosucrose have been approved as FOSHU. As a sweetener and prebiotic ingredient, lactosucrose has also been commercially used in diverse food and beverage systems in Japan, including sweets, candies, desserts, yoghurts, coffee, and tea.

7.2 7.2.1

Physiological Benefits of Lactosucrose Improve the Intestinal Microflora

Lactosucrose is a kind of indigestible carbohydrate that is scarcely hydrolyzed by human digestive enzymes in the upper gastrointestinal tract or by acetone powder prepared from rat intestines, but it could be selectively used by intestinal Bifidobacterium (Fuiita et al. 1991). Lots of studies have shown that lactosucrose could promote the proliferation of Bifidobacterium in the intestine and stomach,

7 Enzymatic Production of Lactosucrose by Levansucrase, β-Fructofuranosidase, and. . .

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Fig. 7.2 Physiological benefits of lactosucrose

wherein the growth of Bifidobacterium would further generate more short-chain fatty acids (SCFAs) (Fig. 7.2). The SCFAs are beneficial to host because they could reduce intestinal pH, enhance the mineral bioavailability, and inhibit the proliferation of intestinal pathogenic bacteria like Clostridium perfringens and Bacteroidaceae (Roberfroid et al. 2010). Therefore, the physiological benefits of lactosucrose mainly result from its resistance to digestion and ability to be fermented by intestinal microbiota and impede the formation of toxic products, such as ammonia, phenol, ethyl phenol, indole, and skatol (Terada et al. 1993, 2009). By improving the intestinal microenvironment, lactosucrose was applied as an agent to deal with the Crohn’s disease and treat ulcerative colitis patients (Teramoto et al. 1996). In vivo, Honda et al. reported the protective effect of lactosucrose on intracolonic indomethacin-induced small intestinal ulcers in rats (Honda et al. 1999). An in vitro fermentation study of lactosucrose also showed that lactosucrose could promote the growth of four bacterial strains including Streptococcus salivarius, Lactobacillus casei, Lactobacillus reuteri, and Lactobacillus acidophilus that are recognized as potential prebiotic strains (García-Cayuela et al. 2014).

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Promote the Absorption of Calcium

The production of SCFAs by lactosucrose fermentation could not only lower the intestinal pH value and improve the intestinal microflora but also enhance calcium solubility and absorption from the intestine. Fujita et al. found that feeding lactosucrose and calcium mixture to weanling rats could strikingly increase the ability of the femur and tibial cancellous cartilage to withstand breakage (Fujita et al. 1999). Kishino et al. employed 45Ca as a tracer for calcium during the transport and deposition into the stomach and found that dietary supplementation with lactosucrose might effectively increase the absorption of calcium in the intestine for the growing rats (Kishino et al. 2006). An in vivo study with young women showed a long-term intake of lactosucrose would enhance the intestinal absorption of minerals such as calcium, magnesium, and phosphorus and reduce bone resorption in their body (Teramoto et al. 2006). Meanwhile, the administration study of repeated lactosucrose supplement in a 1- and 2-week way also led to an enhancement of the intestinal calcium absorption for the healthy men (Fujita 2006). The promotion effect of lactosucrose on calcium absorption was not only limited to mammals but also in teleosts. Kihara et al. found out that the dietary supplementation of lactosucrose would affect the calcium metabolism of red sea bream Pagrus major and result in a greater increase of calcium content in their scales.

7.2.3

Inhibit Fat Accumulation and Obesity

Lactosucrose shows great potential in inhibiting body fat accumulation and preventing obesity. Han et al. reported that the level of 2-monoacylglycerol and plasma triacylglycerol was inhibited by rat brush border membrane vesicles in the female mice when supplemented with lactosucrose (Han et al. 1999). Additionally, suppression of fat accumulation in female mice also resulted in a decrease of parametrical adipose tissue weight, even fed with a high-fat diet (Okuda and Han 2001). Moreover, lactosucrose was speculated to interact with triglyceride, which might inhibit intestinal lipid absorption and then reduce adipose tissue accumulation (Mizote et al. 2009).

7.2.4

Regulate the Immune Response

As mentioned above, lactosucrose could improve the intestinal microflora, and the modification of intestinal microenvironment would in turn increase the level of immunoglobulin A (IgA) in the gut and regulate the mucosal immune response (Hino et al. 2007). The formation of antigen-specific immunoglobulin E (IgE) was also found to be suppressed after lactosucrose intake by intraperitoneal

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immunization with ovalbumin/alum in a mice model, suggesting a potential of lactosucrose against IgE-induced allergic disease (Taniguchi et al. 2007). Besides, Schule et al. reported that lactosucrose could be exploited as a significant excipient for spray-dried powders in the stabilization of IgG (Schüle et al. 2008).

7.2.5

Other Physiological Effects

Oku et al. found the abdominal symptom of lactose intolerance was relieved after lactosucrose intake, suggesting lactosucrose could efficiently prevent the lactose intolerance. Lactosucrose also has an excellent water-holding capacity, which could increase bowel peristalsis and improve fecal formation as well as output (Jie et al. 2000). Moreover, lactosucrose possesses a higher laxative threshold, suggesting a great advantage over other lactose-based prebiotics because diarrhea is frequently described as a side effect of prebiotic intake.

7.3

Production of Lactosucrose

Structurally, lactosucrose (O-β-D-galactopyranosyl-(1, 4)-O-α-D-glucopyranosyl(1, 2)-β-D-fructofuranoside) is a synthesized trisaccharide composing one molar D-galactose, one molar D-glucose, and one molar D-fructose (Fig. 7.3), indicating it could be alternatively produced by transferring the D-galactosyl to sucrose or D-fructosyl to lactose. Microbial levansucrase (EC 2.4.1.10), β-fructofuranosidase (EC EC 3.2.1.26), and β-galactosidase (EC 3.2.1.23) are the most three intensively studied enzymes for the biological production of lactosucrose. Although these enzymes could produce lactosucrose with the same substrates of sucrose and lactose, the conversion ratio and maximum production varied significantly. Moreover, the yield of lactosucrose was not expectedly obtained due to the occurrence of lactosucrose hydrolysis, suggesting a great demand for the improvement of synthetic OH

OH

OH O

O OH

O

HO OH

OH OH

O

O OH OH

Fig. 7.3 Chemical structure of lactosucrose

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process by finding more novel and promising enzymes with enhanced activity or minimizing the product degradation.

7.3.1

Production of Lactosucrose by Levansucrase in Its Cell-Free Form

Levansucrase (sucrose:2, 6-β-D-fructan 6-β-D-fructosyltransferase) belongs to the glycoside hydrolase family 68 (GH 68) and responses for the metabolism of sucrose for microorganism in vivo (Nagarajan and Borchert 1991). In vitro, when different fructosyl acceptors are provided, levansucrase could catalyze different reactions including hydrolysis (only water), transfructosylation (water and another fructosyl acceptor), and polymerization (water and only sucrose) (Ni et al. 2018). Herein, levansucrase plays a significant role in the production of different transfructosylated products, FOS, and β-(2, 6) levan. Specifically, microbial levansucrase could produce lactosucrose as its main transfructosyl product from sucrose and lactosucrose. Table 7.1 gives a summary of the situation about lactosucrose production by different microbial levansucrases. So far, more than ten kinds of levansucrases have been reported in the preparation of lactosucrose. Avigad was the first to investigate the production of lactosucrose by the cell-free enzyme from Aerobacter levanicum (Avigad 1957). 50 g/L sucrose and 292 g/L lactose served as initial fructosyl donor and acceptor, respectively. The pH and temperature were set as pH 5.4 and 30
C, and Torulopsis glabrata cells were applied to remove the glucose and fructose from the mixture. The reaction mixture was also incubated with toluene and was slightly agitated. After 9 h, 50 g/L fresh sucrose was fed to the reaction mixture, and 13 h later, ethanol (96% v/v) was added as the precipitation of the enzyme as well as lactose. Finally, in addition to lactosucrose, a small portion of fructose-containing tri- and tetrasaccharides were also detected in the mixture. However, the yield and conversion ratio of lactosucrose by this enzyme was not reported by that research. Later on, the levansucrase from Bacillus subtilis natto was applied to produce lactosucrose. The enzyme was purified by ammonium sulfate precipitated and analyzed by chromatography. From 85.5 g/L sucrose and 85.5 g/L lactose, the maximum lactosucrose production was obtained as 53 g/L at pH 6.2 and 35
C (Takahama et al. 1991a). However, the immobilization of B. subtilis levansucrase onto molecular sieve (5 Å size), DEAE-Toyopearl 650 M, and hydroxyapatite resulted in a lower production of lactosucrose compared to that of free enzyme. Choi et al. employed the levansucrase from Paenibacillus polymyxa to produce lactosucrose. Two different catalysts including the cell-free enzyme and whole cells were employed and compared. As a result, 140 g/L lactosucrose was obtained from 225 g/L sucrose and 225 g/L lactose in 40 min, and this was lower than that of the whole cell (170 g/L; see below). Heterologous expression has been a powerful and efficient method to obtain more target enzyme and facilitate a further application. Seibel et al. expressed the

6.0

6.5

6.0

6.5

4.0

6.0

7.0

6.0

B. subtilis

B. methylotrophicus

B. goodwinii

L. mesenteroides

P. aurantiaca

P. polymyxa

Z. mobilis

Z. mobilis (coupled with glucose oxidase)

30

23

40

45

50

35

180

180

225

510

270

180

180

205

NRb

37

85.5

35

Sucrose concentration (g/L) 100

180

180

225

360

270

180

180

410

85.5

Lactose concentration (g/L) 292

NR Not reported The lactosucrose conversion is calculated of the total sugar concentration

6.2

B. natto

b

a

pH 5.4

Enzyme source A. levanicum

Temperature (
C) 30

Table 7.1 Production of lactosucrose by levansucrase in its cell-free form

100 224

27b 41b

66 (sucrose) 28 (lactose) 66 (sucrose) 33 (lactose)

NR

156

103

140

285

143

36b

NR

131

53

Maximum lactosucrose production (g/L) NRa

77 (sucrose)

54 (sucrose) 42 (lactose)

Conversion of lactosucrose (%) NRa

34.7

25.8

210

142.5

224

50

7.15

21.9

26.5

Productivity (g/(Lh)) NRa

Reference Avigad (1957) Takahama et al. (1991b) Seibel et al. (2006) Wu et al. (2015) Xu et al. (2018) Li et al. (2015) Han et al. (2007) Choi et al. (2004) Han et al. (2009)

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levansucrase from Bacillus subtilis NCIMB 11871 in Escherichia coli and found the recombinant enzyme had a relative broad substrate spectrum toward D-monosaccharides, L-monosaccharides, and disaccharides during transfructosylation. Under this circumstance, the recombinant enzyme from B. subtilis was applied to produce lactosucrose. From 250 g/L sucrose and 410 g/L lactose, the enzyme exhibited a lactosucrose-producing productivity of 21.9 g/(Lh), and the final conversion rate between lactosucrose and sucrose was obtained as 34% (Seibel Jr et al. 2006). In the case of recombinant levansucrase from Zymomonas mobilis, the lactosucrose conversion efficiency was acquired as 28.5% from 180 g/L sucrose and 180 g/L lactose. Moreover, when glucose oxidase was combined with the Z. mobilis levansucrase to favor the reduction of glucose in the reaction mixture, the lactosucrose conversion efficiency was increased to 43.2% (Han et al. 2009). Different concentrations of sucrose and lactose would largely affect the production of lactosucrose. Wu et al. investigated the effect of sucrose and lactosucrose concentration as well as the ratio of sucrose to lactose on the production of lactosucrose. They employed the recombinant levansucrase from Bacillus methylotrophicus and found a high substrate concentration favored the generation of lactosucrose. Under the optimized conditions pH 6.5 and temperature at 37
C, 143 g/L lactosucrose was obtained from 400 g/L total sugars (sucrose and lactose) after 20 h. Additionally, the 1:1 ratio of sucrose to lactose was validated the best for the lactosucrose production as a compared result of 1:2 or 2:1 (Wu et al. 2015). The conclusion was also drawn by Xu et al. when the recombinant levansucrase from Brenneria goodwinii (Xu et al. 2018) was applied to produce lactosucrose from sucrose and lactose. The condition was optimized as pH at 6.0, temperature at 35
C, and 5 U mL1. Within a 1:1 ratio of sucrose to lactose, the maximal lactosucrose production was acquired as 100 g/L from 180 g/L sucrose and 180 g/L lactose after 6 h, which was higher than that of the other different ratios. In the case of L. mesenteroides levansucrase, Li et al. found that when the concentrations of sucrose and lactose were set as 270 g/L and 270 g/L, and the ratio was 1:1, the highest lactosucrose production was obtained. Notably, the highest production of lactosucrose was observed only in 1 h by the recombinant levansucrase from L. mesenteroides, suggesting an extremely high productivity of lactosucrose (224 g/(Lh)). In contrast to the other levansucrase, the production and productivity was much higher since most of the purified or crude levansucrases produced lactosucrose with a yield less than 100 g/L. In another abstract report, Han et al. obtained the highest lactosucrose production of 285 g/L at pH 4.5 and temperature at 45
C in 2 h of reaction, but they used a very high substrate concentration of 360 g/L lactose and 540 g/L sucrose. The enzyme was from Pseudomonas aurantiaca, but a detailed information with respect to the enzyme was not provided (Han et al. 2007). Even though levansucrase has not been commercially applied in the production of lactosucrose, it is still a promising catalyst compared to β-fructofuranosidase and β-galactosidase not only because of its more microbial resources but also because of its higher production.

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Production of Lactosucrose by Microorganisms Harboring the Levansucrase Activity

Compared to the cell-free form levansucrase, the whole cell catalyst does not need isolation and the purification of levansucrase but allows a continuous production for both enzyme and target product. Also, cells can be easily harvested from the fermentation culture by centrifugation and are directly added to the reaction mixture. What’s more, enzymes in cells tend to be integrated and more stable than the cellfree form. Notably, when using whole cells to produce lactosucrose industrially, many other enzymes, in addition to the desired enzyme, are still in the reaction media, and this would result in some additional and unwanted by-products (Ishige et al. 2005). The production of lactosucrose by whole cells harboring the levansucrase was firstly reported in the case of Paenibacillus polymyxa IFO 3020 (Table 7.2). After a comparison of the cell-free enzyme from the same source, Choi et al. found the whole cell had a higher optimal temperature and maximum lactosucrose production when producing lactosucrose. The maximum lactosucrose production of 170 g/L was detected after a 6 h reaction from 225 g/L sucrose and 225 g/L lactose at 55
C (Choi et al. 2004). After that, the lactosucrose-producing ability of other seven levansucraseharboring microorganisms including the B. subtilis KCCM 32835, Bacillus amyloliquefaciens IFO 15535, Geobacillus stearothermophilus ATCC 12980, Pseudomonas syringae IFO 14086, Paenibacillus polymyxa KCCM 35411, Rahnella aquatilis KTC 2858, and Sterigmatomyces elviae ATCC 18894 were selected and compared by Park et al. (2005). As a result, the B. subtilis cell showed the highest lactosucrose production in a 1-h test and produced 183 g/L lactosucrose from 225 g/ L sucrose and 225 g/L lactose after a 10 h reaction, which was slightly higher than that of P. polymyxa IFO 3020. Among these microorganisms, the S. elviae had a significant big cell size compared to the other bacteria, suggesting an advantage in the continuous production of lactosucrose. Later on, Lee et al. reported a S. elviae ATCC 18894 mutant via the N-methyl-N0 -nitro-N-nitrosoguanidine mutagenesis, and the mutant cell showed a higher lactosucrose production than the wild type. Meanwhile, the cell mass, lactosucrose production, and final lactosucrose yield of S. elviae mutant were increased to 23.3%, 30.42%, and 183.78 g/L, respectively, after optimization of culture medium by introducing analysis of variance (ANOVA) and response surface methodology (RSM) (Lee et al. 2007b). Moreover, the same group reported a continuous lactosucrose employing the immobilized S. elviae mutant cell on calcium alginate beads, and 180 g/L lactosucrose was continuously produced from 250 g/L lactose and 250 g/L sucrose in 48 days (Lee et al. 2007a).

a

NR Not reported

S. elviae ATCC 18894 (immobilized cell on calcium alginate beads)

S. elviae ATCC 18894 (mutant)

Strains B. amyloliquefaciens IFO 15535 B. subtilis KCCM 32835 G. stearothermophilus ATCC 12980 K. pneumoniae ATCC 25306 P. polymyxa P. syringae IFO 14086 R. aquatilis KTC 2858 R. aquatilis ATCC 55046 A. mysorens ATCC 33408 S. elviae ATCC 18894 55 40 40 50 50 50 50 50

6.0 6.0 6.0 6.0 6.0 6.0 6.0 6.0

50

55 50

6.0 6.0

6.0

Temperature (
C) 55 55

pH 6.0 6.0

250/250

34/24 300/300 250/250

300/300

225/225 34/24 34/24 300/300

34/24 300/300

Sucrose/Lactose concentration (g/L) 34/24 225/225

Table 7.2 Production of lactosucrose by microorganisms harboring levansucrase activity

192 (batch); 180 (continuous for 48 days)

7.5 91 141; 184 (after medium optimization)

15

170 3.9 5.6 79

10.4 16

Lactosucrose production (g/L) 4.8 183

7.5 6.1 9.4; 12.3 (after medium optimization) 9.6 (batch); NRa

1.0

28 3.9 5.6 5.3

10.4 1.1

Productivity (g/(Lh)) 4.8 18.3

Lee et al. (2007a)

Reference Park et al. (2005)

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Production of Lactosucrose by β-Fructofuranosidase

β-Fructofuranosidase (β-D-fructofuranoside fructohydrolase) exists widely in plants, fungi, and bacteria. CAZy database showed levansucrase, inulosucrase, and β-fructofuranosidase all belong to the GH 68. However, some other βfructofuranosidases were also reported as GH 32 such as the enzymes from Thermotoga maritima and Bifidobacterium longum (Alberto et al. 2006; ÁvilaFernández et al. 2016). Anyway, both of GH 68 and GH 32 β-fructofuranosidases belong to the “GH-J” clan and primarily catalyze the hydrolysis of β-Dfructofuranoside residues in the nonreducing terminal like sucrose and other fructose-containing saccharides. Namely, it can be called invertase or saccharase (Kotwal and Shankar 2009). In addition to hydrolysis, β-fructofuranosidase is able to catalyze the transfructosylation when fructosyl acceptors are given, but different microbial source of β-fructofuranosidases and acceptor structures may largely affect the transfer ratio, resulting in a series of different short-chain FOSs such as fructosylxyloside, fructosyl-stevioside, and isomaltosylfructoside (Fujita et al. 2014) (Fig. 7.4). For instance, the dominant reaction of Aspergillus niger β-fructofuranosidase could be changed from sucrose hydrolysis to transfructosylation when the sucrose concentration was increased accordingly. When sucrose was provided as sole substrate, the enzyme selectively transferred the fructosyl residue of one sucrose molecule to the C-1 position of the fructose residue of another sucrose, which generated 1-kestose, nystose, and fructosyl-nystose (Hidaka et al. 1988). Different from A. niger β-fructofuranosidase, the enzyme from Arthrobacter sp. K-1 merely catalyzed the hydrolysis of sucrose even at a high substrate concentration. The fructose-transferring ability of soil-derived Arthrobacter sp. K-1 was firstly found by Fujita et al. They purified and validated the β-fructofuranosidase activity of Arthrobacter sp. K-1. Unexpectedly, the enzyme showed a broad affinity Levansucrase:

Glc

Fru +

H2O

Glc

Fru + Gal

Sucrose

Glc + Fru

hydrolysis

Glucose Fructose

Sucrose Glc

Lactose

Glc + Gal

Glc

Fru

transfructosylation

Glucose Lactosucrose

b - fructofuranosidase : Glc

Fru +

H2O

Glucose Fructose

Sucrose Glc

Fru + Gal

Sucrose Gal

Glc

hydrolysis

Glc + Fru

Glc

Lactose Fru +

Lactosucrose

H2O

Glc + Gal

Glc

Fru

transfrutosylation

Glucose Lactosucrose Fru + Gal Fructose

Glc

hydrolysis

Lactose

Fig. 7.4 Different reaction types of levansucrase and β-fructofuranosidase (from Arthrobacter sp. K-1) when sucrose and lactose are provided in the reaction

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Fig. 7.5 Overall structure of Arthrobacter sp. K-1 β-fructofuranosidase (PDB: 3VSS, gray, cartoon model; residues of the tunnel were depicted in blue, stick model) (a) and the tunnel at the bottom for substrate binding (tunnel residues and fructose were labeled and shown in stick model) (b)

toward the other fructosyl acceptors including lactose, xylose, isomaltose, and stevioside, but not for sucrose. Notably, lactose was found as the most effective acceptor with more than 50% transferring ratio as a compared result of the other fructosyl acceptors (Fujita et al. 1990). The sequence of β-fructofuranosidase from Arthrobacter sp. K-1 has been released in NCBI with the GenBank accession No. BAB72022.1. (So far, the identical information of Arthrobacter sp. K-1 has been updated as Microbacterium saccharophilum.) Later, the recombinant enzyme was obtained by sub-cloning the identical gene in E. coli and overexpressed by IPTG induction. The recombinant enzyme showed the optimal pH and temperature at 6.5 and 55
C, which is much similar to that of the wild type. Moreover, the recombinant enzyme was found stable when incubated at 60
C for 0.5 h, showing approximately 70% of residual activity (Ito et al. 2002). Tonozuka et al. determined the crystal structure of βfructofuranosidase from M. saccharophilum (PDB No. 3VSR, 3VSS) and implied the catalytic cleft as well as the tunnel at the bottom might benefit for the lactosucrose by comparing the structure of Thermotoga maritima and Bifidobacterium longum β-fructofuranosidase (Fig. 7.5) (Tonozuka et al. 2012). Several water molecules were also observed in the vicinity of the tunnel, and these waters were speculated to function as a water drain or reservoir to pass through the tunnel to favor the production of lactosucrose. A large number of studies have been reported for the Arthrobacter sp. K-1 βfructofuranosidase from a Japanese company: Ensuiko Sugar Refining Company. From 137 g/L sucrose and 137 g/L lactose, the maximum lactosucrose production 93 g/L was obtained by the commercial Arthrobacter sp. K-1 β-fructofuranosidase, and the productivity was 63.4 g/(Lh). When the reaction temperature was set at 43
C, the maximum production was increased to 202 g/L compared to that of 50
C, but the productivity showed a bit decrease (60.6 g/(Lh)). In addition to lactosucrose production, the hydrolysis of sucrose and lactosucrose was also observed during the

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reaction, which might be a side effect for the production of lactosucrose. Meanwhile, Pilgrim et al. reported that the hydrolysis of sucrose and lactosucrose could be prohibited by glucose and lactose, respectively (Pilgrim et al. 2001). To overcome the problem, Kawase et al. employed a stimulated moving bed reactor (SMBR) for the production of lactosucrose and kept the same enzyme source. In this SMBR, enzymatic production and separation of lactosucrose happened simultaneously but in different refining and extract ports. Finally, the maximum production of lactosucrose was acquired as 133 g/L from 171 g/L sucrose and 181 g/ L lactose. However, when the sucrose conversion was 70%, the yield of lactosucrose only reached up to 53%, indicating hydrolysis of lactosucrose still remained, impeding a further increase of the lactosucrose yield (Kawase et al. 2001). Pilgrim et al. optimized the condition in the SMBR and increased the maximum production of lactosucrose to 348 g/L from 342 g/L sucrose and 342 g/L lactose. Also, the amount of lactosucrose obtained from refining port was increased from 9 to 40 g/L when they prolonged the refining time from 1320 to 3005 min (Pilgrim et al. 2006). Mikuni et al. immobilized the Arthrobacter sp. K-1 β-fructofuranosidase onto FE4611 resin and obtained a continuous production of 120 g/L lactosucrose from 200 g/L sucrose and 200 g/L lactose at pH 6.5 and 55
C, suggesting immobilization of Arthrobacter sp. K-1 β-fructofuranosidase could be a feasible way in the industrial preparation of lactosucrose (Mikuni et al. 2000). The Bacillus sp. 417 isolated from soil, fruit, and flowers shows a great potential in the production of β-fructofuranosidase. Meanwhile, the lactosucrose-producing ability was validated when sucrose and lactose were given as fructosyl donors and acceptors. At 45
C and pH 5.6, Bacillus sp. 417 could produce 54 g/L lactosucrose from 200 g/L sucrose and 200 g/L lactose after an 8 h of reaction, which was similar to that of B. natto (Ikegaki and Park 1997). An extracellular β-fructofuranosidase from Bacillus sp. V230 was also reported as lactosucrose producer in a Japanese patent (No. 224665/97). The enzyme was purified by ion-exchange and hydrophobic chromatography, and Saccharomyces cerevisiae was coupled to remove the generated monosaccharides like glucose and fructose as well as improve the lactosucrose production. When pH and temperature was controlled to 4.0–5.5 and 30
C, more than 70% of lactosucrose was produced from 200 g/L sucrose and 200 g/L lactose after 24 h (Hiroyuki Okabe 2008). It is well-known the β-fructofuranosidase from Arthrobacter sp. K-1 would be easily deactivated at a high temperature (Tonozuka et al. 2012). Recently, the recombinant β-fructofuranosidase from Arthrobacter sp. 10,138 has been reported as another kind of lactosucrose-producing β-fructofuranosidase. Even though the maximum lactosucrose was obtained at 40
C, the enzyme was highly thermostable in the range of 40–50
C. At 50
C and pH 6.0, the recombinant enzyme (40 μg/mL,) could efficiently produce 109 g/L lactosucrose from 150 g/L sucrose and 150 g/L lactose in 10 min, with a molar conversion ratio of 49.3%, suggesting a great advantage for the production of lactosucrose (Chunmei et al. 2020). Long et al. reported the co-immobilization of Arthrobacter sp. 10,137 βfructofuranosidase and glucose oxidase to improve the production of lactosucrose. Glucose was a by-product during the transfructosylation of lactose and sucrose;

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herein the glucose oxidase was introduced to transform glucose into gluconic acid to favor the formation of lactosucrose. A sol-gel system consisting of aqueous polyvinyl alcohol, distilled water, aqueous sodium fluoride (NaF), n-octyltriethoxysilane (OTES), and tetraethoxysilane (TEOS) was constructed for the co-immobilization of β-fructofuranosidase and glucose oxidase. The coupled bi-enzymes could keep 85.39% of their initial activity during immobilization and show high operational stability (maintaining 78.5% of their initial activity) after 15 consecutive cycles. Meanwhile, the co-immobilized enzyme generated 160.8 g/L lactosucrose from 200 g/L sucrose and 200 g/L lactose, which is significantly higher than that of the immobilized β-fructofuranosidase from Arthrobacter sp. K-1 (Table 7.3) (Long et al. 2019).

7.3.4

Production of Lactosucrose by β-Galactosidase

β-Galactosidase (β-D-galactoside galactohydrolase) is a kind of glycosyl hydrolases belonging to GH 35 family. It can also be called lactase since it efficiently catalyzes the hydrolysis of lactose into galactose and glucose. In addition to hydrolysis, βgalactosidase could also catalyze the transgalactosylation. So far, β-galactosidase has been intensively applied in the milk industry for the hydrolysis of lactose as well as the synthesis of functional lactulose, epilactose, and galactooligosaccharides (GOSs). The production of lactulose and epilactose was finished in the presence of cellobiose 2-epimerase which has been discussed in a previous section. Here, we focused on the production of lactosucrose by Bacillus circulans β-galactosidase, which is known as the only one β-galactosidase with the lactosucrose-producing ability. The β-galactosidase from B. circulans could transfer the galactosyl to the other galactosyl acceptors like sucrose to produce lactosucrose by forming a β-(1, 4) linkage between galactosyl and glucosyl units. In fact, lactose was also found as a galactosyl acceptor during the transgalactosylation by B. circulans β-galactosidase and formed β-(1, 3) linkage between galactosyl and galactosyl units (Fig. 7.6). Li et al. optimized the condition of substrate concentration, enzyme amount, and temperature and obtained 56 g/L lactosucrose from 300 g/L sucrose and 300 g/L lactose at pH 6.0 and 40
C (Li et al. 2009). Recently, Duarte et al. reported the immobilization of B. circulans β-galactosidase on chitosan marcospheres and increased the enzyme thermostability by 260-fold during lactosucrose production. With the immobilized enzyme, 79 g/L of lactosucrose and 37 g/L of GOSs were produced at 30
C, and the concentration could be 40 and 62 g/L when the temperature increased to 64
C. Compared to the results obtained from free enzyme, immobilized β-galactosidase displayed a potential in the industrial preparation of lactosucrose and functional GOSs (Duarte et al. 2017). As abovementioned, the production of lactosucrose could be finished by levansucrase and β-fructofuranosidase by hydrolyzing sucrose into glucose and fructose and transferring the fructose into lactose, or by β-galactosidase by

5.6

4.0–5.5

Bacillus sp. 417

Bacillus sp. V230

30

45 200/200

200/200

200/200

150/150

200/200

342/342 342/342

171/181

222/137

Sucrose/ Lactose (g/L) 137/137

NR Not reported The lactosucrose conversion is calculated of the total sugar concentration

b

a

6.5

Arthrobacter sp. 10,137

35

50

50 50

NRa NRa

6.0

50

NRa

55

43

NRa

6.5

Temperature (
C) 50

pH NRa

Arthrobacter sp. K-1 (immobilized on FE4611 resin) Arthrobacter sp. 10,138

Enzyme source Arthrobacter sp. K-1

Table 7.3 Production of lactosucrose by β-fructofuranosidase

160.8

40.2%b

NR

NR

NR

54

109

49.3%b

NRa

348 40 (products the refined) using a SMB reactor 120

133

202

Lactosucrose production 93

NRa NRa

Conversion (%) 56 (sucrose) 47 (lactose) 25 (sucrose) 33 (lactose) 70 (sucrose)

NR

6.75

NRa

654

NRa

NRa NRa

NRa

60.6

Productivity (g/(Lh)) 63.4

Mikuni et al. (2000) Chunmei et al. (2020) Long et al. (2019) Ikegaki and Park (1997) Maniatakos (2016)

Kawase et al. (2001) Pilgrim et al. (2006)

Reference Pilgrim et al. (2001)

7 Enzymatic Production of Lactosucrose by Levansucrase, β-Fructofuranosidase, and. . . 139

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Fig. 7.6 Production of lactosucrose and its analogues by B. circulans β-galactosidase from sucrose and lactose

hydrolyzing lactose into glucose and galactose and transferring the galactose into sucrose. Both of the two processes involved a ratio of transglycosylation to hydrolysis, and the ratio would greatly impact the production of lactosucrose. Several factors including the enzyme sources, ratio of donors to acceptors, and intrinsic enzyme properties can affect the ratio. The enzyme from different microbial resources can have different substrate-binding abilities (exclude water) and lactosucrose-producing abilities. For instance, the Arthrobacter sp. K-1 βfructofuranosidase performed highly in the production of lactosucrose as a compared result of other β-fructofuranosidases, and the B. circulans β-galactosidase was the only one β-galactosidase member that can produce lactosucrose. Herein, how to design those unsatisfied lactosucrose-producing enzymes to improve their ability was a big concern. So far, many attempts have been made to improve the portion of transglycosylation to hydrolysis based on the protein engineering. A drastic shift from water to sucrose as the preferred acceptor was observed in the mutant of recombinant invertase from Triticum aestivum, suggesting wheat vacuolar invertase was engineered into a high-affinity sucrose:sucrose 1-fructosyltransferase (Schroeven et al. 2008). A mutagenic study was also conducted in Geobacillus stearothermophilus β-galactosidase, and a simultaneous improvement of the transglycosylation activity and reduction of hydrolysis was obtained, and this also increased the production of transglycosylation products (Placier et al. 2009a).

7 Enzymatic Production of Lactosucrose by Levansucrase, β-Fructofuranosidase, and. . .

7.4

141

Application of Lactosucrose

With numerous physiological benefits, lactosucrose has been applied in the food, pharmaceutical, and cosmetic industries. Also, some patents reported the application of lactosucrose in the pet food and fish feed. Table 7.4 gives a summary of the recent application about lactosucrose. It should be noted that the practical use of lactosucrose has definitely not limited the lists below.

7.5

Conclusion and Future Perspectives

As a significant indigestible trisaccharide, lactosucrose can be selectively used by Bifidobacterium and regulate the intestinal microflora. Due to its numerous physiological functions such as promoting the absorption of calcium, inhibiting fat accumulation and obesity, and regulating immune response, the demand of lactosucrose was intensively increased in the past few years. Biological production of lactosucrose from sucrose and lactose is environmentally friendly, economical, and promising. Three representative lactosucrose-producing enzymes, including levansucrase, β-fructofuranosidase, and β-galactosidase, were discussed and compared in detail with respect to their microbial resources, lactosucrose production, and industrial application. So far, only the β-fructofuranosidase from Arthrobacter sp. K-1 was commercially applied in the industrial preparation in a stimulated Table 7.4 Application of lactosucrose (Partially) Description Low-calorie, low-digestive

Regulates intestinal microflora, reduce unpleasant odor of feces and urine Improves nutrient absorption and decreases self-contamination Nutritional support and prevents skin disease Model as a transglycosylation product

Application In bakery products Ingredient in yogurts Added in ice cream, snacks, cookies, desserts, candies, infant formula Added into the pet food Feed for fish Excipient the pharmaceutical and cosmetic products Enzymatic tests for hydrolysis by βgalactosidase Induces the production of αgalactosidase Stabilizes protein Stabilizes polyplexes

Reference Takumi et al. (2001) Mu et al. (2013) Maniatakos (2016) Bunch (1997) Stefan Bassarab et al. (2005) Placier et al. (2009b) Quang et al. (2003) Schüle et al. (2008) Kasper et al. (2011)

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moving bed reactor, and the highest production of lactosucrose after refining was obtained as 40 g/L from 342 g/L sucrose and 342 g/L lactose. Even though levansucrase has not been industrially applied for lactosucrose production, the enzymes from B. subtilis, P. aurantiaca, and L. mesenteroides exhibited a much higher lactosucrose-producing ability. Herein, finding out more industrial-valuable catalyst for lactosucrose production is still a big issue. Meanwhile, protein engineering plays a significant role in the rational design of enzyme and in improving the catalytic efficiency as well as the yield of lactosucrose. Last but not least, how to minimize the side effect of by-product such as glucose on the lactosucrose production is also important for the downstream process.

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Wittrant Y, Delzenne NM, Cani PD, Neyrinck AM, Meheust A (2010) Prebiotic effects: metabolic and health benefits. Br J Nutr 104(Suppl 2):S1–S63. https://doi.org/10.1017/ s0007114510003363 Schroeven L, Lammens W, Van Laere A, Van den Ende W (2008) Transforming wheat vacuolar invertase into a high affinity sucrose:sucrose 1-fructosyltransferase. New Phytol 180 (4):822–831. https://doi.org/10.1111/j.1469-8137.2008.02603.x Schüle S, Schulz-Fademrecht T, Garidel P, Bechtold-Peters K, Frieb W (2008) Stabilization of IgG1 in spray-dried powders for inhalation. Eur J Pharm Biopharm 69(3):793–807. https://doi. org/10.1016/j.ejpb.2008.02.010 Seibel J Jr, Moraru R, Götze S, Buchholz K, Na’amnieh S, Pawlowski A, Hecht H Jr (2006) Synthesis of sucrose analogues and the mechanism of action of Bacillus subtilis fructosyltransferase (levansucrase). Carbohydr Res 341(14):2335–2349. https://doi.org/10. 1016/j.carres.2006.07.001 Stefan Bassarab KBP, Fuhrherr R, Friess W, Gariderl P, Schutlz-Fademrecht T (2005) Powder comprising new compositions of oligosaccharides and methods for their preparation. US patent Takahama JK, Okano S, Kyoko A, Toshie N, Hiroshi T, Kobayashi T (1991a) Production of Lactosucrose by bacillus natto Levansucrase and some properties of the enzyme. Nippon Shokuhin Kogyo GakkaiShi 38:789–796 Takahama A, Kuze J, Okano S, Akiyama K, Nakane T, Takahashi H, Kobayashi T (1991b) Production of Lactosucrose by bacillus natto Levansucrase and some properties of the enzyme. Nippon Shokuhin Kogyo Gakkaishi 38(9):789–796. https://doi.org/10.3136/nskkk1962.38.789 Takumi H, Ochi H, Okada S, Li S-T, Terada A, Mitsuoka T (2001) Effect of ingesting frozen yoghurt in combination with Lactosucrose consumption on the Fecal microbiota and Fecal Metabolitic activity in healthy adults. Jpn J Food Microbiol 18(2):49–56. https://doi.org/10. 5803/jsfm.18.49 Taniguchi Y, Mizote A, Kohno K, Iwaki K, Oku K, Chaen H, Fukuda S (2007) Effects of dietary lactosucrose (4G-beta-D-galactosylsucrose) on the IgE response in mice. Biosci Biotechnol Biochem 71(11):2766–2773. https://doi.org/10.1271/bbb.70364 Terada A, Hara H, Kato S, Kimura T, Fujimori I, Hara K, Maruyama T, Mitsuoka T (1993) Effect of lactosucrose (4G-beta-D-galactosylsucrose) on fecal flora and fecal putrefactive products of cats. J Vet Med Sci 55(2):291–295. https://doi.org/10.1292/jvms.55.291 Terada A, Hara H, Oishi T, Matsui S, Mitsuoka T, Nakajyo S, Fujimori I, Hara K (2009) Effect of dietary Lactosucrose on Faecal Flora and Faecal metabolites of dogs. Microb Ecol Health Dis 5 (2):87–92. https://doi.org/10.3109/08910609209141294 Teramoto F, Rokutan K, Kawakami Y, Fujimura Y, Uchida J, Oku K, Oka M, Yoneyama M (1996) Effect of 4G-beta-D-galactosylsucrose (lactosucrose) on fecal microflora in patients with chronic inflammatory bowel disease. J Gastroenterol 31(1):33–39. https://doi.org/10.1007/ bf01211184 Teramoto F, Rokutan K, Sugano Y, Oku K, Kishino E, Fujita K, Hara K, Kishi K, Fukunaga M, Morita T (2006) Long-term administration of 4G-beta-D-galactosylsucrose (lactosucrose) enhances intestinal calcium absorption in young women: a randomized, placebo-controlled 96-wk study. J Nutr Sci Vitaminol 52(5):337–346. https://doi.org/10.3177/jnsv.52.337 Tian Y, Deng Y, Zhang W, Mu W (2019) Sucrose isomers as alternative sweeteners: properties, production, and applications. Appl Microbiol Biotechnol 103(21–22):8677–8687. https://doi. org/10.1007/s00253-019-10132-6 Tonozuka T, Tamaki A, Yokoi G, Miyazaki T, Ichikawa M, Nishikawa A, Ohta Y, Hidaka Y, Katayama K, Hatada Y, Ito T, Fujita K (2012) Crystal structure of a lactosucrose-producing enzyme, Arthrobacter sp. K-1 β-fructofuranosidase. Enzym Microb Technol 51(6):359–365. https://doi.org/10.1016/j.enzmictec.2012.08.004 Wu C, Zhang T, Mu W, Miao M, Jiang B (2015) Biosynthesis of lactosylfructoside by an intracellular levansucrase from bacillus methylotrophicus SK 21.002. Carbohydr Res 401:122–126. https://doi.org/10.1016/j.carres.2014.11.001

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Xiao Y, Chen Q, Guang C, Zhang W, Mu W (2019) An overview on biological production of functional lactose derivatives. Appl Microbiol Biotechnol 103(9):3683–3691. https://doi.org/ 10.1007/s00253-019-09755-6 Xu W, Liu Q, Yu S, Zhang T, Mu W (2018) Synthesis of Lactosucrose using a recombinant Levansucrase from Brenneria goodwinii. Appl Biochem Biotechnol 186(2):292–305. https:// doi.org/10.1007/s12010-018-2743-1

Chapter 8

Difructose Anhydrides-Producing Fructotransferase: Characteristics, Catalytic Mechanism, and Applications Mei Cheng, Yingying Zhu, and Wanmeng Mu

8.1

Introduction

Difructose anhydrides (DFAs) are formed by two D-fructose molecules, with the loss of two water molecules. The typical structure of DFAs is cyclic fructodisaccharides with two reciprocal glycosidic linkages. DFAs are commonly formed upon thermal or protonic activation of D-fructose, sucrose, and two basic fructans (inulin and levan) which consisted of fructose residues by fructosyl linkages (Garcíamoreno et al. 2008). In the late 1930s, several DFAs have been isolated from higher plants, including DFA I (α-D-fructofuranose-β-D-fructofuranose 1,20 :2,10 -dianhydride) from Jerusalem artichoke, DFA IV (β-Dfructofuranose-β-D-fructofuranose 20 ,6:2,60 -dianhydride) from Allium sewertzowi, and DFA III (α-D-fructofuranose-β-Dfructofuranose 1,20 :2,30 -dianhydride) from Lycoris radiata (Uchiyama 1983; Garcíamoreno et al. 2008). However, it remains to be thrown light on the true generator of the isolated DFAs in higher plants. To date, 13 DFA isomers with 5 different tricyclic cores have been characterized from the fructose caramel (Arribas et al. 2010). Additionally, four DFA isomers have been synthesized by enzymatic strategy from fructans: DFA I, DFA III, DFA IV, and DFA V (α-D-fructofuranose-β-D-fructofuranose 20 ,6:2,10 dianhydride) (Wang et al. 2015a). DFAs can be formed during the acid-hydrolyzing inulin (McDonald 1946; Wang et al. 2015a). Subsequently, researchers found that DFAs were synthesized by fructose and fructose-containing substrates, under high temperature condition and acidic treatment (Christian et al. 2000; Suárez-Pereira et al. M. Cheng · Y. Zhu State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China W. Mu (*) State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_8

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2010; Trabs et al. 2011). Naturally, DFAs occurred in caramelization of D-fructose, glucose or glucose syrups during food process (Golon and Kuhnert 2013). Meanwhile, DFAs have been treated as chemical markers for caramelized foods. DFAs can be naturally formed from fructans (inulin and levan) by fructotransferases. The investigations on further metabolism of DFAs are mainly focused on DFA III. DFAs are proposed to be decomposed into inulobiose. In the 1980s, DFA I can be partial hydrolysis into inulobiose, and be formed from inulobiose by this reverse reaction (Matsuyama et al. 1982; Matsuyama and Tanaka 1989). Compared with other DFAs, enzymatic hydrolysis of DFA III has been deeply investigated. DFA III can be depolymerized into inulobiose by difructose anhydrate hydrolase (DFAase) (EC 3.2.1.-) which has been classified to glycoside hydrolase family 91 (GH91) (Saito et al. 2003b; Yu et al. 2018). In the metabolic pathway of microorganisms, inulobiose can be further degraded into two fructose molecules by β-fructofuranosidase (Sakurai et al. 1997b). Nowadays, investigations on DFAs have attracted increasing attention, as their beneficial bio-functions and promising properties as functional foods. At the present chapter, the advances in the physiological functions, enzymatic synthesis, biological productions, and applications of DFAs, at the same time, the structures and catalysis mechanisms of related enzymes, are discussed.

8.2

Physiological Functions of DFAs

DFAs are a family of cyclic disaccharide, with two different reciprocal glycosidic linkages (Garcíamoreno et al. 2008). DFAs are white crystals and soluble in water and remain stable at acid and heat treatments. As non-reducing sugars, DFAs are chemically stable. Thus far, nutritional functions of DFAs are mainly investigated by Japanese researchers, and the related reports were primarily published in national Japanese journals. Physiological functions of DFA III, DFA I, and DFA IV have been extensively investigated, which benefits from their efficient production.

8.2.1

Low-Calorie Sweeteners

Commonly, DFAs have been considered as a promising low-calorie sweetener. DFA III, a typical case, is an ideal table sugar substitute, which has half sweetness but 1/15 calorie of sucrose (Kikuchi et al. 2004; Haraguchi 2011). DFA III and DFA IV are indigestible and unabsorbable sugars, in vitro (Saito and Tomita 2000). Additionally, more researches in humans have been implemented. DFA III is indigestible and low fermentable during the early stages after ingestion in humans (Tamura et al. 2003b). Ingestion of DFA III does not change the serum levels of glucose, fructose, and insulin. High-dose repeated oral ingestion of DFA III in humans, 9 g daily and

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divided for 3 times, can lead to transitory diarrhea-related symptoms, but no serious adverse effects (Tamura et al. 2003a). Thus, DFA III is suggested to have a laxative effect, which is similar with the other oligosaccharides. Rats fed with DFA III exhibit decreased body energy accumulation and fat tissue weight (Fujitani et al. 2017).

8.2.2

Prebiotic Function

DFA III and DFA IV can be transformed into short-chain fatty acids by intestinal microbes, which can change the microenvironment of intestinal and improve the population and properties of gut microorganism (Saito and Tomita 2000; Minamida et al. 2006b; Saito et al. 1999). Moreover, DFA III administration facilitates the growth of Ruminococcus sp. M-1 and Ruminococcus productus, the acetateproducing bacteria, which decrease pH of intestinal tract (Minamida et al. 2005a, b, 2006a). Acidification of intestine inhibits the formation of secondary bile acid which is associated with occurrence of colon cancer (Minamida et al. 2005a, 2006a). Therefore, DFA III is recommended as a promising prebiotic with the ability of preventing colorectal cancer.

8.2.3

Mineral Absorption

Both DFA III and DFA IV have been reported to raise the absorption of several essential minerals. After fermented by intestinal microorganism, DFA III can stimulate calcium absorption in rats, and the positive effect is efficient than the calcium utility increased by fructo-oligosaccharides or raffinose (Tomita et al. 1999; Suzuki et al. 1998; Shiga et al. 2003). Supplementation of DFA IV in broilers can improve the absorption of calcium and iron concentrations (Lee and Kim 2018). Feeding DFA III enhances calcium absorption in the duodenum of cows via a paracellular pathway (Teramura et al. 2015a, b). Low dosage of DFA III has a positive effect on calcium absorption in humans (Shigematsu et al. 2004). The mechanism of increase in calcium absorption has been researched both in vitro and in vivo experiment. DFA III, DFA IV, and melibiose promote calcium absorption via affecting the epithelial tissue and activating the passage of tight junctions, in vitro (Mineo et al. 2002; Suzuki and Hara 2006). To increase calcium absorption in vivo, DFA III can directly affect the epithelial tissue in the lumen and indirectly improve the short-chain fatty acids produced by microbial fermentation (Mineo et al. 2003, 2006). DFA III intensify iron absorption through increasing iron transporter in the whole cecum with the mucosal expansion (Hara et al. 2010). Ingestion of DFA III enhances bioavailability of water-insoluble iron and hemoglobin concentration in anemic Vietnamese women, which suggests DFA III can prevent anemia or iron deficiency (Nakamori et al. 2010). 15 g/kg DFA III stimulus intestinal absorption of both calcium and magnesium (Mitamura and Hara 2005). DFA III accompanied with

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zinc gluconate improves zinc absorption in rats (Hachiya et al. 2006). Accordingly, DFA III and DFA IV are recommended to prevent osteoporosis, with the improvements of mineral absorption.

8.2.4

Immune System

Additionally, the other nutritional properties of DFA III have attracted attention. DFA III can increase peak serum immunoglobulin G (IgG) concentration of newborn calves but has an unclear effect on incidence of diarrhea and respiratory disease (Htun et al. 2016, 2018). Rat experiment indicated that DFAIII stimulates absorption of α G-rutin, a water-soluble antioxidant, in the small intestine (Matsumoto et al. 2007). DFA III has a prebiotic effect on cholesterol metabolism via increasing the plasma equol concentration (Tamura et al. 2006).

8.2.5

Adverse Effect

Reports on the adverse effects of DFAs are relatively scarce. Consumption with DFA III in rats shows no adverse effects on food intake, body weight gain, nor the weight of kidneys and liver (Suzuki et al. 1998). Tamura et al. revealed that DFA III has no adverse effects on body weight gain, but lower the food intake (Tamura et al. 2006). DFA III is as safe as palatinose through repeated ingestion in humans, with no differences in blood parameters among the ingestion periods, and its high resistance to enterobacterial fermentation, but occurrence of suffering diarrhea and transitory abdominal symptoms, is still not clear (Tamura et al. 2003a, 2004). In a study of 4-week single and repeated-dose toxicities of DFA IV, a high dosage (2000 mg kg1) had no harmful effect on rats (Lee et al. 2004). In summary, based on the outstanding properties of DFAs, they are expected to be nutritional additive in functional food.

8.3

Enzymatic Synthesis of DFAs

The synthesis of DFAs via biotechnological strategies has attracted growing attention. So far, four DFA isomers have been obtained from fructans (inulin and levan) and fructans-containing raw materials through enzymatic strategy, as shown in Fig. 8.1. DFA I and DFA III can be transferred from inulin, a typical fructan connected by β-(2, 1) glycosidic bonds, by inulin fructotransferase (IFTase) (DFA I-forming) (EC: 4.2.2.17) and (DFA Ш-forming) (EC: 4.2.2.18), respectively (Haraguchi 2011). Additionally, DFA V can be formed by Aspergillus fumigatus in the culture medium which contains inulin as a sole carbon source (Matsuyama

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Fig. 8.1 Enzymatic production of DFAs

et al. 1991). However, enzymes for producing DFA V have not been identified. IFTases were categorized as lyases rather than transferases until the year 2004. The previous enzyme commission (EC) numbers of DFA III-forming IFTase and DFA I-forming IFTase were EC 2.4.1.93 and EC 2.4.1.200, respectively (Wang et al. 2015a). Levan, connected by β-(2, 6) glycosidic bonds, can be catalyzed into DFA IV by levan fructotransferase (LFTase) (DFA IV-forming) (EC 4.2.2.16) (Tanaka et al. 1981; Saito et al. 1997).

8.3.1

IFTase for DFA III-Forming

Among DFAs-forming enzymes, IFTase of DFA III-forming is the earliest and mostly investigated ones. In 1972, a DFA III-forming microorganism has been isolated from the soil and has been characterized as Arthrobacter ureafaciens (Tanaka et al. 1972). The strain can secrete an extracellular enzyme which transforms inulin into DFA III. Thereafter, a IFTase (DFA III-forming) was identified from A. ureafaciens, previously named as inulase II, and the enzymatic properties of the purified enzyme was firstly characterized (Uchiyama et al. 1973). This inulase II effectively catalyzes β-(2, 1)-linked inulin, but unacted on β-(2, 6)-connected levan (Uchiyama 1975).

8.3.1.1

IFTase (DFA III-Forming) Production from Microorganisms

To date, 17 IFTase-producing microbial strains have been reported to obtain DFA III (Table 8.1), and the bulk of strains belong to Arthrobacter species, including

Enzymes IFTase (DFA III-forming)

62

43 44

Native

Recombinant

Native

Native

Native

Bacillus sp. snu-7

A. pascens T13-2

Arthrobacter sp. L68-1 Leifsonia sp. T88-4

44

58

44

Native

49

Native

27

Native 49

43

Recombinant

Native

45

Native

Flavobacterium sp. LC-413 Bacillus sp. snu-7

Arthrobacter sp. H65-7 Arthrobacter sp. A-6

A. globiformis C11-1 A. globiformis C11-1 A. ilicis OKU17B

Microorganisms A. ureafaciens

Native or recombinant Native

74

73

79

88

NR

45

145

100

50

NR

50

Molecular weight (kDa) SDSGel PAGE filtration 80 NRb

5.0

5.5–6.0

5.5–6.0

5.0

6.0

6.0

6.0

5.5

5.5

5.0

5.0

Optimum pH 6.0

Table 8.1 Comparison of IFTases and LFTases form various microorganisms

65

55

50

35

40

50

70

60

60

50–60

55

Optimum temperature ( C) 50

644

NR

NR

1276

48.5

5728

1195

NR

853

227

294

Specific activity (U mg1) 162.4

60; 30 min

80; 60 min

75; 20 min

60; 103 min

60; 10 min

70; >300 min 80; 30 min

40; >20 min

35; 120 min

70; 70 min

55; 239 min

70; 15 min

70; 187 min

70; 30 min

80; 240 min

Thermostability ( C; min)a 75; 20 min

Yu et al. (2015) Zhu et al. (2016) Cheng et al. (2017) Cheng et al. (2017) Cheng et al. (2017) Cheng et al. (2020) Tanaka et al. (1983) Saito et al. (1997) Song et al. (2000)

References Haraguchi and Ohtsubo (2006)

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54

Native

NR

NR

46

6.5

7.0

7.0

45

40

40

1432

145.2

23,202

NR

NR

NR

The values exhibited the enzyme incubating at specific temperature and time, with more than half of initial enzymatic activity NR not reported

b

a

54

Recombinant

Microbacterium sp. AL-210 A. oxydans J17-21

45.2

Native

Microbacterium sp. AL-210 Yang et al. (2002) Jang et al. (2003)

Cha et al. (2001)

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A. ureafaciens (Tanaka et al. 1972; Uchiyama et al. 1973), A. globiformis C11-1 (Haraguchi et al. 1988, 2000), A. ilicis OKU17B (Kawamura et al. 1988), Arthrobacter sp. H65-7 (Yokota et al. 1991a; Saito et al. 2003a; Sakurai et al. 1997a), Arthrobacter sp. A-6 (Park and Choi 1996; Kim et al. 2000a, b), Arthrobacter sp. Buo141 (Jahnz et al. 2001), A. pascens T13-2 (Haraguchi et al. 2002), Arthrobacter sp. L68-1 (Haraguchi et al. 2005; Haraguchi 2013), Arthrobacter sp. B30-2 (Haraguchi 2009), A. ureafaciens D13-3 (Haraguchi 2010), A. aurescens SK 8.001 (Zhao et al. 2011a, 2011b; Zhan et al. 2015; Ko et al. 2019a), Arthrobacter sp. 161MFSha2.1 (Wang et al. 2015b), and A. chlorophenolicus A6 (Zhu et al. 2018a). Meanwhile, IFTase-producing strains from non-Arthrobacter species include Pseudomonas fluorescens No. 949 (Kuramoto et al. 1987), Flavobacterium sp. LC-413 (Cho et al. 1997), Bacillus sp. snu-7 (Kim et al. 2007; Kang et al. 1998), Leifsonia sp. T88-4 (Haraguchi et al. 2006), Microbacterium sp. S48-1 (Haraguchi 2015), and Nonomuraea sp. ID06A0189 (Pudjiraharti et al. 2011, 2014). These microorganisms can secrete IFTase (DFA III-forming) under the condition of inulin as the sole carbon source. To improve the activity of DFA III-forming IFTase, the optimized carbon source is identified as inulin for both Arthrobacter sp. H65-7 and A. aurescens SK 8.001 (Yokota et al. 1991b; Zhao et al. 2013). The most investigated strain is A. aurescens SK 8.00, which has been heterologously expressed in various hosts including, Escherichia coli (Zhao et al. 2011a), Pichia pastoris (Zhan et al. 2015), and Saccharomyces cerevisiae (Ko et al. 2019a). Both prokaryotic and eukaryotic hosts have been utilized for gene cloning of IFTase (DFA III-forming). For the prokaryotic expression system, E. coli is a popular choice. To date, the gene of IFTase (DFA III-forming), screened from ten strains and expressed by E. coli, includes A. globiformis C11-1 (Haraguchi et al. 2000), Arthrobacter sp. H65-7 (Sakurai et al. 1997a), Arthrobacter sp. A-6 (Kim et al. 2000a), Arthrobacter sp. Buo141 (Jahnz et al. 2001), A. aurescens SK 8.001 (Zhao et al. 2011a), Arthrobacter sp. L68-1 (Haraguchi 2013), Arthrobacter sp. 161MFSha2.1 (Wang et al. 2015b), A. chlorophenolicus A6 (Zhu et al. 2018a), Bacillus sp. snu-7 (Kim et al. 2007), and Nonomuraea sp. ID06A0189 (Pudjiraharti et al. 2014). DFA III-forming IFTase is an extracellular protein in wild strains, and the recombinant one could be partially secreted extracellular by E. coli (Wang et al. 2015b). The signal peptide sequence of the recombinant IFTase (DFA III-forming) is the same as that of wild enzymes, which suggests E. coli and original strains share the same cleavage sites of signal peptidase of the enzyme. Additionally, the recombinant Bacillus subtilis DB104, containing the gene of IFTase (DFA III-forming) from Arthrobacter sp. A-6, was able to efficiently secrete the extracellular enzyme (Kim et al. 2000b). In the eukaryotic expression system, Pichia pastoris and Saccharomyces cerevisiae have been investigated. Zhan et al. have deeply investigated the expression of DFA III-forming IFTase-encoding gene from A. aurescens SK 8.00 by the eukaryotic expressional host, P. pastoris (Zhan et al. 2014, 2015). IFTase (DFA III-forming) is efficiently expressed under the control of the formaldehyde dehydrogenase 1 promoter in P. pastoris and the maximum extracellular enzyme activity

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measured as 62.72 U mL1 (Zhan et al. 2014). Subsequently, multicopy P. pastoris exhibited a maximum extracellular DFA III-forming IFTase activity of 105.4 U mL1 in a 5 L fermenter (Zhan et al. 2015). Furthermore, Zhan et al. presumed that the IFTase-encoding gene from A. aurescens SK 8.001 could achieve higher level extracellular expression of IFTase by P. pastoris system when compared with the E. coli system and the wild type strain (Zhan et al. 2015). Extracellular secretory of DFA III-forming IFTase will efficently stimulus industial application of this enzyme and large-scale production of DFA III, with the convience of obtaine crud enzyme by the way of collecting the culture supernatant. A recombinant S. cerevisiae strain, containing the IFTase-encoding gene from A. aurescens SK 8.00, is constructed and equipped with optimal translational fusion partner for high efficiency production of DFA III (161.6 U mL1) (Ko et al. 2019a). Additionally, S. cerevisiae can remove other saccharides and not react to DFAs. S. cerevisiae is the most widely utilized gene-engineered strain, with the superiority of both safe for human and extracellular secretion of recombinant proteins (Ostergaard et al. 2000; Lian et al. 2018).

8.3.1.2

Properties of IFTases (DFA III-Forming)

Enzymatic properties of DFA III-forming IFTases have been deeply investigated. Generally, these properties are associated with the microbial source of the enzyme, as shown in Table 8.1. To date, the relative molecular weight of IFTases (DFA III-forming) is measured by SDS-PAGE and gel-filtration analysis and is ranged from 40 kDa to 150 kDa (Wang et al. 2015a). Thus, the reported DFA III-forming IFTases are suggested to be monomeric, homo-dimeric, and homo-trimeric proteins. Additionally, the corresponding crystal structure parameters of IFTase (DFA III-forming) from Bacillus sp. snu-7 proofed that this enzyme is a homo-trimeric protein (Jung et al. 2007). The native DFA III-forming IFTase from Arthrobacter sp. A-6 is proposed to be a homo-trimeric protein through gel-filtration analysis (Park and Choi 1996), while the recombinant one expressed by E. coli is presumed to be a monomeric enzyme (Kim et al. 2000a). The difference of this case might point that the aggregated state of protein is variable. All the reported IFTases (DFA III-forming) both native and recombinant ones prefer slightly acid condition for enzymatic reaction and exhibit optimum pH value which varies from 5.0 to 6.0. While the optimum temperature of the reported IFTases (DFA III-forming) has relative significant difference with each other, ranged from 35 to 70  C, and a majority of which up 50 to 70  C. The recombinant IFTase (DFA III-forming) from the Bacillus sp. snu-7 and native one from Arthrobacter sp. A-6 exhibits lowest and highest optimum temperature, 35 and 70  C, respectively (Kim et al. 2007). According to the reported investigations, IFTases (DFA III-forming) are not metal ion-dependent enzymes. Uchiyama et al. firstly reported that enzymatic activity of A. ureafaciens IFTase (DFA III-forming) was strongly inhibited by 1 mM HgC12 (Uchiyama et al. 1973). In a case study of Bacillus sp. snu-7 IFTase

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(DFA III-forming), 1 mM metal ions, Cu2+, Hg2+, and Fe2+ have shown significant inhibition to enzyme reaction (Kang et al. 1998). The enzyme activity of IFTase (DFA III-forming) produced by A. aurescens SK 8.001 was slightly inhibited by 1 mM Cu2+, Mg2+, and Ni2+ (Zhao et al. 2011b). Inhibited effect of divalent metal ions for IFTase (DFA III-forming) depends on the microbial sources and expression host. Thermal stability of the enzyme is a vital parameter for evaluating its industrial application. Currently, IFTase (DFA III-forming) from Arthrobacter sp. 161MFSha2.1 exhibits highest thermostability, and the recombinant enzyme is stable up to 80  C for 240 min of incubation (Wang et al. 2015b). In addition, among the reported IFTases (DFA III-forming), the ones from A. chlorophenolicus A6 (Zhu et al. 2018a), A. aurescens SK 8.001 (Zhao et al. 2011a), Arthrobacter sp. L68-1 (Haraguchi et al. 2005), Leifsonia sp. T88-4 (Haraguchi et al. 2006), and Arthrobacter sp. A-6 (Park and Choi 1996) are considered likewise as heat-stable enzymes. Specific activity of enzyme is another fundamental factor needed to be particularly considered for its industrial application. So far, IFTase (DFA III-forming) from Flavobacterium sp. LC-413 shows the highest specific activity, 5728 U mg1 (Cho et al. 1997), and the ones from Arthrobacter sp. A-6, Arthrobacter sp. B30–2, Arthrobacter sp. 161MFSha2.1, Bacillus sp. snu-7, and Nonomuraea sp. ID06A0189 have relatively high specific activity, with more than 1000 U mg1. IFTase (DFA III-forming) can hydrolyze inulin into DFA III as the main product, with the by-products of sucrose (GF) and fructo-oligosaccharide (FOS), such as 1-kestose (GF2), nystose (GF3), and fructofuranosylnystose (GF4) (Wang et al. 2015a). IFTases (DFA III-forming) from A. chlorophenolicus A6 (Zhu et al. 2018a), Arthrobacter sp. 161MFSha2.1 (Wang et al. 2015b), and A. ureafaciens D13-3 (Haraguchi 2010) produce the minor products as GF, GF2, GF3, and GF4. Minor products of IFTase (DFA III-forming) from A. aurescens SK 8.00 (Zhao et al. 2011b) and Bacillus sp. snu-7 (Kang et al. 1998) are determined as GF2, GF3, and GF4, while that of enzymes from Arthrobacter sp. A-6 (Park and Choi 1996), A. pascens T13-2 (Haraguchi et al. 2002), Arthrobacter sp. L68-1 (Haraguchi et al. 2005), and Leifsonia sp. T88-4 (Haraguchi et al. 2006) are confirmed as GF3 and GF4. The smallest substrates of the enzymes have been determined for comprehensive investigation of the IFTase (DFA III-forming). IFTase (DFA III-forming) can specifically catalyze β-(2, 1)-linked inulin. Therefore, FOS is chosen for the investigation of small substrates. IFTases (DFA III-forming) from A. chlorophenolicus A6 (Zhu et al. 2018a), Arthrobacter sp. 161MFSha2.1 (Wang et al. 2015b), A. ureafaciens D13-3 (Haraguchi 2010), A. aurescens SK 8.00 (Zhao et al. 2011b), and Nonomuraea sp. ID06A0189 (Pudjiraharti et al. 2011) catalyze GF3 as the smallest substrate. The one from Bacillus sp. snu-7 showed the smallest substrate as GF4 (Kang et al. 1998).

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IFTase for DFA I-Forming

IFTase (DFA I-forming) convert β-(2, 1) glycosidic bonds-connected inulin into DFA I, chemical structure of which is different from DFA III, with difference in one glycosidic bond. Researches on IFTase for DFA I-forming is obviously skimpy, compared with that of IFTase (DFA III-forming). In 1979, Tanaka et al. first reported DFA I formed by enzymatic reaction of Aspergillus fumigatus (Tanaka et al. 1979) and proposed that the mycelium autolysate could hydrolyze inulin to produce DFA I, with inulin-containing culture medium of A. fumigatus. An enzyme was purified from A. fumigatus to produce DFA I from inulobiose instead of inulin (Matsuyama et al. 1982). IFTases (DFA I-forming) are found in various microbial sources, with the increasing investigation. The crude enzyme of A. globiformis S14-3, isolated from the soil and cultured with 0.3% inulin, can transform inulin into DFA I (Seki et al. 1988). In addition, the enzyme was purified and characterized (Seki et al. 1989). The gene encoding IFTase (DFA I-forming) from A. globiformis S14-3 was cloned and expressed by E. coli under the control of the lac promoter of pUC119 (Haraguchi et al. 1995; Haraguchi et al. 1998). Subsequently, another 11 strains that form DFA I have been reported, including Streptomyces sp. MCI-2524 (Kushibe et al. 1993), Arthrobacter sp. MCI2493 (Ueda et al. 1994), A. ureafaciens A51-1 (Haraguchi et al. 2003a), A. pascens A62-1 (Haraguchi et al. 2003b), Arthrobacter sp. B69–5 (Haraguchi and Ohtsubo 2006), C. clostridioforme AGR2157 (Yu et al. 2015), S. davawensis SK39.001 (Zhu et al. 2016), S. peucetius subsp. caesius ATCC 27952 (Cheng et al. 2020), and Nocardioides family (Nocardioides sp. JS614, N. luteus, N. bacterium Broad-1) (Cheng et al. 2017). Since 2015, genes of six strains, encoding IFTase (DFA I-forming), were entirely expressed by E. coli. According to the analysis of gel filtration, the native IFTases (DFA I-forming) are considered to be monomers and dimers, and the recombinant ones are suggested to be homo-trimeric proteins. The biochemical properties of IFTase (DFA I-forming) are similar with that of IFTase (DFA III-forming). The optimum pH value ranges from 5.5 to 6.5, and the optimum temperatures were measured to be 40 to 65  C. Enzymes from A. ureafaciens A51-1 (Haraguchi et al. 2003a), Nocardioides sp. JS614 (Cheng et al. 2017), C. clostridioforme AGR2157 (Yu et al. 2015), and S. peucetius subsp. caesius ATCC 27952 (Cheng et al. 2020) have the relatively high thermostability. The recombinant one from C. clostridioforme AGR2157 is the most thermostable, which remains half of initial activity after 240 min of incubation at 80  C, and it has the highest specific activity among the reported enzymes, determined as 2,076 U mg1 (Yu et al. 2015). Investigations of metal ions on enzymatic activity demonstrate that IFTase (DFA I-forming) are not metal ion-dependent enzyme. Kushibe et al. firstly reported that Cu2+ inhibited the enzymatic activity of the purified enzyme from Streptomyces sp. MCI-2524 (Kushibe et al. 1993). In another case study, Fe2+ showed strong inhibition, and Ba showed a slight promotion to the recombinant enzyme from S. peucetius subsp. caesius ATCC 27952 (Cheng et al. 2020). All the reported

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native IFTase (DFA I-forming) convert inulin into DFA I, with GF3 and GF4 as the minor byproduct. While the recombinant ones from C. clostridioforme AGR2157 (Yu et al. 2015), S. davawensis SK39.001 (Zhu et al. 2016), and S. peucetius subsp. caesius ATCC 27952 (Cheng et al. 2020) produce GF, GF2, GF3, and GF4 as the minor by-product. In addition, GF and GF2 are the minor product of recombinant enzymes from Nocardioides family (Cheng et al. 2017).

8.3.3

Enzyme for DFA V-Forming

Until now, the enzyme for forming DFA V has not been characterized yet. There is only one investigation on biological synthesis of DFA V. In 1991, Matsuyama et al. found A. fumigatus can transfer inulin into a novel difructose anhydride under the condition of inulin as a sole carbon source and named DFA V as a trivial name (Matsuyama et al. 1991).

8.3.4

LFTase for DFA IV-Forming

Presently, DFA IV can be bio-transformed from levan by LFTase (DFA IV-forming). Tanaka et al. firstly reported that DFA IV was formed by crude enzyme from A. ureafaciens which was cultured in a culture medium containing bacterial levan from Bacillus mesentericus as a sole carbon source (Tanaka et al. 1981). Additionally, the crude enzyme could not catalyze inulin, inulobiose, sucrose, and levan-type fructan (levanbiose and levantriose) to produce DFA IV. The one from A. ureafaciens was subsequently purified and characterized (Tanaka et al. 1983). Later on, another five microbial sources of DFA IV-forming have been identified, including Arthrobacter sp. No.11-E (Murakami et al. 1993), A. nicotinovorans GS-9 (Saito et al. 1997; Takesue et al. 2009), A. ureafaciens K2032 (Song et al. 2000; Lee et al. 2001; Kim et al. 2005), Microbacterium sp. AL-210 (Cha et al. 2001; Yang et al. 2002; Hwang et al. 2009), and Arthrobacter oxydans J17-21 (Jang et al. 2003). LFTases (DFA IV-forming) from A. nicotinovorans GS-9 (Saito et al. 1997), A. ureafaciens K2032 (Lee et al. 2001; Kim et al. 2005), and Microbacterium sp. AL-210 are expressed by E. coli (Yang et al. 2002; Hwang et al. 2009). The heterologous expression of LFTase (DFA IV-forming) from A. nicotinovorans GS-9 is reported, with B. subtilis as host (Takesue et al. 2009). Moreover, Ko et al. reported the eukaryotic expression of LFTase (DFA IV-forming) from A. ureafaciens by S. cerevisiae (Ko et al. 2019b). The recombinant enzyme from A. ureafaciens K2032 exhibited highest extracellular enzymatic activity accumulated to 46,000 U mL1 (Lee et al. 2001). The specific activity of the purified enzyme from Microbacterium sp. AL-210 was relatively high and determined to be 23,202 U mg1 (Cha et al. 2001). Heterologous expression by B. subtilis and S. cerevisiae contributes the utilizations of DFA IV in food industrial applications.

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Among the reported LFTases (DFA IV-forming), the ones from A. ureafaciens (Tanaka et al. 1983) and A. ureafaciens K2032 (Song et al. 2000) were proposed to be dimers, and the ones from A. nicotinovorans GS-9 (Saito et al. 1997) and Microbacterium sp. AL-210 (Cha et al. 2001) were proposed to be monomers. Normally, LFTases (DFA IV-forming) exhibit maximal enzymatic activity in a slightly acidic or neutral environment (5.8–7.0). The optimum temperature of LFTases (DFA IV-forming) varies from 40 to 50  C. The effects of metal ions on the enzymatic activity have been discussed. Song et al. reported that Mn2+, Fe2+, and Hg2+ strongly inhibited the enzymatic activity, while Na+ and Ca2+ increase the activity of enzyme from A. ureafaciens K2032 (Song et al. 2000). The activity of LFTases (DFA IV-forming) from Microbacterium sp. AL-210 was inhibited by Fe2+ and Ag3+ and activated by Mg2+ (Cha et al. 2001). The activity of the purified enzyme from Arthrobacter oxydans J17-21 was inhibited by Fe2+ and Ag+ and activated by Ca2+ (Jang et al. 2003). Additionally, the thermostability of LFTases (DFA IV-forming) was obviously lower than that of reported IFTases.

8.4 8.4.1

Structural Analysis and Catalytic Mechanisms Overall Structures

According to the category of carbohydrate-active enzymes database (CAZy, http:// www.cazy.org/Glycoside-Hydrolases.html), LFTase (DFA IV-forming) is classified as GH32 enzymes, while IFTases and DFA IIIase belong to GH91 enzymes. Additionally, the characteristic structure status of GH32 family enzymes is fiveblade β-propeller, while that of GH91 family proteins is β-helix. With obviously different structure, the products of IFTases and LFTases are isomers with different glycosidic linkages. Thus, investigations of the crystal structures and catalytic mechanisms of these two kinds of enzymes are meaningful and interesting. To date, three references reported the crystal studies on IFTases and LFTase (DFA IV-forming) (Fig. 8.2). As far as structural investigations of IFTases, only the IFTases (DFA III-forming) from A. globiformis and Bacillus sp. snu-7 have been determined the three-dimensional structure by X-ray diffraction (Momma et al. 2003; Jung et al. 2007). In 2012, the crystal structures and functional basis for substrate specificity of LFTase (DFA IV-forming) from A. ureafaciens have been reported (Park et al. 2012). Momma et al. first reported the crystal of IFTase (DFA III-forming) from A. globiformis and the crystal parameters and data collection statistics of the enzyme. They obtained the crystals of the wild-type form and the selenomethionine derivative of A. globiformis IFTase (DFA III-forming), with a 1.5 and 2.3 Å resolution, respectively (Momma et al. 2003). The space group of crystal of the wild-type form belonged to R32, while that of the selenomethionine derivative belonged to C2. Additionally, they proposed that the R32 crystal contained one molecule in the asymmetric unit and three molecules in the asymmetric unit of the C2 crystal.

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Fig. 8.2 The crystal structures of IFTase (DFA III-forming) from Bacillus sp. snu-7 and LFTase (DFA IV-forming) from A. ureafaciens. (1) The overall and apo-form structure of IFTase (DFA III-forming) from Bacillus sp. snu-7 (PDB: 2INU), (2) its monomeric structure, and (3) its holoform with difructosaccharide (PDB: 2INV). (4) The overall and apo-form structure of LFTase (DFA IV-forming) from A. ureafaciens (PDB: 4FFF) with four monomers in the asymmetric unit, (5) its monomeric structure, (6) its holo-form with sucrose (PDB: 4FFH), (7) its holo-form with levanbiose (PDB: 4FFI), (8) its holo-form with DFA IV (PDB: 4FFG)

However, the relative structural information did not submit to the Protein Data Bank (http://www.rcsb.org/pdb/results), and the further crystallographic analysis is still not reported. Until 2007, Jung et al. determined the structure of IFTase (DFA III-forming) from Bacillus sp. snu-7 and released publicly in the PDB (Jung et al. 2007). The apo-form (ligand-free) (PDB: 2INU) and holo-form (ligand-bound) (PDB: 2INV) of the IFTase (DFA III-forming) from Bacillus sp. snu-7 structures were determined and collected at the same resolution of 1.8 Å. Additionally, the complexed ligand was β2,1-linked tetrafructosaccharide. The two structures belonged to the same C2 space group with three molecules in the asymmetric unit, which indicated that IFTase (DFA III-forming) from Bacillus sp. snu-7 was a homo-trimeric protein. Each monomer formed a right-handed parallel β-helix that consisted 13 helical turns of a right-handed coil, monomers shaped like a cylinder. Specifically, each turn composed of β-strands, connecting by loops. The adjacent parallel β-strands interacted by hydrogen bonding, which stabilized the structure and formed the surface of enzyme. The three monomers of the IFTase (DFA III-forming) from Bacillus sp. snu-7 intertwined to form a trimer. The crystal structure of LFTase (DFA IV-forming) from A. ureafaciens have been resolved (Park et al. 2012). The apo-form (PDB: 4FFF) of this protein’s structure was collected at a resolution of 2.57 Å. In addition, three holo-forms in complex with sucrose (PDB: 4FFH), levanbiose (PDB: 4FFI), and DFA IV (PDB: 4FFG) were collected at resolutions of 2.2, 2.3, and 2.3 Å, respectively. All of the structures

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belonged to the space group of P212121 and had four monomers in the asymmetric unit. However, LFTase (DFA IV-forming) from A. ureafaciens is suggested to be monomers in solution through size exclusion chromatography. Overall, LFTase (DFA IV-forming) from A. ureafaciens is structurally consisted of two domains, N-domain (a five-bladed β-propeller) and C-domain (a β-sandwich), and those structural characters are identical to those of other GH32 family members. The identified catalytic pocket shapes in an elongated shape and is composed of a cluster of conserved residues which have various interactions with ligands. Structure of LFTase in complex with sucrose bounds one molecular of sucrose in N-domain and the one in C-terminal region, respectively. Interestingly, the complex of LFTase and levanbiose exhibits the same ligand-binding sites as that of the complex with sucrose and shows additional bounding on the surface of N-domain. DFA IV only binds at the C-domain of LFTase (DFA IV-forming) from A. ureafaciens structure.

8.4.2

Catalytic Mechanisms

The possible catalytic mechanisms for IFTase and LFTase are proposed, with the relative structural and functional analyses. IFTase (DFA III-forming) from Bacillus sp. snu-7 performs an intramolecular fructosyl transfer with an inverting mechanism (Jung et al. 2007), while LFTase (DFA IV-forming) from A. ureafaciens carries out an intramolecular fructosyl transfer reaction with the retaining mechanism (Park et al. 2012). According to the crystallographic analysis associated with site-directed mutagenesis of IFTase (DFA III-forming) from Bacillus sp. snu-7, the substrate-binding sites are suggested to be located at the interface of two monomers. The strictly reserved residue, Glu244, functions as a general base to activate the terminal fructosyl which serves as the acceptor. Another critical residue is Asp233, which probably coordinates the configuration of the donor fructosyl molecular for catalysis. Based on the holo-form of the enzyme structure, only two fructose moieties of the tetrafructosaccharide are captured, with the terminal fructosyl at +2 subsite and the other one at +1 subsite. The fructosyl of the ligand at +2 subsite binds at the inner site of catalytic pocket, while that of at +1 subsite closes to the entrance. The substratebinding pocket is composed of the residues, which are completely or highly conserved and from the two neighbored monomers. The interactions that maintain the catalytic pocket are hydrogen bonds and van der Waals’ interactions. Specifically, hydrogen bonds are directly generated among the fructose moieties, residues, and water molecules, which play critical roles in catalysis. LFTase (DFA IV-forming) from A. ureafaciens is composed of β-propeller N-domain and β-sandwich C-domain. The N-domain functioned as a catalytic domain, and the C-domain is possibly involved in carbohydrate recognition. LFTase (DFA IV-forming) has a “catalytic triad” at the center of the N-domain, which conserved with that of GH32 and GH68 family members. Asp54, Glu236, and Asp186 consist of the characterized “catalytic triad” and are functioned as

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nucleophile, acid/base catalyst, and intermediate stabilization, respectively. The conserved loop region within smaller side chain residues among LFTase (DFA IV-forming) provides the structural basis for the exo-type cleavage and formation of DFA IV. In the pocket of levanbiose-bounded structure, the reducing fructose moiety and a nonreducing terminal fructose moiety are considered as 1 and 2 subsites, respectively. The fructose at 1 subsite is located at the adjacent of nucleophile (Asp54). The β-2,1-glycosidic bond is cleaved by nucleophile on the anomeric carbon of the fructose at 1 subsite. The hydroxyl group of the fructose at 2 subsite is deprotonated by Glu236 and finally generates DFA IV with a new glycosidic linkage. It is the shape and dimension of the catalytic pocket which decides on the substrate specificity and production of DFA IV.

8.4.3

Molecular Modifications

Based on the structural information, molecular modifications are mainly carried out to improve the enzymatic activity and thermal stability of enzymes. At present, rational design, through site-directed mutagenesis, is the predominant method for molecular modifications of IFTases and LFTases (DFA IV-forming). In general, the current molecular modifications of IFTases are mainly to increase the activity and thermal stability of the enzyme. Yu et al. constructed the structural model of IFTase (DFA III-forming) from Arthrobacter sp. 161MFSha2.1 through homology modeling, with the structure of IFTase (DFA III-forming) from Bacillus sp. snu-7 as the template, to improve its thermostability (Yu et al. 2016a). A double mutation (S309T/S333T) significantly increased the thermostability, with the melting temperature (Tm) increased by 5  C and the half-life at 55  C increased by 5 h. Thereafter, site-directed mutagenesis was designed for enhancing the catalytic activity of IFTase (DFA I-forming) from S. davawensis (Yu et al. 2017). Two positive mutants, G121A/T122L (two-sites) and G236S/G281S/A257S/T313S/ A314S (five-sites), were obtained. Compared with the activity of the wild-type enzyme, that of the two-site and five-site mutations increased by 48.7% and 40.5%, respectively. Additionally, the Tm value of the two-site and five-site mutations was measured to be 4.5  C and 3.2  C higher, respectively, than that of the wild-type enzyme. Site-directed mutagenesis studies mainly focused on revealing the critical residues of LFTases (DFA IV-forming), before resolving the crystal structures of LFTase (DFA IV-forming) from A. ureafaciens. Three highly conserved residues, Asp63, Asp195, and Glu245, are expected to be crucial in catalysis of LFTase (DFA IV-forming) from Microbacterium sp. AL-210 (Sung et al. 2003). Moon et al. reported that the residue, Asn85, is critical for transfructosylation activity of LFTase from Microbacterium sp. AL-210, through error-prone PCR mutagenesis process (Moon et al. 2008).

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Biological Production and Application of DFAs

Biological production of DFAs by enzymatic strategy has been investigated and commercially utilized. Industrial production of DFA III has been achieved by the Shimizu Factory (Tokachi district, Hokkaido, Japan) of Nippon Beet Sugar Mfg. Co., Ltd., through enzymatic bioconversion of inulin by Arthrobacter sp. H65-7 IFTase (DFA III-forming) (Kikuchi et al. 2009). To large-scale and practical production of DFAs, investigations with various enzymatic substrates have increasingly attracted attention, including inulin- or levan- contained raw materials. Kikuchi et al. reported the economical and industrial production of DFA III by Arthrobacter sp. H65-7 IFTase, and the crude inulin extracted from chicory roots was treated as substrate (Kikuchi et al. 2009). The recombinant S. cerevisiae, with the ability of hyper-secreting IFTase (DFA III-forming) from A. aurescens SK 8.001, was directly fermented with crude extract of Jerusalem artichoke tuber powder (Ko et al. 2019a). Recently, one-pot conversion of DFAs has been researched. Enzymatic production of DFA III and DFA IV from sucrose was achieved. DFA III was prepared from sucrose by coupled the fructosyltransferase from A. niger AS0023 and IFTase (DFA III-forming) from A. aurescens SK 8.001, and the DFA III yield is 100 mg g1 (DFA III weight/sucrose weight) (Hang et al. 2013). Inulosucrase from Lactobacillus johnsonii NCC533 formed inulin from sucrose, and IFTase from Arthrobacter sp. 161MFSha2.1 was used for producing DFA III (Yu et al. 2016b). DFA IV was efficiently converted by A. nicotinovorans GS-9 LFTase (DFA IV-forming) from the microbial levan which was formed from sucrose by levansucrase from Serratia levanicum NN (Kikuchi et al. 2010). The crude DFA IV was directly biosynthesized from sucrose by co-fermenting of two recombinant yeasts, which were used for secreting levansucrase and LFTase (DFA IV-forming) (Ko et al. 2019b). Immobilization of enzymes has been widely investigated and applied for industrial applications. Jahnz et al. reported that immobilized IFTase (DFA III-forming) from Arthrobacter sp. Buo141 had an activity of 196 U mg1 (Jahnz et al. 2001). The IFTase from A. aurescens can be used for efficient production of DFA III, through embedded in curdlan-based mesoporous silica microspheres, and this immobilized enzyme has a good reusability and excellent storage stability (Liu et al. 2019). LFTase from A. ureafaciens K2032 exhibited a high activity for DFA IV formation, and the enzyme remained approximately 60% of its initial-enzymatic activity after 20 cycles utilization (Lim et al. 2001). Additionally, an ultrafiltration membrane bioreactor (UFMB) system was used for the highly efficient production of DFA III from inulin by IFTase (DFA III-forming) from A. aurescens SK8.001 (Hang et al. 2012b). Compared to the batch reactor, the UFMB system contributed to recycling of the enzymes, continuous preparation, increasing productivity and purity of DFA III, and eliminating product inhibition effect (Hang et al. 2012a, b). An enzymatic membrane reactor (EMR) system, equipped with nanofiltration and ultrafiltration membranes, was applied for bioconversion of high concentration of DFA III from inulin. The integrated EMR

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system improved the DFA III concentration to approximately 400 g L1 (Hang et al. 2015). In Japan, a DFA III-contained health food, named “Twintose,” has been put on the Japanese market by a FANCL company, since 1994. And the slogan claimed that the Twintose could stimulate the absorption of minerals (Zhu et al. 2018b). In addition, Jiangnan University and the Jiangsu Liangfeng food company proposed the project “Functional sweeteners-DFA III bio-processing key technology and industrialization.” Based on the provided bio-processing technology, the annual production of DFA III could reached to 100 tons (Hang 2017). In another study, DFAs are the by-products during the reaction and act as slow-converting and protective intermediates that increase the yield of fructose to 5-hydroxymethylfurfural which is an intermediate to high-energy chemical compounds furans (Svenningsen et al. 2018).

8.6

Concluding Remarks

Sucrose, as a high-calorie sweetener, is commonly utilized in food industry, and excessive intake of sucrose is related to the metabolic syndrome which clinically increase the risk for several diseases, including type 2 diabetes, obesity, cardiovascular disease, and so on (Eckel et al. 2005; Bray and Popkin 2014; Malik et al. 2010). Therefore, DFAs have attracted cumulative attention, as the promising substitute of table sugar. Benefit from the biotechnology, four types of DFAs, including DFA III, DFA IV, DAF I, and DFA V, have been characterized by enzymatic methods or microbial reaction. The most investigated ones are DFA III and DFA IV, from bio-formation to physiological functions and applications. Interestingly, fructotransferases catalyzed inulin for synthesis of DFA III and DAF I, belonging to GH91 family, but LFTases (DFA IV-forming) are classified as the members of GH32 family. Based on the structural information, the former ones and the latter ones adopt the different catalytic mechanisms, inverting and retaining, respectively. Although the IFTase (DFA I-forming) has a relative high homology with IFTase (DFA III-forming), its catalytic mechanism still needs to be clarified, as extremely similar between DFA I and DFA III. Additionally, investigations on DFA V are relatively rare. The specific DFA V-forming enzymes are required to be identified for the further investigations, synthesis, and applications. To better serve for human health, the basic researches and the practical applications of DFAs deserve attentions.

References Arribas B, Suárez-Pereira E, Ortiz Mellet C, García Fernández JM, Buttersack C, RodríguezCabezas ME, Garrido-Mesa N, Bailon E, Guerra-Hernández E, Zarzuelo A, Gálvez J (2010)

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Teramura M, Wynn S, Reshalaitihan M, Kyuno W, Sato T, Ohtani M, Kawashima C, Hanada M (2015b) Supplementation with difructose anhydride III promotes passive calcium absorption in the small intestine immediately after calving in dairy cows. J Dairy Sci 98(12):8688–8697 Tomita F, Yokota A, Kasai T, Ham H, Sayama K (1999) An efficient production of DFA III and its potential utility as a physiologically functional food. In: Food for Health in the Pacific Rim: 3rd International Conference of Food Science and Technology. Wiley Online Library, pp 353–362 Trabs K, Kasprick N, Henle T (2011) Isolation and identification of Di-D-fructose dianhydrides resulting from heat-induced degradation of inulin. Eur Food Res Technol 233(1):151–158. https://doi.org/10.1007/s00217-011-1507-8 Uchiyama T (1975) Action of Arthrobacter ureafaciens inulinase II on several oligofructans and bacterial levans. Biochim Biophys Acta (BBA) Enzymol 397(1):153–163 Uchiyama T (1983) Formation of Di-d-fructose anhydride III from inulin by the root of Lycoris radiata Herbert. Agric Biol Chem 47(2):437–439. https://doi.org/10.1080/00021369.1983. 10865656 Uchiyama T, Niwa S, Tanaka K (1973) Purification and properties of Arthrobacter ureafaciens inulase II. Biochim Biophys Acta (BBA) Enzymol 315(2):412–420 Ueda M, Sashida R, Morimoto Y, Ohkishi H (1994) Purification of inulin fructotransferase (DFA I-producing) from Arthrobacter sp. MCI2493 and production of DFA I from inulin by the enzyme. J Agric Chem Soc Japan 58(3):574–575 Wang X, Yu S, Zhang T, Jiang B, Mu W (2015a) From fructans to difructose dianhydrides. Appl Microbiol Biotechnol 99(1):175–188. https://doi.org/10.1007/s00253-014-6238-x Wang X, Yu S, Zhang T, Jiang B, Mu W (2015b) Identification of a recombinant inulin fructotransferase (difructose dianhydride III forming) from Arthrobacter sp. 161MFSha2.1 with high specific activity and remarkable thermostability. J Agric Food Chem 63 (13):3509–3515 Yang SJ, Park NH, Lee TH, Cha J (2002) Expression, purification and characterization of a recombinant Levan fructotransferase. Biotechnol Appl Biochem 35(3):199–203. https://doi. org/10.1111/j.1470-8744.2002.tb01189.x Yokota A, Enomoto K, Tomita F (1991a) Purification and properties of an inulin fructotransferase (depolymerizing) from Arthrobacter sp. H65-7. J Ferment Bioeng 72(4):262–265. https://doi. org/10.1016/0922-338X(91)90160-I Yokota A, Hirayama S, Enomoto K, Miura Y, Takao S, Tomita F (1991b) Production of inulin fructotransferase (depolymerizing) by Arthrobacter sp. H65-7 and preparation of DFA III from inulin by the enzyme. J Ferment Bioeng 72(4):258–261 Yu S, Wang X, Zhang T, Jiang B, Mu W (2015) Characterization of a thermostable inulin fructotransferase from clostridium clostridioforme AGR2157 that produces difructose dianhydride I from inulin. J Mol Catal B Enzym 120:16–22. https://doi.org/10.1016/j. molcatb.2015.06.012 Yu S, Wang X, Zhang T, Jiang B, Mu W (2016a) Probing the role of two critical residues in inulin Fructotransferase (DFA III-producing) Thermostability from Arthrobacter sp. 161MFSha2.1. J Agric Food Chem 64(31):6188–6195. https://doi.org/10.1021/acs.jafc.6b02291 Yu S, Zhu Y, Zhang T, Jiang B, Mu W (2016b) Facile enzymatic production of difructose dianhydride III from sucrose. RSC Adv 6(105):103791–103794. https://doi.org/10.1039/ c6ra23352j Yu S, Zhang Y, Zhu Y, Zhang T, Jiang B, Mu W (2017) Improving the catalytic behavior of DFA I-forming inulin Fructotransferase from Streptomyces davawensis with site-directed mutagenesis. J Agric Food Chem 65(34):7579–7587. https://doi.org/10.1021/acs.jafc.7b02897 Yu S, Shen H, Cheng Y, Zhu Y, Li X, Mu W (2018) Structural and functional basis of difructose anhydride III hydrolase, which sequentially converts inulin using the same catalytic residue. ACS Catal 8(11):10683–10697 Zhan R, Mu W, Jiang B, Zhou L, Zhang T (2014) Efficient secretion of inulin fructotransferase in Pichia pastoris using the formaldehyde dehydrogenase 1 promoter. J Ind Microbiol Biotechnol 41(12):1783–1791. https://doi.org/10.1007/s10295-014-1516-2

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Zhan R, Mu W, Jiang B, Li Y, Zhou L, Zhang T (2015) High-level extracellular expression of inulin fructotransferase in Pichia pastoris for DFA III production. J Sci Food Agric 95(7):1408–1413. https://doi.org/10.1002/jsfa.6931 Zhao M, Mu W, Jiang B, Hang H, Zhou L, Zhang T (2011a) Cloning and extracellular expression of inulin fructotransferase from Arthrobacter aurescens SK 8.001 in E. coli. J Sci Food Agric 91 (15):2715–2721. https://doi.org/10.1002/jsfa.4582 Zhao M, Mu W, Jiang B, Zhou L, Zhang T, Lu Z, Jin Z, Yang R (2011b) Purification and characterization of inulin fructotransferase (DFA III-forming) from Arthrobacter aurescens SK 8.001. Bioresource Technology 102(2):1757–1764. https://doi.org/10.1016/j.biortech. 2010.08.093 Zhao M, Jiang B, Hang H, Fang Y, Jiang F, Phillips GO (2013) Efficient induction of inulin fructotransferase by inulin and by difructose anhydride III in Arthrobacter aurescens SK 8.001. Eur Food Res Technol 236(6):991–998. https://doi.org/10.1007/s00217-013-1962-5 Zhu Y, Yu S, Huang D, Zhang T, Jiang B, Mu W (2016) Identification of a novel DFA I-producing inulin fructotransferase from Streptomyces davawensis. Int J Biol Macromol 92:723–730. https://doi.org/10.1016/j.ijbiomac.2016.07.092 Zhu Y, Wang X, Yu S, Zhang W, Zhang T, Jiang B, Mu W (2018a) Bioconversion of inulin to difructose anhydride III by a novel inulin fructotransferase from Arthrobacter chlorophenolicus A6. Process Biochem 75:130–138. https://doi.org/10.1016/j.procbio.2018.07.003 Zhu YY, Yu SH, Zhang WL, Zhang T, Guang CI, Mu WM (2018b) Recent advances on biological production of difructose dianhydride III. Appl Microbiol Biotechnol 102(7):3007–3015. https:// doi.org/10.1007/s00253-018-8834-7

Chapter 9

Characteristics of Levansucrase and Its Application for the Preparation of Levan and Levan-Type Oligosaccharides Wei Xu, Wenli Zhang, and Wanmeng Mu

9.1

Levan and Its Existing Resources

Levan is a homopolysaccharide consisting of one glucose and numerous repeated fructoses via a β-(2, 6) linkage. It is a non-structural carbohydrate and exists in plant, fungi, yeast, and bacteria (Arvidson et al. 2006). For plants, levan can serve as an energy reservoir to protect them from hostile environment such as soil salinity, drought, and extreme temperatures (Ritsema and Smeekens 2003). For bacteria, levan can favor the formation of exopolysaccharides (EPS) matrix and microbial biofilm (Franken et al. 2013). Notably, the formed biofilm could have different functions when presented in different strains. For instance, they can shield the microorganism such as Bacillus subtilis, from desiccation as the water level changes, help glue cells in a beneficial environment, and protect the community from predatory organisms (Dogsa et al. 2013). Secondly, for the plant pathogens such as Erwinia amylovora and Pseudomonas syringae (Koczan et al. 2009; Mehmood et al. 2015), levan as a biofilm component also contributes to the virulence. In the case of endophyte Gluconacetobacter diazotrophicus, the levan-based biofilm acts as an oxygen difussional barrier supporting the microaerobic conditions required for nitrogen fixation (Hernández et al. 2000). Last but not least, the biofilms of levan constitutes a nutrient reservoir that could be alternatively utilized as energy source by bacteria when they are under starvation conditions.

W. Xu · W. Zhang State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China e-mail: [email protected] W. Mu (*) State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_9

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Levan can be synthesized by a limited plant species and a wide variety of microorganisms. The plants Agropyron cristatum, Dactylis glomerata, and Poa secunda are well known as levan-producing plants, wherein the sucrose:fructan 6-fructosyltransferase (6-SFT; EC 2.4.1.10), also known as sucrose:sucrose 6-fructosyltransferase (6-SST), is involved in the synthesis of levan (Srikanth et al. 2015a). The enzyme is able to use both fructan molecule and sucrose as the acceptor substrate to form the β-(2, 6) fructosyl linkage. Alone, 6-SFT can synthesize linear levan chains directly from sucrose via the intermediate 6-kestose (He et al. 2015). In bacteria, levansucrase (LSase, EC 2.4.1.10) is the key enzyme responsible for the production of levan through a transfructosylation (Hernández et al. 1999). Many different types of strains including Lactobacillus gasserii, Lactobacillus johnsonii, Bacillus subtilis, Aerobacter levanicum, and Streptococcus salivarius are able to produce LSase. LSase is constitutive and endocellularly formed in A. levanicum but can also be extracellularly produced in the case of B. subtilis (Abdel-Fattah et al. 2005), indicating there are two types of excretion mechanism for LSase among different microorganisms. Firstly, LSase is accumulated in periplasm and then excreted into the outer member by the two different mechanisms. For the grampositive bacteria like Bacillus stearothermophilus, B. subtilis, and Bacillus amyloliquefaciens, two steps including signal peptide cleavage and protein folding are required for the excretion of LSase (Donot et al. 2012). By contrast, a one step for LSase secretion into the outer environment triggered by signal peptide is found for the gram-negative microorganisms such as Erwinia amylovora, Glycinea, Rahnella aquatilis, Zymomonas mobilis, and P. syringae pv. phaseolicola (Avigad et al. 1956).

9.2

Chemical Structure of Levan and Its Physicochemical Property

The property of polymers is distinguished largely depending on their different structures. Herein, a good knowledge of polymer structure is of significance in both the scientific and practical application fields. Levan employs the repeated β(2, 6) fructosyl linkage as its backbone and occasionally branches at the β-(2, 1) point (Fig. 9.1). For plants-derived levan, the degree of polymerization (DP) is limited to 101–102 fructosyl units, whereas the bacterial levan has a much higher DP (>104) and molecular mass. Notably, the molecular mass of levan could largely affect its property and industrial application.

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Fig. 9.1 Chemical structure of levan polysaccharide

9.2.1

Solubility

Levan does not swell in water and has an excellent water solubility compared to the other polysaccharides. It can be completely soluble in hot water but show a different solubility in cold water. It is insoluble in the most of organic solvents including methanol, ethanol, n-propanol, isopropanol, methylethylketone, acetone, toluene, ethyl lactate, methyl caprylate/caprate, methylpalmitate/oleate, d-limonene, propylene carbonate, ethylene carbonate, 1-vinyl-2-pyrrolidone, methoxypolyethylene glycol, polyethylene glycol, dimethyl formamide, ethoxyethyl acetate, acetic anhydride, and furfuryl alcohol (Patel and Patel 2011). More importantly, levan could hardly dissolve in oil, although many papers have described levan as oil-soluble.

9.2.2

Viscosity

The intrinsic viscosity [η] is usually used to describe the solute’s contribution to the viscosity of its solution. Early studies found the intrinsic viscosity of levan in its aqueous solution was between 0.07 and 0.18 dL/g for the molecular weight ranging from 16  106 to 24  106 Da (Newbrun and Baker 1968), suggesting it has an extreme low intrinsic viscosity compared to that of the other high molecular weight molecules. Harada et al. found that at the pH of 4–11, the viscosity of Z. mobilis levan was stable, but the viscosity could be affected by increasing salt concentrations as well as the temperature (70  C) (Vina et al. 1998). Ullrich et al. described the 50% levan solution as a kind of chewing gum. Xu et al. also investigated the apparent viscosity of levan produced by the recombinant levansucrase from Brenneria sp. At a low concentration (3%) of levan aqueous solution, there was no change about the apparent viscosity at the shear rates between 1 and 100 1/s, indicating an essentially Newtonian behavior. At 6, 9, and 12% concentrations, the solution showed a shear

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thinning behavior (non-Newtonian pseudoplastic fluid), showing a decent apparent viscosity with an increasing shear rate (Xu et al. 2018a).

9.2.3

Tensile Strength

The hybrid structure of levan results in its cohesive strength, and the large number of hydroxyl groups enables levan to form adhesive bonds with a variety of substrates. The tensile strength of levan on aluminum was determined to be 500–1500 psi, which is higher than that of many petrochemical based adhesives. Besides, levan is also regarded as a “green adhesive” because the dried levan could be washed easily with water.

9.2.4

Thermostability and Safety

The melting temperature of levan is around 225  C, suggesting a high heat stability (Ling 2009). Also, levan meets all of the safety criteria from Human Repeated Insult Patch Test (HRIPT), Chorioallantoic Membrane Vascular Assay (CAMVA), and Agar Diffusion Test, which implies a totally non-toxicity and ocular non-irritating of levan (Corp. 2015).

9.3

Biological Production of Levan

Compared to inulin, the yield of levan extracted from plants is much lower; at the same time, biological production of levan by microbial fermentation or enzymatic conversion is highly promising both on the industrial feasibility and product yield (González-Garcinuño Al et al. 2018). As abovementioned, levan exists widely in microorganisms, including the gram-positive and gram-negative bacteria. Biological production of levan by gram-positive microorganism was summarized in Table 9.1. So far, the genera including Bacillus and Panebacillus have been the most intensively studied gram-positive bacteria for the production of levan.

9.3.1

Biological Production of Levan by Gram-Positive Microorganisms

The gram-positive bacteria B. subtilis (natto) Takahashi from soybean has been intensively studied for the levan production. Under the optimized conditions

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Table 9.1 Production of levan by gram-positive microorganisms Initial pH 6.0

Temperature ( C) 25–40

Speed (rpm) 150–200

Sucrose (g/L) 200

5.6–5.8

37

150

200

7.4

37

150

250

56 (48 h)

7.0

37

175

250

B. subtilis (natto) CCT 7712

NR

37

NR

400

61 (24) batch 100 (120) fed-batch 111 (16 h)

B. methylotrophicus SK 21.002

6.0

37

NR

300

100 (16 h)

B. lentus V8

6.5

30

150

250

B. licheniformis NS032

7.0 7.4

NR NR

NR NR

196.8 397.6

57.95 (60 h) from sucrose 49.86 (60 h) from sugar cane molasses 47.8 99.6

B. amyloliquefaciens NK-ΔLP

6.0

37

180

250

22.6 (48 h)

B. amyloliquefaciens NK-Q-7

6.0

37

180

250

31.1 (48 h)

M. laevaniformans PTCC 1406

6.0

NR

NR

200

P. polymyxa EJS-3

8.0

24

150

188.2

48.9 (48 h, from sucrose) 10.48 (48 h, from 250 g/L syrup) 35.26 (60 h)

Strains Bacillus subtilis (natto) Takahashi B. subtilis (natto) Takahashi (immobilized) B. subtilis (natto) Takahashi B. subtilis (natto) Takahashi

NR Not reported

Production (time) 49.4 g/L (21 h) 70.6 g/L (72 h)

Reference Shih et al. (2005) Shih et al. (2010b)

Wu et al. (2013)

Dos Santos et al. (2013) Zhang et al. (2014) Aboutaleb et al. (2015)

Kekez et al. (2015) Feng et al. (2015a) Feng et al. (2015b) MoosaviNasab et al. (2010)

Liu et al. (2010)

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including initial pH, temperature, substrate type and concentration, and rotation speed, Shih et al. obtained 49.4 g/L levan from 200 g/L sucrose after submerged fermentation for 21 h. The yield of levan on available fructose was around 50%, and this Takahashi strain was the most efficient levan-producing strain among all of the tested B. subtilis strains in the least time (21 h) under the common cultivation condition (Shih et al. 2005). Later on, the yield of levan was increased by the immobilized B. subtilis (natto) Takahashi cells. After immobilization onto alginate beads, more than 70% of the initial bioactivity was remained after 72 h long reaction cycles, and 70.6 g/L levan was produced from 200 g/L sucrose (Shih et al. 2010b). A tandem and eco-friendly production of levan and ethanol by co-fermentation with B. subtilis (natto) Takahashi and Zymomonas mobilis was also investigated by Shih et al. When cultivated for 48 h, the B. subtilis (natto) Takahashi could produce 56.0  0.6 g/L of levan in a medium containing 250 g/L sucrose, with a 45  0.5% yield on available fructose. The proposed process realized a fully utilization of sucrose substrate without wasting any by-products in the process; furthermore, the generated ethanol could be used to precipitate levan, reducing the recovery cost of levan (Shih et al. 2010a). Recently, with the same strain, Wu et al. studied the production of levan in a 10-L stirred tank bioreactor with batch and fed-batch cultures. As a result, 61 g/L (batch, 24 h) and 100 g/L levan (fed-batch, 120 h) was generated under the optimized condition: pH at 7.0, temperature at 37, agitation speed 175 rpm, and 250 g/L sucrose. In addition to the B. subtilis (natto) Takahashi, the isolated strain B. subtilis (natto) CCT 7712 was also employed to produce 111.6 g/L levan from 400 g/ L sucrose in 16 h. Efforts to produce levan by the other Bacillus species also made some progresses. Zhang et al. reported the levan production by B. methylotrophicus SK 21.002 strain that is isolated from soil samples collecting from beet and sugar cane gardens in China. Finally, 100 g/L levan was produced from 300 g/L sucrose by this strain, suggesting a great potential of B. methylotrophicus SK 21.002 in the industrial preparation of levan, especially for the low molecular weight levan (Zhang et al. 2014). Abou-taleb et al. reported the production of levan by B. lentus V8 strain from two different substrates, including sucrose and sugar cane molasses. Under the optimized conditions, 57.95 g/L or 49.86 g/L levan was obtained from sucrose and sugar cane molasses, respectively. Kekez et al. studied the effect of initial pH, sucrose, and ammonium chloride concentrations on the final production of levan by the B. subtilis NS032 strain. As a result, the maximum levan yield 47.8 g/L was observed under the condition, pH 7.0, sucrose 196.8 g/L, and ammonium chloride 2.4 g/L, while in a higher sucrose system, the maximal production was 99.2 g/L (pH 7.4, sucrose 397.6 g/L, ammonium chloride 4.6 g/L) (Kekez et al. 2015). Feng et al. increased the levan production by tenfold times by deleting the pgs cluster encoding for polyglutamic acid (PGA) synthesis in B. amyloliquefaciens NK-ΔLP, resulting in 22.6 g/L levan from 250 g/L sucrose (Feng et al. 2015a). Later on, the same group further improved the yield of levan to 31.1 g/L from the same sucrose concentration by disrupting six extracellular protease gene and the tasA gene encoding for the biofilm matrix protein (Feng et al. 2015b).

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The Panebacillus genus was another gram-positive bacterium that had been employed to produce levan from sucrose. The production of levan by P. polymyxa was firstly reported by Hestrin et al. Later on, Liu et al. optimized the culture medium including 188.2 g/L of sucrose, 25.8 g/L of yeast extract, 5 g/L K2HPO4, and 0.34 g/ L CaCl2 and produced 35.26 g/L levan under pH 8.0 and 24  C for 60 h (Liu et al. 2010). The Microbacterium laevaniformans PTCC 1406 was another levanproducer that has been studied by Moosavi-Nasab et al. They reported the optimum condition for levan production was at concentration of 20% sucrose for 48 h and 25% of date syrup for 48 h at pH 6.0. Meanwhile, a longer fermentation time would result in some decrease of levan production. Finally, 48.9 g/L and 10.48 g/L levan was produced from sucrose and date syrup under the optimized conditions (Marzieh Moosavi-Nasab et al. 2010).

9.3.2

Biological Production of Levan by Gram-Negative Microorganisms

So far, many gram-negative bacteria had been reported to produce levan, and the results were concluded in Table 9.2. Among these microorganisms, the acetic acid bacterium from Gluconacetobacter diazotrophicus was the most intensively studied. Table 9.2 Production of levan by gram-negative microorganisms Strains A. xylinum NCIM 2526

Substrate (concentration, g/L) Sucrose (60)

Levan production (g/L) 13.25

P. fluorescens NCIM 2059

Sucrose (60)

15.42

H. smyrnensis AAD6T

Sucrose (50) + mannitol (30 g/L) Sucrose (50) + boric acid (50 mM) Beet molasses (30)

6.60

S. levanicum NN

200

G. dizaotrophicus PAl-5

Sucrose (100)

39 (10 h) 50 (7 days) 24.7

Z. mobilis ZAG-12

Sucrose (250)

14.67

Z. mobilis CCT 4494 (mutant) Z. mobilis CCT 4494 (immobilized) Z. mobilis B-14023

Sucrose (300)

42.67

Sucrose (299)

112.53

Sucrose (299)

40.2

Reference Srikanth et al. (2015b) Jathore et al. (2012) Hande Kazak et al. (2015)

8.84 12.4

Küçükaşik et al. (2011) Kikuchi et al. (2010) Idogawa et al. (2014) Calazans et al. (2007) Senthilkumar et al. (2004) Lorenzetti et al. (2015) Silbir et al. (2014)

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The G. diazotrophicus PAl 5 strain was able to produce 24.7 g/L levan from 100 g/L sucrose in batch cultures in the presence of biological N2-fixation (BNF), suggesting a high potential of G. diazotrophicus PAl 5 since it does not require extra N-source supplementation for culture (Boiardi and Molinari 2013). Simultaneously, a high concentration of phosphate was found able to improve the formation of levan (Idogawa et al. 2014). In the case of A. xylinum NCIM 2526 (now classified as Gluconacetobacter swingsii), Srikanth et al. employed a one-factor-at-a-time and factorial design to improve the levan production from 0.54 to 13.25 g/L with 60 g/L sucrose (Srikanth et al. 2015b). Another gram-negative bacterium from Halomonas sp. was also found as a levanproducer microorganism. The strain was further characterized as a novel species of Halomonas sp. and called Halomonas smyrnensis AAD6T (Poli et al. 2009). Lots of optimization studies were carried on H. smyrnensis AAD6T because it is highly halophilic and osmoadaptable and is able to produce levan via an unsterile production under high salinity. For instance, Kazak Sarilmiser et al. reported that the addition of boric acid into the culture medium can effectively enhance the production of levan from 1.84 to 8.84 g/L after 160 h (Hande Kazak et al. 2015). Küçükaşik et al. found the substitute of the sugar beet molasses with sucrose could also be fermented by H. smyrnensis AAD6T, and 12.4 g/L levan was generated from 30 g/L pretreated beet molasses (Küçükaşik et al. 2011). The other gram-negative microorganisms including Zymomonas, Pseudomonas, Serratia sp., and Erwinia species were identified with the levan-producing capacity. Dawes et al. firstly reported the production of levan in Z. mobilis when sucrose was presented. After optimization, 14.67 g/L levan was reported for the ZAG-12 from 250 g/L sucrose (Calazans et al. 2007). After treated with N-methyl N-nitro N-nitrosoguanidine (NTG), the mutant strain of Z. mobilis CCT 4494 showed an increase in the levan production from 30.3 g/L to 42.67 g/L in 24 h (Senthilkumar et al. 2004). Silbir et al. compared the production of levan by Z. mobilis B-14023 in batch and continuous fermentation, and 40.2 g/L levan was generated from 299 g/L sucrose in the batch culture. When fermented in packed bed bioreactor with Ca-alginate immobilized Z. mobilis cells, the highest concentration 31.8 g/L of levan was obtained (Silbir et al. 2014). However, increasing the dilution rate would decrease the formation of levan and increase the residual sugar concentrations. Introducing the immobilized Z. mobilis CCT 4494 cells entrapped in alginate and PVA beads, levan could be produced in the range of 18.84–112.53 g/L when given different pH, sucrose concentrations, and fermentation periods (Lorenzetti et al. 2015).

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Table 9.3 Characteristics of microbial LSases Temperature ( C) 25–45

>50

Subunit (kDa) 52

pH 6.0–6.2

Cation Ni2+

Specific activitya 415 U/mg

NR

5.0

NRb

NR

49

6.0

NR

620 U/mg

Brenneria sp.

49

6.5

NR

570 U/mg

L. sanfranciscensis TMW Z. mobilis

90

5.4

NR

NR

NR

7.0

NR

NR

L. reuteri LTH 5448

76

6.0

500 U/mg

B. subtilis DSM 347 L. reuteri 121

51  2

5.5

Ca2+, Mn2+, Fe2+ Mn2+

NR

4.5–5.5

Ca2+

NR

NR

6.0–6.5

NR

NR

6.0

NR

1274.3 U/ l 714 U/mg

Microorganisms B. amyloliquefaciens B. licheniformis 8-37-0-1 B. goodwinii

G. stearothermophilus R. aquatiltis

336 U/mg

Reference Phengnoi et al. (2020) Lu et al. (2011) Liu et al. (2017) Xu et al. (2018b) Tieking et al. (2005) Han et al. (2009) Ni et al. (2018) Szwengiel et al. (2016) van Hijum et al. (2004) Inthanavong et al. (2013) Kim et al. (1998)

a

Specific activity was calculated as the total activity (including sucrose hydrolysis and transfructosylation) b NR Not reported

9.4 9.4.1

Characteristics of Microbial LSase LSase from Bacillus Species

Bacillus subtilis was firstly found as LSase-producible by Dedonder (Table 9.3). The purified enzyme was about 39 kDa and showed the optimum pH between 5.8 and 6.0 and optimum temperature at 37  C. Also, the presence of Zn2+, Fe2+, and Al3 was found to enhance the thermostability of B. subtilis LSase (Dedonder 1966). Steinmtz et al. reported the expression level of extracellular B. subtilis LSase was regulated by three genes including sacUh, amyB, and pap (Steinmetz et al. 1976). Later on, more LSases were found in the other Bacillus species including the B. megaterium (Korneli et al. 2013), B. amyloliquefaciens (Phengnoi et al. 2020), and B. lichenifomis. Lu et al. optimized the fermentation conditions for levan production by B. lichenifomis 8-37-0-1 LSase and obtained the maximum levan production of 41.7 g/L from 100 g/L sucrose (Lu et al. 2011). By contrast, the B. licheniformis RN-01 LSase generated the levan-type fructooligosaccharide (L-FOS) (more than

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11 DP) as the main product. As a result, the immobilized enzyme could produce L-FOS of 7.35 g from sucrose of 25 g (Sangmanee et al. 2016).

9.4.2

LSase from Gram-Negative Species

LSases could also be produced from the gram-negative bacteria, including the Gluconacetobacter, Zymomonas, Pseudomonas, Erwinia, Brenneria, and Rahnella species. Notably, the LSase from Gluconacetobacter distinguished from the other gram-negative LSase in that the LSase was excreted in a signal peptide-dependent pathway, which was unique among the other gram-negative bacteria (Arrieta et al. 2004). Recently, Liu et al. reported a long-chain levan-producing LSase from the gram-negative B. goodwinii, by which 185 g/L levan was generated from 500 g/L sucrose at pH 6.0 and 35  C after a 12 h reaction. Moreover, the Mw of levan was around 1.3  105 kDa, which was larger than the ones produced by other gramnegative LSases (Liu et al. 2017).

Table 9.4 Molecular weight distribution of microbial levan Microorganisms GramB. natto positive B. subtilis CCT 7712 P. polymyxa NRRL B-18475 P. polymyxa EJS-3 B. methylotrophicus SK 21.002 B. subtilis NRC 33a

Gramnegative

Molecular weight (Da) 1.8  106; 1.2  104 5.7  105; 5  104 2  106 1.22  106; 0.896  106 4–5  103

Liu et al. (2010)

5–6  104

Abdel-Fattah et al. (2005) Ben Ammar et al. (2003) Lu et al. (2011) Hernandez et al. (1995) Brandt et al. (2016) Poli et al. (2009)

Bacillus sp. TH4-2

6  103;6.6  105

B. licheniformis 8-37-0-1 G. dizaotrophicus K. baliensis H. smyrnensis AAD6T

9.6  106 2  106 2.5  109 9.861  106; 1.483  106 0.457  106 4.4  106 1.1–1.6  106

Z. mobilis Serratia sp. E. herbicola ATCC 15552

Reference Shih et al. (2005) Dos Santos et al. (2013) Han and Clarke (1990)

Zhang et al. (2014)

Calazans et al. (2000) Kikuchi et al. (2010) Keith et al. (1991)

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185

Molecular Weight Distribution of Microbial Levan

The molecular weight (Mw) distribution of levan varied largely among different microorganisms (Table 9.4). As reported, the B. subtilis (natto) Takahashi showed two product peaks of 1.8  103 kDa and 11.8 kDa. In the case of B. subtilis CCT 7712, two levan fractions corresponding to the Mw of 5.7  105 and 5  104 Da was also observed (Shih et al. 2005). By comparison, the levan produced by gramnegative bacteria was higher than that of the gram positive due to the presence of outer membrane as well as the periplasm and usually had an average Mw that larger than 106 Da (González-Garcinuño et al. 2017). Notably, the Mw of levan produced by K. baliensis was high to 2.47  109 Da, which was the highest value for all of the bacteria levan (Brandt et al. 2016). However, some other studies also found that the LSase from gram-positive bacteria had a higher transfructosylation performance than that of the gram-negative LSase, which prefer to generate levan polysaccharides rather than the short-chain oligosaccharides, and the differences of enzyme structures were supposed as the main reason for this variation.

Fig. 9.2 Crystal structures of LSase from different microorganisms

Fig. 9.3 Two different elongation mechanisms for levan production

186 W. Xu et al.

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187

Crystal Structures Analysis of LSase

So far, five microbial LSases have been determined by their protein structures by X-ray diffraction (Fig. 9.2). The B. subtilis LSase structure was the first one that is obtained by multiple isomorphous replacement and determined within a 1.5 and 2.1 Å resolution for its apo-form and sucrose-complex, respectively (Meng and Futterer 2003). Overall, from N-terminal to C-terminal stretch, the B. subtilis LSase structure folded like a “funnel” shape with a five-blade β-propeller catalytic domain. Notably, each β-propeller was composed of four antiparallel β-strands adopting a “W” topology (Fig. 9.3a). Moreover, the residues D86, D247, and E342 were validated as the “nucleophile,” “transition stabilizer,” and “general acid” for the B. subtilis LSase. Based on the complexed structures of sucrose, the catalytic mechanism for sucrose hydrolysis was found as a “ping-pong” mechanism. Later on, the structures of LSase from B. megaterium (Homann et al. 2007), gram-negative bacteria G. diazotrophicus (Martínez-Fleites et al. 2005), and E. amylovora (Wuerges et al. 2015) were also determined in the resolution range of 1.75–2.77 Å. In general, there was no big difference in the catalytic domain of LSases from different microorganisms, which are all enclosed by a five-blade βpropeller and employed the putative catalytic residues. In addition to these conserved features, the loops that connect each β-propeller and the β-strand were found variable among gram-positive and gram-negative LSases. For instance, Wuerges et al. compared nine loops of E. amylovora LSase with that of the other LSases, including the residues numbers and shape and speculated the loops residing at the rim of the active site funnel were significant for the product length (Wuerges et al. 2015).

Table 9.5 Mutants of LSase with different product length Enzyme resource B. subtilis

B. megaterium

Z. mobilis

Oligosaccharides R360S, Y429N and R433A only produced oligosaccharides H331L, H331S and H331I only formed the trisaccharide (kestose)

Polymer H243L only generated high molecular weight Levan

N252A formed short-chain oligosaccharides (nystose) The variants N251A and K372A mainly synthesized tri- and tetrasaccharides R369A, R369S, and R369K resulted in elimination of polysaccharide synthesis W47N, W118H and R193K nearly lost the abilities to synthesize Levan

The degree of polymerization (DP) was up to 25

Reference Ortiz-Soto et al. (2008) Chambert and Petit-Glatron (1991) Strube et al. (2011) He et al. (2018)

Li et al. (2011)

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Mutation Studies of LSase in Altering Product Size

As abovementioned, the putative catalytic triad including “nucleophile,” “transition stabilizer,” and “general acid” was conserved for all LSases and responsible for the sucrose hydrolysis. Herein, mutation of these residues would result in a total loss of catalytic activity (Table 9.5). For example, Li et al. reported that the residue of His296 in Z. mobilis LSase did not participate in sucrose hydrolysis, but it acted as a site for acceptor recognizing or binding (Schroeven et al. 2008). In 2011, the same team investigated the conserved 1 site of Z. mobilis LSase and found the mutants W118N, W118H, and R193K almost lost the capacity in producing long-chain levan, indicating the changes in 1 subsites residues could not only influence enzyme activity for sucrose hydrolysis, but also affect the polymerization and product size (Li et al. 2011). In addition, the residue of R360 was also found as an important residue to maintain the polymerization activity of B. subtilis LSase (Meng and Futterer 2003). Correspondingly, the residues of R370 and N252 in B. megaterium LSase were proved as key “switch” in deciding the kind of enzymatic reaction and the final product size. Strube et al. investigated the product spectrum of B. megaterium LSase and found that the mutation of Y247, N252, and K373 also could largely affect the product length and substituted them with other amino acid would cause a remarkable decrease in the formation of polysaccharides. Particularly, the mutant N252A could merely generate short-chain oligosaccharides of up to three fructosyl units (nystose) rather than any other polysaccharide. In the case of variant K373A, the formation of tri- and tetrasaccharides was hampered, but the synthesis of penta- and hexasaccharides was slightly enhanced. In short summary, although these residues did not belong to the sucrose binding subsites (+1 and 1 subsites), they could take part in the acceptor binding and thus impact the synthesis of polysaccharides by LSase (Strube et al. 2011). The different product sizes could be attributed to the different ratio of transfructosylation to hydrolysis (T/H) in LSase. Ortiz-Soto et al. investigated the S173 and S422 as well as the Y421 and Y439 and found they played important roles in determining the T/H ratio B. megaterium LSase. These residues resided in the first shell of the catalytic pocket, and mutants S173A/S422S, S173A/S422A, and S173A/ S422T displayed a remarkable increase in the T/H ratio compared to that of the wildtype LSase (Ortiz-Soto et al. 2017). Recently, Xu et al. reported the A154S mutant of LSase from Brenneria sp. EniD312 could shift the T/H ratio from 2.70 of the wildtype enzyme to 1.04 of the mutant, wherein the total activity was almost retained (Xu et al. 2018b).

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Two Types of Elongation Mechanism for Levan

Early in 1956, Avigad et al. studied the donor-acceptor specificity of LSase from A. levanicum and reported a two-step procedure for the production of levan from sucrose. Two steps mainly involved a double replacement of different fuctosyl acceptors. When water molecules was accepted, then the hydrolysis reaction happened; otherwise the transfructosylation happed when the other fuctosyl acceptor was provided (Avigad et al. 1956). Later on, Paulis et al. specified the process as splitting of the α1-β2 glycosidic bond in sucrose, formation of an enzyme-fructosyl intermediate, and transferring of fructosyl units to the non-reducing glucose end of sucrose with the formation of β -(2, 6) fructan. As we know, the elongation of fructosyl to polymer required that the generated new fructan molecules must be able to utilize as new fructosyl acceptor after the first step of sucrose cleavage (Paulis and Barry 1969). Chambert and Denoder proved that the fructose unit was added one by one for each time onto an acceptor molecule through the kinetic and chemical investigations of B. subtilis LSase (Chambert et al. 1974). However, based on the solved crystal structure, the residues in the vicinity of sucrose known as 1 and +1 subsites were primarily responsible for the sucrose hydrolysis, but not for the binding of other fructosyl acceptors. Moreover, these residues were relatively conserved for all of the gram-positive and gram-negative LSase, which could not fully explain the difference of product spectrum. In the study of LSase and ISase from L. reuteri 121, Ozimek et al. proposed a processive and non-processive (semi-processive) mechanism for the synthesis of fructan for the first time. They compared the different product sizes produced from LSase (mainly levan polymer) and ISase (mainly inulin-type fructooligosaccharides up to DP 15) and supposed there should be two different product formation mechanisms for levan and inulin. They found the affinity of substrate-binding sites in ISase was higher for the shorter-chain FOSs than that of longer FOSs; but on the contrary, the affinity was higher for larger polymer for LSase. Herein, the ISase was proposed to adopt a non-processive mechanism during the elongation of inulin-type fructan, while the LSase adopts a processive route for levan production (Ozimek et al. 2006). During the investigation of B. subtilis LSase, Raga-Carbajal et al. also employed two elongation mechanisms for the formation of levan from sucrose (Raga-Carbajal et al. 2016). They studied both the effect of enzyme concentration and substrate concentration on the elongation mechanism. As a result, they concluded that the generation of low molecular weight (LMW) levan was non-processive and accompanied by a plenty of intermediate size levan-type FOSs with DP up to 70. On the other hand, the high molecular weight (HMW) levan was synthesized through a processive mechanism, and only few oligosaccharides with DP 90%) DP 3-65 DPav 23-42

Tuber

36.45

DPav 19-23

Dendelion (Taraxacum officinale) Garlic (Allium sativum)

Root

26

DP 2-40

Tuber

12.5–23.5

Jerusalem artichoke (Helianthus tuberosus L.) Onion (Allium cepa) Salisfy (Tragopogon porrifolius L.) Sweet leaf (Stevia rebaudiana) Yacon (Smallanthus sonchifolius)

Tuber

14–19

Tuber –

1.1–7.1 3.7

Root

29

DPav 16 and DPav 21 DP 2-80/ 2521 DP 1-12 DP 2-7 (88.6%) DPav 28

Zhu et al. (2019) and Maumela et al. (2020) Loo et al. (2014) Beiro-Da-Costa et al. (2009) Lopes et al. (2015)

Root

35

NR

Castro et al. (2013)

Fig. 10.2 The production process of inulin from plants

References Apolinário et al. (2017) Singh and Bhermi (2008) Shalini and Antony (2015) Nemeth et al. (2015) Cao et al. (2018) Glyn et al. (2004) and Hossein et al. (2018) Petkova and Denev (2018) Schütz et al. (2010) Zhang et al. (2013)

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2011), microwave (Petkova et al. 2018), and pulsed electric field processing (Zhu et al. 2012) are used during this procedure; (3) purification, the inulin juice with high purity is purified further; purification procedure such as ultrafiltration, specific crystallization, precipitation, centrifugation, and decantation may be selected according to the DP of inulin (Gupta et al. 2019); and (4) drying, spray-drying is the most common way to obtain the final product inulin. In contrast, microbial inulin has not been produced on an industrial scale. However, some recent studies focusing on the properties and functions of microbial inulin revealed some potential applications, which may arouse attention about the commercial production of microbial inulin.

10.3

Inulin-Forming Enzymes

10.3.1 Inulin-Forming Enzymes from Plants Two enzymes, sucrose:sucrose 1-fructosyltransferase (1-SST EC: 2.4.1.99) and fructan:fructan 1-fructosyltransferase (1-FFT EC: 2.4.1.100), participate in the synthesis of inulin in plants. They both belong to the Glycoside Hydrolase 32 family (GH32) and play different roles during the production of inulin. 1-SST catalyzes the transfructosylation reaction of two molecules of sucrose, transferring the fructosyl moiety of sucrose to another sucrose to generate 1-kestose (α-D-glucopyranosyl(1-2)-β-D-fructofuranosyl-(1-2)-β-D-fructofuranose), releasing glucose at the same time. Then the formed 1-kestose is used as the acceptor of the fructosyl to 1-SST to generate 1-nystose (α-D-glucopyranosyl-(1-2)-β-D-fructofuranosyl-(1–2)-β-Dfructofuranosyl-(1-2)-β-D-fructofuranose). By that analogy, the extended saccharide chains can also be used as acceptors. Thus, inulin is synthesized.

10.3.2 Inulin-Forming Enzyme from Microorganisms Compared to vegetal inulin, the synthesized process of microbial inulin is more straightforward. One enzyme in some microorganisms, ISase, uses sucrose as the sole substrate to produce inulin by a series of continuous reaction. Firstly, sucrose is cleaved into a molecule of glucose and enzyme-frucosyl complex intermediate. Secondly, if a water molecule is the acceptor of frucosyl, fructose will be released, hydrolysis reaction occurring. Similarly, if the frucosyl is transferred into another sucrose molecule, 1-kestose will be formed, transfructosylation reaction occurring (Fig. 10.3). Then, the produced fructooligosaccharides (FOSs) are utilized as fructosyl acceptors to generate long-chain inulin. Based on the particular reaction process, total activity, hydrolysis activity, and transfructosylation activity are defined, which are represented by the amount of glucose and fructose and the subtraction of glucose to fructose, respectively.

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H2O a-(1, 2)

Fru

Fru

Hydrolysis

Enzyme

Glu

+ Fru

a-(1, 2)

Glu

Fru Glu Glu

Sucrose

b-(1, 2)

E-Fru intermedite

Transfrutosylation Fru a-(1, 2)

Fru

Glu

Glu

1-Kestose

a-(1, 2)

b-(1, 2)

a-(1, 2)

Fru

Glu

Fru

Fru

n

Glu

Inulin

Fig. 10.3 The catalytic process of inulosucrase

10.4

Characterization of ISase

10.4.1 Microbial Sources of ISase Since the first fungus, A. sydowi IAM 2544, was found to generate inulin, more than 15 microorganisms have been determined as inulin-producing individuals. The enzymatic properties of nine ISases in the native or recombinant forms have been identified in detail. The DP and molecular weight of mature inulin were also determined to be almost in the range of 106–107 Da. In this part, the enzymatic properties and factors affecting ISase activity or inulin production were discussed comprehensively.

10.4.2 Sequence Analysis of ISase Lactobacillus-derived ISases are the most widely studied ones due to the safety of Lactobacillus. The whole sequence of ISase from Lactobacillus consists of four regions: (I) (II) (III) (IV)

Signal peptide, it possesses about 40 amino acids. N-terminal variable region. Catalytic domain. C-terminal variable region, it includes a cell-wall-anchoring sequence.

On the one hand, the signal peptide guides the secretion of ISase, but the cell wallanchoring sequence at the C-terminal region prevents the secretion. On the other hand, the N-terminal C-terminal variable regions are unnecessary to structural integrity. Therefore, the N-terminal C-terminal truncation strategies are always used in the enzymatic properties, inulin synthesis, and structural analysis. Similarly, LSases from Lactobacillus species also show the sequence trait. Figure 10.4 shows the truncated ways of ISases and LSases from some Lactobacillus species, including ISases from L. johosonii NCC533 (Pijning et al. 2011), L. gasseri DSM 20604 (Anwar et al. 2010), L. gasseri DSM 20243 (Anwar et al. 2010) and L. reuteri

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Fig. 10.4 Schematic representation of ISases and LSases from Lactobacillus species showing four organized domains. (I) Signal peptide, (II) N-terminal variable region, (III) catalytic domain, (IV) C-terminal variable region. Δ represents the remained amino acids for the construction of the recombinant enzymes

121 (Van Hijum et al. 2002), LSases from L. reuteri 121 (Van Hijum et al. 2004), L. reuteri LTH5448 (Ni et al. 2018), and L. reuteri 20,077 (Anwar et al. 2010).

10.4.3 The Effects of pH and Temperature The primary enzymatic properties of reported ISases were summarized in Table 10.2. pH and temperature are two essential elements affecting the activity of enzymes. Only in optimal pH and temperature, enzymes can show their roles adequately. However, few enzymatic properties’ information of ISases from A. Sydowi, Lactobacillus reuteri TMW1.106, Streptococcus sp., and Weissella confusa MBFCNC-2(1) are available. Only the molecular weight of produce inulin was measured. According to the recombinant ISases from Lactobacillus gasseri DSM 20243 (Anwar et al. 2010), Lactobacillus gasseri DSM 20604 (Ni et al. 2017), Lactobacillus johnsonii NCC 533 (Anwar et al. 2008), Lactobacillus reuteri 121 (Van Hijum et al. 2002), Streptococcus mutans GS-5 (Heyer et al. 1998), and S. viridochromogenes DSM40736 (Frasch et al. 2017) and native ISase from Leuconostoc citreum CW28 (Olivaresillana et al. 2002), an apparent optimal pH preference could be summarized that ISase tends to show maximum activity at slightly acidic and neutral environments, except ones from Bacillus sp. including

ND AAO25086.1

ND

ACZ67286.1

2YFR_A

OJI11288.1

CAL25302.1 ND AAA88584.1

L. gasseri DSM 20243

L. gasseri DSM 20604

L. johnsonii NCC 533

L. reuteri 121

L. reuteri TMW1.106 S. mutans BHT S. mutans GS-5

GenBank CAB89083.1 ATN45518.1

Microorganisms A. sydowi IAM 2544 B. agaradhaerens WDG185 Bacillus sp. 217C-11 L. citreum CW28

Recombinant (E. coli) Recombinant (E. coli) Recombinant (E. coli) Recombinant (E. coli) Native Native Recombinant (E. coli)

Native or recombinant Native Recombinant (B. subtilis) Native Native Recombinant (E. coli)

ND/ND ND/ND 5.4/30

5–5.5/50

7.0/55

5.5/35

4.5–5.5/50

7–8/45–50 6.5/45

Optimized pH/Temperature ( C) ND/ ND 6.0–10.0/ 60

ND ND ND

ND

50  C, 180 min, 84% ND

ND

ND 35  C, 420 min, 50% 50  C, 30 min, 0

Thermostability (T1, T2, L) ND ND

Table 10.2 Comparison of enzymatic properties of ISases form different microorganisms

ND ND ND

Effect

Effect

Effect

Effect

Ca2+ ND No effect ND Effect

Schwab and Gänzle (2006) Ebisu et al. (1975) Heyer et al. (1998)

Van Hijum et al. (2002)

ND/>107 ND/ND 8/ND ND/7  107

Anwar et al. (2008)

Ni et al. (2017)

Wada et al. (2003) Olivaresillana et al. (2002), Ortiz-Soto et al. (2004), del Moral et al. (2008) and Olivares-Illana et al. (2003) Anwar et al. (2010)

References Kawai et al. (1973) Kralj et al. (2018)

ND/5. 858  106 ND/4  107

ND/ND

10–18/ND ND/ 2.6–3.4  103 1.35–1.6 106

DP/Mw (Da) ND/2  107 3–25/3  103

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ADB27748.1

ND ND EFL36273.1

Native Native Recombinant (E. coli) Recombinant (E. coli and B. subtilis) ND/ ND

ND/ND ND/ ND 5.0–7.0/50–55 ND

ND ND ND

ND not described, T1, T2, L incubation temperature, incubation time, relative activity

S. mutans JC-1 S. mutans JC-2 S. viridochromogenes DSM40736 W. confusa MBFCNC-2(1)

ND ND No effect ND ND/ND

27/ND ND/2  107 ND/2.5  107 Malik (2012) and Malik et al. (2015)

Ebisu et al. (1975) Rosell and KG (1974) Frasch et al. (2017)

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Bacillus agaradhaerens WDG185 (Kralj et al. 2018) and Bacillus sp. 217C-11 (Wada et al. 2003), which showed optimal pH at weak alkaline conditions. The optimal temperatures of half of the reported ISases fall in the range of 40–45  C. The recombinant ISases from B. agaradhaerens WDG185 (Kralj et al. 2018) and S. mutans GS-5 (Heyer et al. 1998) showed the optimal temperatures at 60  C and 30  C, respectively, which are two extremes. Frequently, transfructosylation activity and hydrolysis activity showed the maximum at different temperatures. For instance, ISase from L. gasseri DSM 20604 exhibited optimal transfructosylation activity and hydrolysis activity 25  C and 45  C, respectively (Ni et al. 2017). The phenomenon that low temperatures facilitate transfructosylation and high temperatures aggrandize hydrolysis also occurs in some LSases, such as LSases from Lactobacillus reuteri LTH5448 (Ni et al. 2018) and Leuconostoc mesenteroides B-512 FMC (Li et al. 2015b). Nevertheless, L. johnsonii NCC 533 ISase was an opposite example, and it preferred hydrolysis at 40  C and transfructosylation at 55  C (Anwar et al. 2008).

10.4.4 Thermostability of ISase Thermostability is one of the most crucial factors deciding the application of enzymes in industry. However, the information about the thermostability of ISase is limited, and only two ISases are measured the thermostability. The half-life of L. citreum CW28 ISase is 420 min at 35  C, but it is extremely unstable when the temperature over 50  C (Moral et al. 2008). L. gasseri DSM 20604 ISase shows good stability at temperatures below 50  C, but the activity decreases rapidly once the temperatures are more than 55  C (Ni et al. 2017). Summarily, the thermostability of ISase is not so good enough to satisfy the need for industrial applications. Therefore, exploring novel ISases with high thermostability or improving the thermostability of existing ISases by molecular modification is desirable.

10.4.5 The Effect of Sucrose Concentration and Enzyme Dosage The typical feature of the effect of sucrose concentration on the activity of ISase is that a high level of sucrose facilitates the transfructosylation activity, and ISase prefers hydrolyzing sucrose at low sucrose concentration (Ni et al. 2017; Kralj et al. 2018; Peña-Cardeña et al. 2015). For instance, the transfructosylation activity of L. reuteri 121 ISase occupied for 90% of the total activity when the sucrose concentration was 1.7 M (Ozimek et al. 2006). The effect of sucrose concentration on the activity of LSase exhibited consistent traits with that of ISase, and the molecular weight of the synthesized levan could be controlled by sucrose

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concentration (González-Garcinuño et al. 2017). The phenomenon may result from a high sucrose concentration that repels water molecules from the catalytic domain, decreasing the possibility of water molecules as the acceptors of fructosyls. Thus hydrolysis reaction is inhibited (Chuankhayan et al. 2010). The effect of enzyme dosage on the activity of ISase is opposite to that of sucrose concentration on its activity. Peña-Cardeña et al. found that the hydrolysis activity increased and the products synthesized were nearly oligosaccharides rather than polysaccharides when the enzyme dosage increased (Peña-Cardeña et al. 2015). Additionally, Ni et al. (2017) synthesized inulin using L. gasseri ISase, and the inulin output decreased once the enzyme dosage excessed 4.5 U/mg sucrose, which indicated that high enzyme dosage was adverse to inulin biosynthesis. Therefore, it is essential to balance the amount of sucrose concentration and enzyme dosage in the process of inulin synthesis.

10.4.6 Kinetic Parameters Based on some studies probing the kinetic parameters of ISase, the transfructosylation and total activities cannot be saturated by sucrose (Anwar et al. 2008, 2010; Van Hijum et al. 2003), which would lead to high standard errors with curve fits. Therefore, the Hill equation is more appropriate to analyze the kinetic parameters of ISases instead of Michaelis-Menten kinetics. According to Anwar et al. (2008), the non-Michaelian behavior of ISase may result from FOSs synthesized at the early stage of the reaction which are more available as acceptors of fructosyls than the growing polysaccharide chains. The non-Michaelian phenomenon also occurs in some LSases such as these from L. reuteri 121 (Van Hijum et al. 2004), Bacillus amyloliquefaciens (Tian and Karboune 2012), and Lactobacillus sanfranciscensis TMW 1.392 (Tieking et al. 2005). The sources and truncated ways of ISases may affect the kinetic behavior. ISase from L. reuteri 121 is the most representative one that follows the Hill equation (Van Hijum et al. 2003). However, some exceptions are confirming the Michaelian behavior. Moral et al. (2008) constructed four truncated ISases (IslA- IslA4) by removing different fragments from the whole sequence, and the results showed that only IslA3 truncating C-terminal region was best described by Hill equation, while IslA, IslA2, and IslA4 exhibited Michaelis-Menten-type kinetics. Recently, a novel ISase from B. agaradhaerens WDG185 was characterized, and the Michaelis kinetics was observed. The Km values of transfructosylation activity and total activity for sucrose were 1.4  0.2 and 2.2  0.2, respectively (Kralj et al. 2018). However, the relative structural feature and the molecular mechanism controlling the kinetic behavior remained unclear.

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Structure and Molecular Modification

10.5.1 Overall Structure Currently, only one ISase from L. johnsonii NCC533 (Lajo-ISase) has been resolved the crystal structure. The recombinant Lajo-ISase was constructed by N- and C-terminal truncated ways, remaining the residues from 145 to 708. The apo structure and the complex structures with sucrose and 1-kestose were obtained. The overall structure presents the five-bladed β-propeller fold (Fig. 10.5), which is a representative bimodular arrangement of CH-J enzymes. Generally, each blade consists of four antiparallel β-strands, and all β-strands built the funneled pocket. The residues of these β-strands are highly conserved among ISases, especially the ones from Lactobacillus species. The sequence difference is mainly embodied in the loops connecting the β-strands. Up to now, the crystal structures of five LSases have been resolved. Compared to the LSase structures, an apparent difference can be found that the N-terminal of ISase is longer. But the specific roles of the ISase long N-terminal have remained unclear.

10.5.2 Subsites in the Catalytic Pocket ISase also adopts the classic double-displacement reaction mechanism to catalyze the hydrolysis and transfructosylation, and the putative processive and non-processive mechanisms are referred to depict the saccharide-chain extension, which is the same as LSase (Chap. 9). The catalytic residues of ISases and LSases are totally conserved, composing of two aspartic acids and a glutamic acid. In

Fig. 10.5 The overall structure of Lajo-ISase. Left, the apo structure of Lajo-ISase (PDB: 2YFR). Right, the complex structure with sucrose (PDB: 2YFS). The five-bladed β-propeller fold (I-V) is labeled by ruby, lime, slate, yellow, and cyan, respectively. N-terminal and C-terminal are colored by orange and salmon. The green spheres represent the Ca2+. The sucrose at the catalytic core is marked as red. The second sucrose having little interaction with the enzyme is tabbed as sky blue

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Fig. 10.6 Saccharides binding sites of Lajo-ISase. (a) Two sucrose binding sites. The red sucrose molecule is located at the catalytic core domain, and the blue one binds at the surface. (b) Sucrose binding site at the catalytic core domain. (c) 1-Kestose binding site at the catalytic core domain

Lajo-ISase, the catalytic triads, Asp272, Asp425, and Glu524, function as the nucleophile, transition-state stabilizer, and general acid/base, respectively. By socking the inactive variation D272N crystals in a high concentration of sucrose, the complex structure with sucrose was obtained. As is shown in Fig. 10.6a, two sucrose molecules are bound to the structure, one of them is merged in the catalytic core, and the other one is attached at the surface. The second sucrose has little interaction with the enzyme; only the fructosyl moiety makes hydrogen bonds with Asn301, Asn 305, and Arg623. The first sucrose merging in the catalytic domain forms numerous direct hydrogen bonds with protein side chains of residues (Fig. 10.6b). The fructosyl moiety lies in the 1 subsite, forming hydrogen bonds with the Trp271, Asn272, Ser340, Arg424, Asp425, and Glu524, which are conserved in GH68 enzymes. Due to the strict conservation of 1 subsite, only donor fructosyl moiety can bind here. In contrast, the conservation of +1 and +2 subsites are weaker than 1 subsite, both fructosyl and glucosyl can attach here. When sucrose binds at the catalytic core, Glu522 and Arg 542 form hydrogen bonds with the glucosyl moiety at +1 subsite. Arg 542 shows alternative conformation, so when 1-kestose binds at the catalytic core, Arg 542 and Arg 545 makes hydrogen bonds with the glucosyl at +2 subsite (Fig. 10.5c).

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10.5.3 Ca2+ Binding Sites The complex structure with sucrose reveals two Ca2+ binding sites at the periphery of the five-bladed β-propeller fold. However, the apo structure and the complex structure with 1-kestose only uncover the first Ca2+ binding site (Fig. 10.7a), which is highly conserved in ISases. Five residues, Asn419, Gln450, Trp487, Asn489, and Asp521 can make interactions with the Ca2+, which may stabilize the surrounding microenvironment. The second Ca2+ binding site is farther from the catalytic core than the first one, but there are still four residues making interactions with it (Fig. 10.7b). Ca2+ can improve the activity of some ISases. For example, in the absence of Ca2+, the L. reuteri 121 ISase activity decreased to 79% (Van Hijum et al. 2002). Generally, the activity of ISase from Lactobacillus can be promoted by Ca2+ (Table 10.2), except ones from B. agaradhaerens WDG185 and S. viridochromogenes DSM40736 due to the missing of Ca2+ binding site (Frasch et al. 2017; Kralj et al. 2018).

10.6

Modulation of Product Chain Length

10.6.1 The Chain-Length Distribution of Microbial Inulin The molecular weight of microbial inulin synthesized enzymatically from sucrose varies from each other. Generally, the molecular weight of microbial inulin is hundreds of times more massive than vegetal inulin. Some microbial inulin has been measured the molecular weight which distributes in 106–107 Da, including these from L. johnsonii NCC 533 (4  107 Da) (Anwar et al. 2008), S. viridochromogenes DSM40736 (2.5  107 Da) (Frasch et al. 2017), S. mutans JC-2 (2  107 Da) (Rosell and KG 1974), S. mutans GS-5 (7  107 Da) (AduseOpuko et al. 1989), L. reuteri 121 (>107 Da) (Van Hijum et al. 2002), and L. gasseri DSM 20604 (5.9  106 Da) (Ni et al. 2017). Different from the abovementioned

Fig. 10.7 The Ca2+ binding sites of Laje-ISase. The green spheres represent Ca2+, and the dummy lines indicate as the interaction with residues

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ISases synthesizing single molecular weight inulin, L. citreum CW28 ISase could produce high-molecular-weight inulin (1.35  106–1.6  106 Da) and lowmolecular-weight inulin (2.6  103–3.4  103 Da) (Ortiz-Soto et al. 2004). ISase from B. agaradhaerens WDG185 is the sole one that only produces low-molecularweight inulin with the molecular weight of 3  103 Da (Kralj et al. 2018). Additionally, the sole fungus-derived ISase, A. sydowi IAM 2544 ISase, could produce inulin with the molecular weight of 7  107 Da (Kawai et al. 1973), which is the highest one compared to other microbial inulin.

10.6.2 The Chain-Length Modulation of Microbial Inulin The molecular weight of polysaccharides is one of the most significant factors impacting their properties, functions, and applications. The molecular weight of levan produced by LSase could be regulated by sucrose concentration, enzyme dosage, ionic strength, organic solvents, and protein immobilization (Ortiz-Soto et al. 2019; Raga-Carbajal et al. 2016). Producing tailor-made inulin by controlling reaction conditions or modifying ISase is an important research direction. Therein, molecular modification is the most frequently used way. Ozimek et al. (2006) firstly reported the enzymatic inulin chain modulation by site-director mutagenesis. A single mutation was carried out on the three completely conserved residues at the 1 site of L. reuteri 121 ISase, and three variations, W271N, W340N, and R423K, with changed product chain length were obtained. Compared to the wide-type ISase, W271N and R423K synthesized more FOSs with the DP exceeding 10, and W340N could only produce kestose (GF2) and nystose (GF3). Anwar et al. (2012) constructed the variation N543S, which lost the capability to synthesize long-chain inulin and only produced FOSs with the DP below 6. By altering the residues around the active site, Rodríguez-Alegría et al. (Rodríguez-Alegría et al. 2010) obtained three L. citreum ISase variations that changed the chain length, S425A, L499F, and R618K. Therein, S425A and R618K could only generate oligosaccharides, indicating the importance of the two residues in controlling inulin extension. L499F remained the capability to synthesize inulin, and many oligosaccharides were also observed in the reaction mixture. The reports involving the chain-length modulation are mainly focused on the ISase from L. reuteri 121. Two main procedures are often used in relevant researches: (1) eliminating or weakening the interaction between sucrose and enzyme at the binding site, which can be realized by alanine replacement, and (2) blocking the track for sucrose binding and saccharide chain extension, which can be realized by aromatic amino acids’ replacement. Charoenwongpaiboon et al. (2019c) analyzed the residues interacting with the carbohydrate molecules in the binding site from the crystal structures of 84 carbohydrate-enzyme complexes and found that 80% of the residues were charged and polar amino acids. Accordingly, they selected some charged and polar amino acids around the binding site as the targeted residues and two potential chain extension tracks

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were speculated, track A (N543, W551, R483, D479, S482 and D478) and track B (N543, N561, N555 and D689). These residues were replaced with alanine to eliminate the potential hydrogen bond and polar interaction, which lead to the declining the capability to synthesize inulin, while the production of FOSs with the DP below 10 (GF9) was promoted. Taking R483A as an example, it generated more FOSs with the DP below 6 (GF5). Moreover, Charoenwongpaiboon et al. (2019a) studied the second chain-length modulating way by computer-aided rational protein mutagenesis. Binding GF4 in the catalytic site is the indispensable precondition to form GF5. Therefore, docking GF4 with the variations R483A, R483F, R483Y, and R483W and counting the binding free energies, respectively, they found that all the binding free energies of the variations decreased except R483W. Furthermore, they analyzed the chainlength distribution of the products of the four variations and found that they produced more FOSs with the DP ranging from 4 to 8 than the wild type and the highest DP of the products was lower than 12 (GF11). For modulating the inulin chain length of ISase, the crux is to solve the deeper sites of produced FOSs. Currently, the complex structure of ISase with kestose (GF2) has been solved, revealing the occurrence of +2 subsite and the relative residues (Pijning et al. 2011). Hence, resolving the complex structures of ISase with higher DP of FOSs is crucial to disclose the deeper subsites and corresponding residues, which is meaningful to the chain-length modulation.

10.7

Applications of ISase

10.7.1 Production of High-Molecular-Weight Inulin The molecular weight of polysaccharide is one of the most significant factors affecting its properties and functions. Some studies about levan-type fructan determined that high-molecular-weight levan showed more superior properties in some aspects. For example, the same amount of levan with different molecular weight was supplemented into baking bread, and the result exhibited that the bread added highmolecular-weight levan showed softer texture and larger volume than that added low-molecular-weight levan (Tharalinee et al. 2017). The apparent difference between microbial inulin and vegetal inulin is their molecular weight. Generally, the molecular weight of microbial inulin is hundreds of times larger than vegetal inulin. A few studies probing the properties of microbial inulin uncovered the better solubility, gel-forming properties, and storage stability than vegetal inulin (Ni et al. 2020b; Wada et al. 2005). Therefore, the potential applications of microbial inulin deserve further exploration. Currently, inulin produced and applied commercially is from plants. Although vegetal inulin is widespread, the extraction process is complex, and some impurities like other polysaccharides inevitably exist in the final product. Microbial inulin is synthesized by inulosucrase with sucrose as the sole substrate, and the reaction

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system only contains fructose, glucose, sucrose, and inulin. Therefore, the process of purification is more straightforward, and the purity of produced inulin is higher than vegetal inulin. The study focusing on the production of microbial inulin by inulosucrase from L. gasseri DSM 20604 showed that the maximum yield reached 53 g/L at optimized conditions (Ni et al. 2017). Additionally, the by-product of the sugar industry like molasses may also be used as the substrate to produce inulin due to the high content of sucrose, which will reduce the cost and be conducive to the comprehensive utilization of resources.

10.7.2 Production of FOSs FOS is a type of significant prebiotics possessing many essential physiological effects, such as the roles of intestinal microbiota. It stimulates the growth and activity of beneficial colonic bacteria like Bifidobacteria and inhibits the harmful groups, thus reducing the incidence of gastrointestinal infections and keeping the intestine healthy (Flores-Maltos et al. 2016). FOSs are formed at the early stage of producing inulin. Hence, plants reserving inulin as energy storage can also accumulate FOSs. Traditionally, FOSs are extracted from inulin-rich plant materials. At present, using biotechnological strategies to produce FOSs has been commercialized. Two enzymes, endo-inulinase (EC 3.2.1.7) and β-fructofuranosidase (EC 3.2.1.26), utilizing inulin and sucrose to synthesize FOSs, respectively, have been widely studied. The chain length of produced inulin could be controlled by changing some residues, making inulosucrase lose the capability to synthesize long-chain inulin and efficiently obtain tailor-made FOSs. For instance, the variation R483A of inulosucrase from L. reuteri 121 could synthesize FOSs with DP no more than 10 (Charoenwongpaiboon et al. 2019c). Then, the variation was used to prepare the cross-linked enzyme aggregates to produce FOSs, and the production of FOSs reached 12 g/L after 4 h at optimized conditions with 50 g/L sucrose as the substrate (Charoenwongpaiboon et al. 2019b). Additionally, N- and C-terminal sequence truncation and effective secretion signal peptide insertion strategies were applied to construct the recombinant ISase from L. reuteri 121, and it was expressed in Saccharomyces cerevisiae. 128.4 g/L medium-chain FOSs with the DP 2-20 were obtained from 300 g/L sucrose by fed-batch fermentation, and the conversion yield of sucrose reached 85.6% (Ko et al. 2019). Moreover, the production of the smallest FOS, 1-kestose, reached 22.9 g/L (Ko et al. 2019). Some latest studies disclosed that short-chain FOSs, 1-kestose in particular, showed more efficient roles in improving the intestinal environment and activating the gut immune system than long-chain FOSs (Kim et al. 2018; Shimomura et al. 2017). Charoenwongpaiboon et al. (2018) improved the pH stability and thermostability by immobilizing inulosucrase on the core-shell chitosan beads, and a continuous fixed-bed bioreactor was created to produce FOSs with 1-kestose as the main composition.

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10.7.3 Production of Novel Oligosaccharides Using water molecules and sucrose as the acceptors of fructosyls, ISase causes hydrolysis reaction and transfructosylation reaction, respectively, which generates glucose and inulin, respectively (Fig. 10.2). Transglycosylation capability is a crucial factor in evaluating the value and application of glycosyltransferase. Some scarce and unnatural saccharides or derivates could be synthesized by transglycosylation reaction, which is a research hotspot of enzymes owning the transglycosylation capability, such as the sucrose-type enzyme including glucansucrase (Te Poele et al. 2016), amylosucrase (Daude et al. 2013), and LSase (Ruiz-Aceituno et al. 2017). Some studies showed that not only could water molecules and sucrose be acceptors but some other saccharides may also be acceptors of fructosyl to generate novel oligosaccharides. The transfructosylation capability of ISase was firstly probed by RodríguezAlegría et al. (2010). Sucrose was used as donner, and xylose and maltose were used as acceptors, respectively; the results showed that besides FOSs and inulin, some other oligosaccharides also occurred in the reaction mixture, which was speculated as novel transfructosylation products. However, the specific structures of these novel products were not identified. ISase from L. gasseri DSM 20604 is the most common one used to study the synthesis of novel oligosaccharides by acceptor reaction. Díez-Municio et al. (2016) explored the enzymatic production of novel oligosaccharides by ISase from L. gasseri DSM 20604 using sucrose and raffinose as co-substrates and characterized the structures of produced oligosaccharides. The structural information determined that the transferred fructosyls were connected to raffinose by β-(2, 1) glycoside bonds and raffinosyl-oligofructosides with DP 4-8 (αD-galactopyranosyl-(1,6)-α-D-glucopyranosyl-(1,2)-β-D-fructofuranosyl-((1,2)-βD-fructofuranoside)n) were formed. Additionally, sucrose and maltose were used as co-substrates to produce maltosyl-fructosides with DP 3-6, and α-D-glucopyranosyl(1,4)-α-D-glucopyranosyl-(1,2)-β-D-fructofuranoside was the main product (Diezmunicio et al. 2013). ISase can transfer fructosyl and form new β-(2, 1) linkages with the acceptors. However, studies about acceptor reaction are not as many as other sucrose-type enzymes, which may ascribe to the limited sources of ISase. Many saccharides, aliphatic alcohols, and aromatic alcohols could be used as acceptors of fructosyls by LSase, and the produced oligosaccharides and derivates have promising applications in the food industry (Mena-Arizmendi et al. 2011; Li et al. 2015a). Therefore, the characterization of novel ISase with high transfructosylation capability and screen the acceptor spectra of fructosyl are essential for the application of ISase.

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10.7.4 Production of Inulin Nanoparticles Polysaccharide-based nanoparticles have attracted much attention due to their versatile performance and showed promising biomedical applications (Seidi et al. 2018; Swierczewska et al. 2016). Vegetal inulin nanoparticles showed potential as a hepatoprotective and encapsulated media (Abdelwahhab et al. 2018; Cui et al. 2018). Recently, the production and application of high-molecular-weight inulin nanoparticles have been studied. For example, Jiménez-Sánchez et al. (2019) produced inulin using ISase from L. citreum, and stable spherical nanoparticles with an average diameter of 112 nm were formed. The inulin nanoparticles are not harmful to the peripheral blood mononuclear cells at concentrations below 200 μg/mL. Temperature is one of the most crucial factors impacting the formation and diameter of inulin nanoparticles. Charoenwongpaiboon et al. (2019d) optimized the production conditions of inulin nanoparticles, including temperature, sucrose concentration, and enzyme dosage, obtaining a high yield of inulin nanoparticles. Then, the inulin nanoparticles were complexed with quercetin and fisetin, and the results showed that microbial inulin nanoparticles could promote the solubility and antioxidation of these flavonoids. Therefore, microbial inulin nanoparticles could be used as biocompatible materials to improve the solubility and prevent the degradation of natural functional materials.

10.8

Conclusions

Inulin, a versatile polysaccharide, has been applied widely in the food industry. Recently, high-molecular-weight microbial inulin has shown many more superior properties. ISase, as an efficient transfructosylation tool, could be used to produce microbial inulin. In this chapter, the outstanding properties and applications of inulin have been summarized briefly. More attention has been paid on the ISase, including the enzymatic properties, crystal structure, and chain-length modulation. Moreover, the promising applications of ISase in the production of microbial inulin, FOSs, novel oligosaccharides and inulin nanoparticles light the perspective of ISase.

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Li W, Yu S, Zhang T, Jiang B, Mu W (2015a) Recent novel applications of levansucrases. Appl Microbiol Biotechnol 99(17):6959–6969 Li W, Yu S, Zhang T, Jiang B, Stressler T, Fischer L, Mu W (2015b) Efficient biosynthesis of Lactosucrose from sucrose and lactose by the purified recombinant Levansucrase from Leuconostoc mesenteroides B-512 FMC. J Agric Food Chem 63(44):9755–9763 Loo JV, Coussement P, Leenheer LD, Hoebregs H, Smits G (2014) On the presence of inulin and oligofructose as natural ingredients in the western diet. C R C Crit Rev Food Technol 35 (6):525–552 Lopes SM, Krausová G, Rada V, Gonçalves JE, Gonçalves RA, de Oliveira AJ (2015) Isolation and characterization of inulin with a high degree of polymerization from roots of Stevia rebaudiana (Bert.) Bertoni. Carbohydr Res 411:15–21 Malik A (2012) Molecular cloning and in silico characterization of fructansucrase gene from Weissella confusa MBFCNC-2 (1) isolated from local beverage. Asia Pac J Mol Biol Biotechnol 20(1):33–42 Malik A, Hapsari MT, Ohtsu I, Ishikawa S, Takagi H (2015) Cloning and heterologous expression of the ftfCNC-2 (1) gene from Weissella confusa MBFCNC-2 (1) as an extracellular active fructansucrase in Bacillus subtilis. J Biosci Bioeng 119(5):515–520 Maumela P, Rensburg EV, Chimphango AFA, Grgens JF (2020) Sequential extraction of protein and inulin from the tubers of Jerusalem artichoke (Helianthus tuberosus L.). J Food Sci Technol Mysore 57(2):775–786 Mena-Arizmendi A, Alderete J, Águila S, Marty A, Miranda-Molina A, López-Munguía A, Castillo E (2011) Enzymatic fructosylation of aromatic and aliphatic alcohols by Bacillus subtilis levansucrase: reactivity of acceptors. J Mol Catalysis B Enzymatic 70(1–2):41–48 Mensink AM, Hinrichs JWL, Maarschalk VK, Frijlink WH (2015a) Inulin, a flexible oligosaccharide. II: review of its pharmaceutical applications. Carbohydr Polym 134:418–428 Mensink MA, Frijlink HW, van der Voort MK, Hinrichs WL (2015b) Inulin, a flexible oligosaccharide I: review of its physicochemical characteristics. Carbohydr Polym 130:405–419 Meyer D, Bayarri S, Tarrega A, Costell E (2011) Inulin as texture modifier in dairy products. Food Hydrocoll 25(8):1881–1890 Micka A, Siepelmeyer A, Holz A, Theis S, Schn C (2017) Effect of consumption of chicory inulin on bowel function in healthy subjects with constipation: a randomized, double-blind, placebocontrolled trial. Int J Food Sci Nutr 68(1):82–89 Milani E, Koocheki A, Golimovahhed QA (2011) Extraction of inulin from burdock root (Arctium lappa) using high intensity ultrasound. Int J Food Sci Technol 46(8):1699–1704 Moradali MF, Rehm BH (2020) Bacterial biopolymers: from pathogenesis to advanced materials. Nat Rev Microbiol 18:195–210 Moral SD, Olvera C, Rodriguez ME, Munguia AL (2008) Functional role of the additional domains in inulosucrase (IslA) from Leuconostoc citreum CW28. BMC Biochem 9(1):6 Nemeth C, Andersson AAM, Andersson R, Mangelsen E, Sun C, Åman P (2015) Relationship of grain Fructan content to degree of polymerisation in different barleys. Food Nutr Ences 5 (6):581–589 Ni D, Zhu Y, Xu W, Bai Y, Zhang T, Mu W (2017) Biosynthesis of inulin from sucrose using inulosucrase from lactobacillus gasseri DSM 20604. Int J Biol Macromol 109:1209–1218 Ni D, Xu W, Bai Y, Zhang W, Zhang T, Mu W (2018) Biosynthesis of Levan from sucrose using a thermostable levansucrase from lactobacillus reuteri LTH5448. Int J Biol Macromol 113:29–37 Ni D, Xu W, Zhu Y, Zhang W, Zhang T, Guang C, Mu W (2019) Inulin and its enzymatic production by inulosucrase: characteristics, structural features, molecular modifications and applications. Biotechnol Adv 37:306–318 Ni D, Zhu Y, Xu W, Pang X, Lv J-P, Mu W (2020a) Production and physicochemical properties of food-grade high molecular weight lactobacillus inulin. J Agric Food Chem 68:5854–5862 Ni D, Zhu Y, Xu W, Pang X, Mu W (2020b) Production and physicochemical properties of foodgrade high-molecular-weight lactobacillus inulin. J Agric Food Chem 68(21):5854–5862

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Nwafor IC, Shale K, Achilonu MC (2017) Chemical composition and nutritive benefits of chicory (Cichorium intybus) as an ideal complementary and/or alternative livestock feed supplement. Sci World J 2017:7343928 Olivaresillana V, Wacherrodarte C, Borgne SL, Lopezmunguia A (2002) Characterization of a cellassociated inulosucrase from a novel source: a Leuconostoc citreum strain isolated from Pozol, a fermented corn beverage of Mayan origin. J Ind Microbiol Biotechnol 28(2):112–117 Olivares-Illana V, López-Munguía A, Olvera C (2003) Molecular characterization of inulosucrase from Leuconostoc citreum: a fructosyltransferase within a glucosyltransferase. J Bacteriol 185 (12):3606–3612 Ortiz-Soto ME, Olivares-Illana V, López-Munguía A (2004) Biochemical properties of inulosucrase from Leuconostoc citreum CW28 used for inulin synthesis. Biocatal Biotransformation 22(4):275–281 Ortiz-Soto ME, Porras-Domínguez JR, Seibel J, López-Munguía A (2019) A close look at the structural features and reaction conditions that modulate the synthesis of low and high molecular weight fructans by levansucrases. Carbohydr Polym 219:130–142 Ozimek LK, Kralj S, Kaper T, Van Der Maarel MJ, Dijkhuizen L (2006) Single amino acid residue changes in subsite 1 of inulosucrase from lactobacillus reuteri 121 strongly influence the size of products synthesized. FEBS J 273(17):4104–4113 Peña-Cardeña A, Rodríguez-Alegría ME, Olvera C, Munguía AL (2015) Synthesis of Fructooligosaccharides by IslA4, a truncated inulosucrase from Leuconostoc citreum. BMC Biotechnol 15(1):1–9 Petkova NT, Denev P (2018) Characterization of inulin from dahlia tubers isolated by microwave and ultrasound-assisted extractions. Int Food Res J 25(5):1876–1884 Petkova N, Sherova G, Denev P (2018) Characterization of inulin from dahlia tubers isolated by microwave and ultrasound-assisted extractions. Int Food Res J 25(5):1876–1884 Pijning T, Anwar MA, Böger M, Dobruchowska JM, Leemhuis H, Kralj S, Dijkhuizen L, Dijkstra BW (2011) Crystal structure of inulosucrase from lactobacillus: insights into the substrate specificity and product specificity of GH68 fructansucrases. J Mol Biol 412(1):80–93 Raga-Carbajal E, Carrillo-Nava E, Costas M, Porras-Dominguez J, López-Munguía A, Olvera C (2016) Size product modulation by enzyme concentration reveals two distinct Levan elongation mechanisms in Bacillus subtilis levansucrase. Glycobiology 26(4):377–385 Rao M, Gao C, Xu L, Jiang L, Xu Y (2019) Effect of inulin-type carbohydrates on insulin resistance in patients with type 2 diabetes and obesity: a systematic review and meta-analysis. J Diabetes Res 2019(2):1–13 Rehm BH (2010) Bacterial polymers: biosynthesis, modifications and applications. Nat Rev Microbiol 8(8):578–592 Roberfroid M, Slavin J (2000) Nondigestible oligosaccharides. Crit Rev Food Technol 40 (6):461–480 Rodríguez-Alegría ME, Enciso-Rodríguez A, Ortiz-Soto ME, Cassani J, Munguía AL (2010) Fructooligosaccharide production by a truncated Leuconostoc citreum inulosucrase mutant. Biocatalysis 28(1):51–59 Rosell K, KG R (1974) An inulin-like fructan produced by Streptococcus mutans, strain JC2. Acta Chem Scand Series B Organic Chem Biochem 28:589 Rosica V, Petya K, Inés M, Jens W, Michael G (2018) Inulin-type fructans improve active ulcerative colitis associated with microbiota changes and increased short-chain fatty acids levels. Gut Microbes 10(3):334–357 Ruiz-Aceituno L, Sanz ML, Blanca DLR, Muñoz R, Kolida S, Jimeno ML, Moreno FJ (2017) Enzymatic synthesis and structural characterization of theanderose through transfructosylation reaction catalyzed by levansucrase from Bacillus subtilis CECT 39. J Agric Food Chem 65 (48):10505–10513 Schütz K, Muks E, Carle R, Schieber A (2010) Separation and quantification of inulin in selected artichoke (Cynara scolymus L.) cultivars and dandelion (Taraxacum officinale WEB. ex

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Chapter 11

Amylosucrase: A Versatile Sucrose-Utilizing Transglucosylase for Glycodiversification Yuqing Tian, Qiuming Chen, and Wenli Zhang

11.1

Introduction

Carbohydrates are undoubtedly the most abundant organic compounds in nature, with diverse chemical structures and biological functions. The diversity of carbohydrates is determined by a series of enzymes, named Carbohydrate-Active enZymes (CAZymes), that catalyze the assembly, cleavage, and modification of carbohydrates (Lombard et al. 2014). CAZymes can be classified into glycoside hydrolases (GH), glycosyltransferases (GT), polysaccharide lyases (PL), carbohydrate esterases (CE), carbohydrate binding modules (CBM), and auxiliary activities (AA) which accounts for 47%, 36%, 1%, 5%, 10%, and 1%, respectively (André et al. 2014). With the rapidly increased number of CAZymes members, many facile enzyme-based synthetic routes to complex carbohydrates can be accessible. However, numerous enzymatic or chemo-enzymatic pathways are still hampered by the lack of some desirable catalysts, and thus CAZymes with requisite properties and specificities are urgently demanded (Benkoulouche et al. 2019). Among the CAZymes, GH family enzymes attract increasing attention in terms of glycodiversification recently, owing to their ability to utilize abundant biomasses such as starches and sucrose instead of expensive nucleotide-active sugars as donor substrates. Sucrose (2-O-α-D-glucopyranosyl-D-fructose), which naturally exists in sugar cane and sugar beet, has been considered as the most available low molecular weight carbohydrate and is an indispensable sweetener in the diet (André et al. 2010). However, excessive intake of sucrose is often thought to be related to the occurrence of some chronic diseases, such as type 2 diabetes (Mu et al. 2014). Therefore, it has attracted attention that converts sucrose to functional sweeteners or other highly valuable carbohydrates (Tian et al. 2019a). In the GH 70 and 13 families, there are

Y. Tian · Q. Chen · W. Zhang (*) State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China e-mail: [email protected]; [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_11

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some enzymes known as sucrose-utilizing transglucosidases that can utilize sucrose as the glucosyl donor and catalyze the transfer of a glucosyl group to an acceptor molecule such as other sugars or polyols (Daudé et al. 2014). Most of sucroseutilizing transglucosidases belong to GH70 family, such as dextransucrase, alternansucrase, reuteransucrase, and mutansucrase. In addition to these enzymes belonging to the GH70 family, there is an atypical member of sucrose-utilizing transglucosidases, amylosucrase (ASase), belonging to the GH13 family. The GH13 family, the largest sequence-based GH family, is also known as the α-amylase family (Stam et al. 2006). GH13 family contains a series of enzymes possessing the ability to act on α-glycosidic bonds, such as α-amylase, cyclodextrin glucanotransferase, pullulanase, and sucrose phosphorylase which share a (β/α)8-barrel structure with ASase (Janeček and Gabriško 2016). ASase (EC 2.4.1.4), also known as sucrose-1,4-α-glucan glucosyltransferase and sucrose:1,4-α-D-glucan 4-α-D-glucosyltransferase, can catalyze various types of reactions including polymerization, isomerization, transglycosylation, hydrolysis, and disproportionation. In 1946, ASase was first discovered in Neisseria perflava grown on sucrose-containing medium and named after its ability to synthesize the amylose-like polymer from sucrose with no need of any nucleotide-activated sugars (Hehre and Hamilton 1946). In 1997, ASase from N. polysaccharea (NP-ASase) was first sequenced, and heterologous expressed in Escherichia coli XL1-Blue (Büttcher et al. 1997), which make it possible to biochemically characterize this enzyme. After 2000, the crystal structures of NP-ASase were solved in its apo form (Skov et al. 2000) and in complex with glucose, fructose, sucrose, and oligosaccharide (Mirza et al. 2001, 2002), and its catalytic mechanism was then explained. Subsequently, the ability of ASase to catalyze the isomerization of sucrose (de Montalk et al. 2000) and the disproportionation of maltooligosaccharides (MOS) (Albenne et al. 2002) was discovered. After 2005, ASases were also identified from other strains other than the genus Neisseria, which indicates that their distribution in nature is wider than previously thought (Pizzut-Serin et al. 2005). Then, molecular modifications through random mutagenesis and rational design was started to improve the enzymatic properties of reported ASases (van der Veen et al. 2004; Daudé et al. 2019). Around 2010, ASase was used to catalyze the glycosylation of salicin (Jung et al. 2009). Since then, in addition to sugars, various polyphenolic compounds and glycoside molecules have also been proved to be effective acceptors of ASase, which indicates that ASase can be applied to the modification of many bioactive compounds. As a multifunctional sucrose-utilizing transglucosylase, ASase has numerous applications in producing functional sweeteners, dietary fibers, carbohydrate-based carrier materials, bioactive compounds, and cell surface oligosaccharides. This chapter will in detail exhibit the characteristics of ASase, including the catalytic mechanism, reaction type, substrate specificity, heterologous expression, and other biochemical properties, and its structural features and structure-based molecular modification in activity, product specificity, acceptor specificity, and thermostability. The applications of ASase in glycodiversification recently are also systematically classified and summarized based on its substrate and product.

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11.2

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Characteristics

As shown in Table 11.1, ASase is often found in the genus Neisseria, Deinococcus, and Bifidobacterium and exists in other strains, including Arthrobacter chlorophenolicus, Alteromonas macleodii, Cellulomonas carboniz, Calidithermus timidus, Methylomicrobium alcaliphilum, Methylobacillus flagellates, Synechococcus sp., Truepera radiovictrix, and Xanthomonas campestris, as well as in the metagenome of a thermal aquatic habitat. Among them, NP-ASase and DG-ASase have been well-studied and were usually used in related applications.

11.2.1 Catalytic Mechanism ASase belongs to GH subfamily 13_4 and follows the double-displacement mechanism as other members of GH13 family (Stam et al. 2006). First, the glycosyl donor substrate, sucrose, enters the catalytic active site with the glucosyl ring binding at the
1 subsite and the fructosyl ring binding at the +1 subsite. Next, glutamic acid (Glu 328 in NP-ASase) protonates the oxygen in the glycosidic bond and assists aspartic acid (Asp 286 in NP-ASase) to exert a nucleophilic attack on the anomeric carbon in α-configuration. Simultaneously, the covalent β-glucosyl-enzyme intermediate is formed, and the fructose is released. This step goes through an oxocarbenium ion transition state. Second, glutamic acid (Glu 328 in NP-ASase) acts as a base to activate a glycosyl acceptor which attacks the anomeric carbon. Then, the covalent bond is broken, and the anomeric carbon thus returns to the α-configuration (Jensen et al. 2004; Skov et al. 2006). When the glycosyl donor is sucrose and the glycosyl acceptor is water, fructose, glucose, or extrinsic acceptors, the reaction is called hydrolysis, isomerization, polymerization, or tranglycosylation, respectively (Fig. 11.1). In addition, when using MOS as the sole substrate, ASase can catalyze the transfer of the glucosyl unit between MOS molecules, which is known as disproportionation reaction (Albenne et al. 2002; Schneider et al. 2011). Therefore, reaction ASase that mainly catalyzes largely depends on the enzyme itself and reaction conditions used, which will be exhibited in the next part in detail.

11.2.2 Substrate Specificity For a transglucosylase, the substrate specificity includes the specificity of glycosyl donor substrate and the specificity of glycosyl acceptor substrate. ASase has high specificity for sucrose as the glycosyl donor. Daudé et al. (2013a) tested the donor substrate specificity of NP-ASase toward 11 sucrose analogues, including 3-ketosucrose, raffinose, sucralose, sucrose-6-acetate, sucrose-6,60 acetate, glucose-1-phosphate, α-D-galactopyranosyl-1,2-β-D-fructofuranoside,

Enzyme ACASase AMASase BDASase BI-ASase BPASase BTASase CCASase CTASase DGASase DRdASase DRpASase MAASase METASase MFASase Dimer Dimer

WP_018466847.1 ABF44874.1 WP_010887578.1 – CCE22312.1 QCZ35432.1 ABE50875.1

C. timidus DSM 17022

D. geothermalis DSM 11300

D. radiodurans ATCC 13939

D. radiopugnans ATCC 19172

M. alcaliphilum 20Z

Uncultured bacterium

M. flagellatus KT

Dimer

Monomer

Dimer

Dimer

Tetramer

Monomer

KGM11272.1

C. carbonis T26

45

60

30

40

30

45

55

40

50

–

WP_044279707.1

B. thermophilum ATCC 25525

45 30

– –

WP_101026183.1 WP_118284197.1

B. longum 51A B. pseudocatenulatum

35

–

WP_003842696.1

B. dentium

45

–

BAG82876.1

A. macleodii KCTC 2957

Topt ( C) –

Oligomeric state Monomer

Protein ID ACL41561.1

Microbial sources A. chlorophenolicus A6

Table 11.1 Comparison of enzymatic properties of recombinant ASase

50.6

–

–

50.7

–

58.9

74.3

47.8

61.9

48.2 49.0

47.8

48.1

Tm ( C) 42.6

60  C; 60 min 55  C; 4 min

50  C; 8 min –

60  C; 55 min 50  C; 5 min 70  C; 65 min 55  C; 408 min –

– –

Half-life 45  C; 3 min 45  C; 30 min –

8.5

9.0

8.0

8.0

8.0

8.0

Jeong et al. (2014)

Agarwal et al. (2019)

But et al. (2015)

Kim et al. (2014)

Pizzut-Serin et al. (2005)

Seo et al. (2008)

Tian et al. (2019b)

Wang et al. (2017)

7.0 7.0

Choi et al. (2019)

Kim et al. (2020b) Kim et al. (2020b)

Kim et al. (2020b)

Ha et al. (2009)

Reference Seo et al. (2012a)

6.0

5.0 8.0

5.0

8.0

pHopt 8.0

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a

CAA09772.1 EET43598.1 EFC51554.1 SMQ77851.1 ADI14158.1 AAY47880.1

N. polysaccharea ATCC 43768

N. sicca ATCC 29256

N. subflava ATCC 49275

Synechococcus sp. PCC 7002

T. radiovictrix DSM 17093

Xanthomonas campestris pv. campestris 8004

35a

30

–

–

45

–

45

45

–

Dimer

37

Monomer

–

69.6

–

–

–

49.6

65  C; 73 min –

50  C; 15 min 50  C; 9 min 50  C; 6 min –

7.5a

7.5

7.0

8.0

8.5

8.0

Zhu et al. (2019)

Perez-Cenci and Salerno (2014) Zhu et al. (2018a)

Park et al. (2018a)

Kim et al. (2019)

Büttcher et al. (1997)

These data were determined using sucrose as donor and hydroquinone as acceptor, while other data use sucrose as the sole substrate

NPASase NsiASase NSuASase SSASase TRASase XCASase

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Fig. 11.1 The reactions catalyzed by ASase using sucrose as the sole substrate

α-D-xylopyranosyl-1,2-β-D-fructofuranoside, p-nitrophenyl-α-D-glucopyranoside ( pNP-Glc), pNP-α-D-galactoside, and pNP-α-D-mannoside. These 11 potential donors were chosen based on the preliminary molecular modeling studies; however, only pNP-Glc could be recognized as glycosyl donor, showing that its
1 subsite has high specificity for the glucosyl moiety. ASase has broad specificity for the glycosyl acceptors substrate, showing that its +1 subsite has high plasticity. Generally, water, fructose, glucose, and MOS were known as natural acceptors of ASase. In addition to these natural acceptors, there are more than 50 compounds that can be recognized as effective glycosyl acceptors by ASase, including some monosaccharides, oligosaccharides, starches, sugar alcohols, phenolic compounds, polyphenolic compounds, polyhydroxy compounds, and glycoside molecules. These effective acceptors will be exhibited in detail in Sect. 11.4. However, some compounds were still recognized as poor acceptors of ASase so far in the literature, such as L-fucose (Daudé et al. 2013a), cinnamic acid (Kim et al. 2016b), orientin, luteolin-30 ,7-diglucoside, isorhoifolin, chrysin (Jang et al. 2018), 3-hydroxyflavone, 5-hydroxyflavone, 7-hydroxyflavone, 20 -hydroxyflavanone, 7-flavonol, galangin (Rha et al. 2019a), taxifolin, 6-hydroxydaidzein, 8-methyldaidzein, formononetin, biochanin A (Overwin et al. 2016), morin, myricetin, quercetin, and kaempferol (Overwin et al. 2015b). In addition, ASases from different microbial sources also have some difference in substrate specificity. For example, DG-ASase can recognize apigenin, baicalein (Rha et al. 2019a), daidzein (Jung et al. 2020b), and luteolin (Jang et al. 2018) as effective glycosyl acceptors, while NP-ASase cannot (Overwin et al. 2015b, 2016).

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11.2.3 Effect of Reaction Conditions As shown in Sect. 11.2.1, ASase follows double-displacement mechanism and will release fructose in its first step when in reaction with sucrose. Therefore, the enzyme activity of ASase is usually investigated using the sucrose as the sole substrate and by estimating the amounts of released fructose. The enzyme activity calculated from the release of fructose is also called total activity (van der Veen et al. 2004; Wang et al. 2017). And because the hydrolysis reaction is the only reaction that releases glucose, the amount of glucose released is used to calculate the hydrolysis activity (Ha et al. 2009). The total activity minus hydrolysis activity is used to calculate the polymerization activity or tranglycosylation activity in some literature (van der Veen et al. 2006; Wang et al. 2017; Zhu et al. 2018a), while some other researchers evaluated its polymerization and isomerization ability directly from the amounts of released amylose-like polymer and sucrose isomers, respectively (Choi et al. 2019; Kim et al. 2019). As shown in Table 11.1, most of ASases showed pH optima at neutral or alkaline pH, while BD-ASase, BI-ASase, and BP-ASase from the genus Bifidobacterium showed highest enzyme activity at slightly acidic pH. The optimal pH of the total activity and hydrolysis activity of some ASases is the same, but there are also some enzymes that prefer catalyzing different reactions at different pH. For example, CC-ASase exhibited highest hydrolysis activity and total activity at pH 5.0 and at pH 7.0, respectively (Wang et al. 2017), and AM-ASase showed highest hydrolysis activity and total activity at pH 7.0 and at pH 8.0, respectively (Ha et al. 2009). Metal ions usually show negative effects on the activity of ASase. For example, the enzyme activity of CC-ASase was completely inhibited in the presence of 1 mM Cu2+ (Wang et al. 2017). Agarwal et al. (2019) investigated the effect of metal ions on isomerization activity of MET-ASase and found that the addition of 5 mM of Cu2 + , Zn2+, or Ni2+ can cause its turanose production ability completely loss. Most of ASases showed optimum temperature between 30 and 50  C, except the CT-ASase and MET-ASase showing optimum temperature at 55 and 60  C, respectively. Like pH, the reaction temperature will also affect the product ratio. For example, Seo et al. (2008) reported that the optimal temperature of DG-ASasecatalyzed hydrolysis reaction was 20  C higher than that of transglycosylation reaction. TR-ASase showed highest hydrolysis activity and transglycosylation activity at 55  C and 45  C, respectively (Zhu et al. 2018a). Thermostability is an important property needed for industrial applications of an enzyme. However, most of ASases cannot maintain 50% of its activity after incubating for 15 min at 50  C, which may limit its application. Recently, some ASases with high thermostability have been found from thermophilic microorganisms or thermal habitat, such as TR-ASase (Zhu et al. 2018a), CT-ASase (Tian et al. 2019b), and MET-ASase (Agarwal et al. 2019). In addition, several structural studies showed that the thermostability may be associated with its oligomeric state, which will be presented in detail in Sect. 11.3.1.

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11.2.4 GRAS-Grade Expression Recent studies also focused on the food-grade expression of ASase, such as the expression in Corynebacterium glutamicum (Chin et al. 2020) and Bacillus subtilis (Kim et al. 2018; Su et al. 2020). In 2018, Kim et al. (2018) attempted to heterogeneously express ASase in B. subtilis KCTC3135 using vector pUBRTAMY. DG-ASase has been successfully expressed and showed similar enzymatic properties as expressed in E. coli, while NP-ASase cannot be expressed functionally. In 2020, Su et al. (2020) successfully expressed NP-ASase in B. subtilis WS11 using the modified expression vector pHY300PLK. In addition, DG-ASase has been functionally expressed in C. glutamicum ATCC 13032 using vector pXMJ19 and interestingly showed a 1.onefold enhancement in luteolin transglucosylation activity than that expressed in E. coli owing to a lower strand/helix ratio (Chin et al. 2020).

11.3

Structure-Based Molecular Modification

11.3.1 Structural Features At present, the three-dimensional structures of NP-ASase (Skov et al. 2001), DG-ASase (Guerin et al. 2012), and DRd-ASase (Skov et al. 2013) have been resolved. Among them, NP-ASase has many structures in complex with various ligands, such as sucrose (Mirza et al. 2001), fructose (Mirza et al. 2002), turanose (Guerin et al. 2012), and MOS (Skov et al. 2002) and remains the most investigated ASase in terms of structure so far. As shown in Fig. 11.2, ASase can be divided into five domains in structure. Taking the structure of NP-ASase as the example, the domain A is composed of residues 90–183, 260–394, and 460–553 and forms a (β/ α)8-barrel structure which is common in GH13 family. The domains B and C are also shared in GH13 family, which are composed of residues 184–259 and residues 554–628, respectively. The domain B is located between β3-strand and α3-helix of the (β/α)8-barrel, while the domain C is located at C-terminal. The domains N and B0 are specific to ASase. The domain N is located at N-terminal consisting of residues 1–89, and the domain B0 is composed of residues 395–459 inserted in β-strand 7 and α-helix 7. Compared with monomer NP-ASase, both DG-ASase and DRd-ASase exist in the form of dimers in solution and show a dimeric assembly in their crystal structure. A set of residues are found to be involved in dimerization and can be divided into 7 regions. Taking the structure of DG-ASase as the example (Fig. 11.2), the region 1 to 7 are composed of residues 21–35, 73–88, 318–320, 334–344, 378–385, 560–568, and 584–590. The salt bridges and hydrogen bonds that are formed between these seven regions are believed to play an important role in stabilizing the dimer structure of the enzyme (Guerin et al. 2012; Skov et al. 2013).

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Fig. 11.2 The three-dimensional structure of DG-ASase (PDB ID: 3UCQ). Domain A, B, B0 , C, N are shown in blue, green, orange, pink, and yellow, respectively. Seven regions involved in dimerization are shown in black, while other part of chain B are shown in light blue

11.3.2 Enhancing Activity and Thermostability Activity and thermostability are two key factors for the industrial application of an enzyme. Recently, Seo et al. (2019) constructed the site-saturated mutagenesis of DG-ASase at Pro219, Phe225, and Ala226 in loop 3 of the B domain and screened out the mutant A226N, which has improved polymerization activity due to reduced flexibility of loops. Salt bridges and hydrogen bonds often have influence on the thermal stability of ASase. The double mutant of NP-ASase, R20C-A451T, have a tenfold increased half-life at 50  C owing to the reorganization of surface salt bridges and the introduction of a hydrogen-bonding interaction (Emond et al. 2008). Tian et al. (2019b) conducted a truncation mutation to break the salt bridges and hydrogen bonds in subunit interface of CT-ASase, and these mutants showed a 10  C decrease in melting temperature (Tm), which indicated the importance of these interactions for thermostability. In addition, Daudé et al. (2013b) reported that reshaping the subsite
1 can change the melting temperature of the NP-ASase by 8  C.

11.3.3 Altering Substrate and Product Specificity As shown in Sect. 11.2.2, ASase has high specificity for sucrose and can also use pNP-Glc as glycosyl donor substrate. Recently, a mutant of NP-ASase (D37HV331D-D526N-V609I) was isolated and showed 25 times enhancement in catalytic efficiency toward pNP-Glc as the donor substrate (Daudé et al. 2019). In addition to donor substrate, the specificity of acceptor substrate also attracts increasing attention.

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Through computer-aided engineering, the active site of NP-ASase have been successfully reshaped, and several mutants were isolated with novel acceptor specificity towards intermediate saccharides for synthesis of Shigella flexneri antigenic oligosaccharides (Champion et al. 2009a, b, 2012; Vergès et al. 2015; Daudé et al. 2019). Furthermore, owing to conformational changes in loop 4, two mutants of DRp-ASase, Q299K and Q299R, showed tenfold and fourfold increase in rutin glycosylation product (Kim et al. 2016b). As shown in Table 11.2, several mutants with desirable product specificity were obtained through random and site-directed mutagenesis. For example, as early as 2003, Arg226 was found to have a great effect on the product specificity of NP-ASase (Albenne et al. 2003). Then, in 2014, by constructing saturation mutation of NP-ASase on Arg226, R226N with a tenfold increase in catalytic efficiency was obtained, and it was confirmed that Arg226 plays an important role in oligosaccharide elongation (Cambon et al. 2014). Through directed evolution, mutant E227G of NP-ASase were obtained with improved product specificity for polymer, which can synthesize amylose with average DP (degree of polymerization) of 108 from 20 mM sucrose (van der Veen et al. 2006). Recently, Vergès et al. (2017b) reported that after reshaping the active site, two novel sucrose derivatives of potential interest, erlose (α-D-glucosyl-1,4-α-D-glucosyl-1,2-β-D-fructose) and panose (α-D-glucosyl-1,6-αD--glucosyl-1,4-α-D-glucose), can be synthesized by NP-ASase.

11.4

Versatile Functions and Its Applications

As introduced in Sect. 11.2.1, ASase can catalyze several types of reactions and thus has numerous applications in producing functional sweeteners, dietary fibers, carbohydrate-based carrier materials, bioactive compounds, and cell surface oligosaccharides. This section will systematically classify and summarize the applications of ASase based on its substrate and product. For example, when using sucrose as the sole substrate, ASase mainly catalyzes polymerization reaction of glucose unit to synthesize α-1,4 glucan. This amylose-like polymer can be directly used in the preparation of amylose microparticles or being used as the substrate of some starch-converting enzymes to produce some functional oligosaccharides composed of different numbers glucose units. ASase can also recognize some sugars or polyphenolic compounds as the glycosyl acceptor and catalyze the transfer of the glucose group from the donor sucrose to the acceptor. When using fructose, oligosaccharides, or starches as the acceptors, ASase can produce turanose, immunogenic oligosaccharides, or starches with tailored digestibility, respectively. When using bioactive compounds as the acceptors, ASase can modify these compounds to increase their stability, solubility, and bioavailability.

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Table 11.2 Summary of the structure-based molecular modification of ASases Characteristics Activity

Thermostability

Substrate

Enzyme NPASase

Mutant V389LN503I

Changed properties Total activity (U/mg)

Wildtype 1.7

Mutant 6.4

NPASase

R20CF598S

Total activity (U/mg)

1.7

7.2

NPASase

N387D

Relative activity (50  C)

5%a

60%

NPASase

P351S

t1/2 (50  C)/min

1

25

NPASase

R20CA451T

t1/2 (50  C)/min

3

32

NPASase

A170VQ353L

t1/2 (50  C)/min

3

10

NPASase

H392P

Tm/ C

48

50

NPASase

D37HV331DD526NV609I I228Y

kcat/Km( pNPGlc) / s
1 mM
1

0.03

0.76

A’b conversion ratio

0%

44%

NPASase

F290K

D’c conversion ratio

6%

78%

NPASase

A289PF290L

kcat/Km(D’c) / s
1 mM
1

0.002

0.790

NPASase

F3d

D0 A’e conversion ratio

0%

4.3%

DRpASase NPASase

Q299K

Rutin conversion ratio Insoluble glucan ratio

Onefold

Tenfold

45%

0%

45

108

NPASase

Product

NPASase

D394A/ R446A/ R415A E227G

Average DP

Reference van der Veen et al. (2004) van der Veen et al. (2004) van der Veen et al. (2006) Emond et al. (2007) Emond et al. (2008) Emond et al. (2008) Daudé et al. (2013b) Daudé et al. (2019) Champion et al. (2009a) Champion et al. (2009a) Champion et al. (2012) Vergès et al. (2015) Kim et al. (2016b) Albenne et al. (2003) van der Veen et al. (2006) (continued)

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Table 11.2 (continued) Characteristics

Enzyme NPASase

Mutant R226N

Changed properties Insoluble glucan ratio

Wildtype 44%

Mutant 82%

Reference Cambon et al. (2014)

NPASase

30H3b

MOS ratio

78%

100%

NPASase

37G4b

Erlose ratio

0%

46%

NPASase

39A8b

Turanose ratio

19%

46%

NPASase DGASase

G396S

Turanose yield/ gL
1 Polymerization/ hydrolysis ratio

349

410

0.6

0.9

Vergès et al. (2017a) Vergès et al. (2017b) Vergès et al. (2017b) Su et al. (2020) Seo et al. (2019)

A226N

a

These results are estimates from the chart A’, methyl α-L-rhamnopyranoside c 0 D , allyl 2-N-acetyl-2-deoxy-α-D-glucopyranoside d 37G4 refers to a mutant containing 10 mutations, R226K-I228V-A289I-F290Y-E300I-V331TG396S-T398V-Q437R-N439D. 39A8 refers to a mutant containing 11 mutations, R226K-I228VA289I-F290Y-E300I-V331T-G396S-T398V-Q437S-N439D-C445R. 30H3 refers to a mutant containing 9 mutations, R226K-I228V-A289I-F290Y-E300I-V331T-Q437S-N439D-C445A. F3 refers to a mutant containing 7 mutations, R226L-I228V-F290Y-E300V-V331T-G396S-T398V e ED’A’, allyl (2-deoxy-2-trichloroacetamido-β-D-glucopyranosyl)-1,2-α-L-rhamnopyranoside b

11.4.1 Reactions with Sucrose As the Sole Substrate 11.4.1.1

Utilization of Amylose-Like Polymer

ASase can catalyze the polymerization reaction using sucrose as the sole substrate to produce glucans linked by α-1,4 bonds, also known as α-1,4 glucans, and release fructose at the same time. This amylose-like polymer can spontaneously selfassemble in an aqueous solution to form microparticles due to the hydrogen bond and hydrophobic interaction between chains of the polymer (Lim et al. 2014, 2015). Therefore, this biological method of enzymatic polymerization and self-assembly was investigated to synthesize some amylose-based microparticles, with the advantage that the product can be obtained in one step. In 2014, Lim et al. (2014) reported a novel method for preparing amylose microbeads and the amylose microbeads single walled carbon nanotube composite microbeads by enzymatic synthesis using DG-ASase. Then, Lim et al. (2015) used a similar method to synthesize amylose magnetic beads possessing high binding capacity and recyclability, which can be used in column-free purification of target proteins. After being fused with maltose-binding protein and streptococcal

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protein G, the amylose magnetic beads can also be used to isolate target bacteria from some food matrix, such as milk (Lim et al. 2016a). This kind of enzymatically synthesized α-1,4 glucans can also self-assemble with β-carotene in an aqueous solution to generate amylose microparticle-embedded β-carotene (Letona et al. 2017). In addition, some studies have focused on optimizing the reaction conditions to obtain amylose-based microparticles with better morphology, size, and other properties. Short-chain fatty acids can induce the self-association of α-1,4 glucans through hydrogen bonds and are ejected by the amylose microparticles after they are formed (Lim et al. 2016b). Luo et al. (2018) used dextran-coated iron oxide nanoparticles as a seeding agent to obtain well-defined spherical and stable superparamagnetic amylose microparticles. Letona et al. (2019) used lecithin as a steric surfactant to increase the nucleation rate of α-1,4 glucans and thereby obtained amylose microparticles with well-defined size.

11.4.1.2

Synergistic Action with Starch-Converting Enzymes

Starch-converting enzymes play an indispensable role in glycodiversification; however, using starch as the substrate, the bioconversion process usually require a timeconsuming and energy-intensive substrate dissolving step (Kim et al. 2011b; Koh et al. 2016). Recent studies revealed that through the synergistic action of starchconverting enzymes and ASase, highly soluble sucrose can be used as the raw material instead of starch to synthesis some functional carbohydrates such as cycloamyloses, cyclodextrins, MOS, and trehalose. Cycloamyloses and cyclodextrins are cyclic oligosaccharides composed of different numbers of glucose units linked by α-1,4 bonds, which can form an inclusion complex with some hydrophobic and unstable materials, and thus widely used as solubilizing and stabilizing agents (Del Valle 2004). They are usually produced from starch using 4-α-glucanotransferase (EC 2.4.1.25) and cyclodextrin glucanotransferase (EC 2.4.1.19) as catalyst, respectively. Kim et al. (2011b) have investigated the bioconversion of sucrose to cycloamyloses by the synergistic action of ASase and 4-α-glucanotransferase and achieved a yield of 9.6% (w/w). One-pot synthesis of cyclodextrins has also been conducted using ASase and cyclodextrin glucanotransferase with a conversion ratio of 21.1% (w/w) (Koh et al. 2016). MOS refer to a series of oligosaccharides composed of 2 to 10 glucose units linked by α-1,4 linkages, which possess many physical and chemical properties, such as the high water-holding capacity and crystallization-prevention ability (Pan et al. 2017). Maltooligosaccharide-forming amylase is a well-known starchconverting enzyme which can hydrolyze starch to specifically synthesize MOS. Zhu et al. (2018b) have investigated the bioconversion of sucrose to MOS by the synergistic action of ASase and maltooligosaccharide-forming amylase and achieved a yield of 34.1% (w/w). Trehalose (1-O-α-D-glucopyranosyl-D-glucose) can also be produced from sucrose using multi-enzyme system containing ASase. Trehalose, a non-reducing

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disaccharide, is about 45% as sweet as sucrose (Elbein et al. 2003). It is of great interest for its low hygroscopicity, high glass transition temperature, and protein protection properties (Kim et al. 2011a). Jung et al. (2013) have investigated the one-step bioconversion of sucrose to trehalose using the combined cross-linked enzyme aggregates of ASase, maltooligosyltrehalose synthase (EC 5.4.99.15), and maltooligosyltrehalose trehalohydrolase (EC 3.2.1.141) and achieved a production yield of about 8% (w/w) in each cycle.

11.4.2 Reactions with Sucrose and Other Carbohydrates 11.4.2.1

Production of Alternative Sweeteners

Sucrose is an indispensable sweetener in the diet, but the excessive intake of sucrose may bring about some health problems (Mu et al. 2014). Therefore, alternative sweeteners which have physiological benefits and feasible production processes are urgently needed. Turanose (3-O-α-D-glucopyranosyl-D-fructose) and trehalulose (1-O-α-D-glucopyranosyl-D-fructose) are the product of the sucrose isomerization reaction catalyzed by ASase. These non-cariogenic sugars are about 50% as sweet as sucrose and have some physiological benefits such as slow digestion properties, anti-adipogenesis, and anti-inflammatory effects (Tian et al. 2019a; Kim et al. 2019). In general, the sucrose isomerization reaction catalyzed by ASase will produce turanose and trehalulose at the same time, but the yield of turanose is usually higher than or equal to that of trehalulose (Guerin et al. 2012). Therefore, research focused on the use of ASase to catalyze the synthesis of turanose. When using sucrose as the sole substrate, most ASases can convert about 20% sucrose into turanose (Tian et al. 2018). Recent studies have shown that when a high concentration of sucrose is used as a substrate, the yield of turanose often increases. Furthermore, adding additional fructose as a glycosyl acceptor substrate can also increase the yield of turanose (Wang et al. 2012; Park et al. 2016, 2019). In addition, the production of turanose was also studied using ASase immobilized on functionalized iron nanoparticles as biocatalyst and using unrefined or partially refined sugar as substrates, such as jaggery and muscovado (Agarwal et al. 2019). Furthermore, site-directed mutation of the acceptor binding site of ASase has been carried out to increase their product specificity toward turanose by increasing steric hindrance (Su et al. 2020). The mutant G396S of NP-ASase can produce 523 gL
1 turanose with the conversion rate of 76.5%. In addition, Kim et al. (2020b) reported that the trehalulose yield of BI-ASase was much higher than its turanose yield, suggesting the potential application of ASase in trehalulose production. When using 600 mM sucrose as the substrate, BI-ASase can convert about 58.8% sucrose into trehalulose after 140 h of reaction, while the conversion rate of turanose was only about 12.0%. Furthermore, some glucose analogues, including arabinose, allose, altrose, D-fucose, galactose, mannose,

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xylose, etc., can also be recognized as glycosyl acceptors by ASase with conversion rates of about 15–100% (Schneider et al. 2011; Daudé et al. 2013a). Although the products of these reactions and their physiological functions have not been well identified, it is sufficient to show that the monosaccharide acceptor reactions catalyzed by ASase have the potential to become new routes for the synthesis of functional oligosaccharides.

11.4.2.2

Synthesis of Microbial Cell Surface Oligosaccharides

Microbial cell surface oligosaccharides have aroused extensive attention because they can elicit an immune response and be used in the development of glycoconjugate vaccines (Benkoulouche et al. 2019). Since about a decade ago, ASase has been used in the chemo-enzymatic synthesis of S. flexneri cell surface oligosaccharides. Through semi-rational engineering approach, Champion et al. (2009b, 2012) isolated some NP-ASase mutants with new specificity toward methyl α-L-rhamnopyranoside or enhanced catalytic efficiency toward allyl 2-acetamido-2deoxy-α-D-glucopyranoside. These mutants can be used to synthesize oligosaccharides that mimic the O-antigen motif of S. flexneri serotypes 1b and 3a. In 2015, a multipoint mutant of NP-ASase were isolated with the ability to glycosylate allyl (2-deoxy-2-trichloroacetamido-β-D-glucosyl)-(1,2)-α-L-rhamnoside for the synthesis of S. flexneri serotypes 1a, 1b, or 1d O-SPs (Vergès et al. 2015). Recently, through neutral drift-based engineering, a NP-ASase variant were isolated with the ability to glycosylate α-L-Rhap-OMe for the synthesis of a precursor of antigenic oligosaccharides of S. flexneri (Daudé et al. 2019).

11.4.2.3

Modification of Starches

Starch is the main component of cereal grains and plays an indispensable role in the diet. Its fractions can be nutritionally classified into rapidly digestible starch (RDS), slowly digestible starch (SDS), and resistant starch (RS) (Englyst et al. 1992). Since the digestibility of starch has a certain impact on people’s health, many researchers have tried to modify starch using physical, chemical, or biological methods. In terms of biological approach, α-amylase, β-amylase, branching enzyme, and pullulanase are commonly used (Zhang et al. 2019). Since about a decade ago, ASase has also been applied to prepare starches with tailored digestibility (Ryu et al. 2010; Shin et al. 2010). ASase can recognize starches as the glycosyl acceptor and catalyze the glucose moiety transfer from donor substrate sucrose to the non-reducing end of acceptor starch and thereby elongate the starch chain. Among various microbial sources, NP-ASase and DG-ASase are commonly used in the modification of starches. Generally, after ASase treatment, the crystalline structure of starches changed to B-type, and their thermal properties have been improved, and the proportion of rapidly digestible starch in modified product was significantly reduced. Up to now,

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ASase has been investigated for the modification of potato starch (Kim et al. 2020a), chestnut starch (Lee et al. 2020), rice starch (Jung et al. 2020a), corn starch (Zhang et al. 2018), adlay starch (Kim et al. 2016a), and barley starch (Kim et al. 2015). Among them, ASase-treated rice starches have shown slow glucose generation rates estimated by mammalian α-glucosidase (Jung et al. 2020a), while ASase-modified chestnut starch has shown positive effect on attenuating obesity in diet-induced obese mice (Lee et al. 2018, 2020). In addition, ASase can also be applied to prepare some novel materials from starch, such as microparticles, nanohydrogels, and other carbohydrate-based carriers. Recently, Zhang et al. (2019) modified maize amylopectin through combined action of ASase and pullulanase and obtain novel starch microparticles which contain 67.2% of resistant starch component. Jung et al. (2019b) modified normal corn starch through combined action of β-amylase and ASase and obtain an α-glucan-coated porous starch granules showing a loading efficiency of 20.9  0.8% toward crocin. Nguyen Doan et al. (2019) used ASase-modified corn starch as a texture modifier to obtain rice starch gels with hardness increased to 60%. Lim et al. (2019) used ASase-modified waxy corn starch to prepare starch–lipid complexes with myristic acid and palmitic acid. Zhang et al. (2020) reported a novel nanohydrogels with controlled size prepared by complexing ASase-modified maize amylopectin with sodium palmitate.

11.4.3 Reactions with Sucrose and Bioactive Compounds 11.4.3.1

Glycosylation of Flavonoids

Flavonoids refer to a large class of bioactive compounds sharing a 15-carbon skeleton which consists of two phenyl rings named ring A and B and a heterocyclic ring named ring C (Xiao et al. 2014). Therefore, the content related to flavonoids will be introduced separately in this section. Flavonoids naturally exist in plant and fungus and possess numerous physiological effects, such as anti-oxidation, antiinflammatory, anti-obesity, etc., and thus widely used as dietary supplements and antioxidants in food industry (Rha et al. 2019b). However, most of flavonoids have poor bioavailability, which brings difficulties to their practical applications (Pandey et al. 2016). The glycosylation of flavonoids has been proved to increase their solubility and bioavailability, as well as to protect them from oxygen and light degradation (Slámová et al. 2018). Leloir glycosyltransferases are commonly used as transglycosylation tools of flavonoids, but they need expensive activated sugars as glucosyl donors. Recently, transglycosylation reactions using Non-Leloir glycosyltransferases, such as ASase, cyclodextrin glucanotransferase, and glucansucrases (EC 2.4.1.5 or 140), have also attracted the attention of researchers due to the low price of their substrates (Hofer 2016). Among various microbial sources, NP-ASase (Kim et al. 2018), DG-ASase (Chin et al. 2020), and DRp-ASase (Kim et al. 2016b) have been investigated in the

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glycosylation of flavonoids. As shown in Table 11.3, ASase has been used in the modification of 18 flavonoid aglycones and 4 flavonoid glycosides, including the members of flavone, flavanone, flavonol, flavanol, isoflavone, and chalcone. Most of these flavonoids have a conversion rate of more than 60%, and their glycosylation usually occur at the 30 , 40 , and 7 position. Furthermore, raw materials such as soybeans extract can also be utilized as the acceptor substrate of ASase (Jung et al. 2020b). The optimal glycosylation conditions are usually mild, pH ranges from 5.0 to 8.0, and temperatures range from 30  C to 45  C. Rha et al. (2019a) investigated the effect of pH on transglycosylation products and found that a low pH is good for the gross yield, while a relatively high pH appears to be good for obtaining single product. In addition to the optimization of reaction conditions, molecular modification and food-grade expression have also been attempted to improve the transglycosylation activity of ASase toward flavonoids (Malbert et al. 2014; Kim et al. 2016b, 2018; Chin et al. 2020). For example, Kim et al. (2016b) reported that a variant of DRp-ASase, Q299K, showed a tenfold improvement in rutin glycosylation ability.

11.4.3.2

Glycosylation of Other Bioactive Compounds

In addition to flavonoids, ASases have also been widely used in the glycosylation of other compounds, including polyols, phenolic compounds, polyphenolic compounds, polyhydroxy compounds, and glycoside molecules. The advantage of the glycosylation reaction catalyzed by ASase lies mainly in its strong enantioselectivity, as well as the high production yields, and the utilization of cheap donor substrate (Moulis et al. 2016). Several bioactive compounds have been glycosylated using ASase, including β-glucosylglycerol (Jung et al. 2019a), aesculin, aesculetin (Park et al. 2018b), caffeic acid phenethyl ester (Moon et al. 2017), glycerol (Jeong et al. 2014), D-arabitol, maltitol, D-mannitol, myoinositol, D-sorbitol, xylitol (Daudé et al. 2013a), piceid (Park et al. 2012), and salicin (Jung et al. 2009). The additional glycosides can protect aglycone from oxidative degradation, thereby extending its half-life in cells and allowing it to exert beneficial biological properties. For example, the solubility of caffeic acid phenethyl ester-4-O-α-D-glucopyranoside synthesized by DG-ASase is increased by 770 times, the half-life is increased by 10 times, and the bioavailability in RAW 264.7 cells is also improved (Moon et al. 2017). ASase can be applied to the feasible synthesis of some bioactive compounds, such as α-arbutin (Zhu et al. 2019). α-Arbutin (4-hydroxyphenyl α-Dglucopyranoside), an unnatural compound, has strong inhibitory effect on the tyrosinase activity and thus can be used as skin lightening agent in cosmeceutical industry (Zhu et al. 2018c). Until now, XC-ASase (Yang et al. 2019), CC-ASase (Yu et al. 2018), and DG-ASase (Seo et al. 2012b) have been used to produce α-arbutin from hydroquinone. CC-ASase has been reported to have high transglycosylation activity and thus can efficiently catalyze the synthesis of

Flavanone

Isoflavone

Type Flavone

DG-ASase DG-ASase DG-ASase NP-ASase DG-ASase

40 , 7

40 , 7

40 , 7, 8

7, 8

40 / 7-O-glucoside

Daidzein

Glycitein

8-Hydroxydaidzein

6

4’

6-Hydroxyflavanone

4’-Hydroxyflavanone

7,8-diHydroxy40 -methylisoflavone Daidzin

DG-ASase

DG-ASase

DG-ASase

Genistein

DG-ASase DG-ASase

6, 7 30 , 40 , 5, 7/ 6-Cglucoside 40 , 5, 7

DG-ASase

40 , 5, 7

Apigenin

6,7-Dihydroxyflavone Homoorientin

DG-ASase

5, 6, 7

Baicalein

Enzyme DG-ASase DG-ASase

Hydroxyl/Glycosyl location 30 , 40 , 5, 7 6

Acceptor Luteolin 6-Hydroxyflavone

G, G2

G2

G, G2

7-G

7-G

7-G

NR

NR

NR NR

G2

G2

Main product 40 -G G2

Table 11.3 The summary of flavonoids glycosylation reactions catalyzed by ASases

5.0

5.0

8.0

7.0

7.0

5.0

5.0

5.0

8.0 8.0

24 h

45  C

24 h 24 h 24 h

45  C 45  C

24 h

30  C 45  C

0.5 h

24 h 40  C

45  C

24 h

24 h 24 h

37  C 37  C

45  C

24 h

45  C

24 h

45  C

5.0 5.0

Time 24 h 24 h

Temperature 37  C 45  C

pH 7.0 5.0

60a,b

100a,b

99

64

89.3

88.8a

96.9a

98.2a

56 57

60a,b

100a,b

Yield/ % 86 100a,b

Jung et al. (2020b) Jung et al. (2020b) Jung et al. (2020b) Chang et al. (2019) Overwin et al. (2016) Kim et al. (2016b) Rha et al. (2019a) Rha et al. (2019a)

Reference Jang et al. (2018) Rha et al. (2019a) Rha et al. (2019a) Rha et al. (2019a) Jang et al. (2018) Jang et al. (2018)

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DRp-ASase/ Q229K DG-ASase NP-ASase NP-ASase

4, 40 , 6’

DG-ASase

DG-ASase

30 , 40 , 5, 7/ 3-Oglucoside 30 , 40 , 5, 7/ 3-Orutinoside 3, 30 , 40 , 5, 7 3, 30 , 40 , 5, 7

3, 6

Yield of all products Estimating from the chart. NR, not reported

Phloretin

Chalcone

b

a

Catechin Epicatechin

Rutin

Isoquercitrin

3,6-diHydroxyflavone

DG-ASase

40 , 5, 7

Naringenin

Flavanol

Flavonol

DG-ASase

40 , 7

Liquiritigenin

(n ¼ 1 ~ 3)

30 -G,
G2 40 -G, 30 -G2,
G3 40 -Gn

G3, 400 -G,
G2 400 ‘-G

G2

G2

G2

7.0

8.0 7.0

8.0

7.0

5.0

5.0

5.0

30  C

8h

1h 24 h

16 h

40  C 30  C 30  C

24 h

24 h

45  C 45  C

24 h

24 h

45  C

45  C

95a

97a 81a

NR

97.6

100a,b

60a,b

60a,b Rha et al. (2019a) Rha et al. (2019a) Rha et al. (2019a) Rha et al. (2019c) Kim et al. (2016b) Cho et al. (2011) Overwin et al. (2015b) Overwin et al. (2015a)

11 Amylosucrase: A Versatile Sucrose-Utilizing Transglucosylase for. . . 241

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α-arbutin in vitro with no need to add ascorbic acid to protect the substrate from oxidation (Yu et al. 2018). Zhu et al. (2019) evaluated scale-up production in 5000 L reactor using whole cell of XC-ASase expressed in E. coli and achieved conversion rates of about 95% and productivities of about 4.9 kg kL
1 h
1. In addition, Hun et al. (2018) has investigated the immobilization of the DG-ASase on Amicogen LKZ118 beads, and the results showed that its transglycoslation activity toward hydroquinone can maintain 35 cycles.

11.5

Conclusion and Perspectives

In recent years, ASase has been attracted widespread attention as a sucrose-utilizing transglucosylase for glycodiversification. Attempts have been made to discover ASases from novel microorganisms and express them heterogeneously in foodgrade strains. Structural analysis was also carried out to understand the catalytic mechanism of the enzyme, following by the molecular modification to improve its properties, and thereby broadening its application range. However, the poor thermal stability and the lack of desirable specificity still hamper their applications to some extent. Therefore, computer-aided design is underway to obtain ASase variants with requisite specificity and improved thermostability. It is believed that rapid development of enzyme engineering technology will help solve these problems of ASase.

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Seo DH, Jung JH, Ha SJ, Cho HK, Jung DH, Kim TJ, Baek NI, Yoo SH, Park CS (2012b) Highyield enzymatic bioconversion of hydroquinone to alpha-arbutin, a powerful skin lightening agent, by amylosucrase. Appl Microbiol Biot 94(5):1189–1197. https://doi.org/10.1007/ s00253-012-3905-7 Seo D-H, Jung J-H, Park C-S (2019) Improved polymerization activity of Deinococcus geothermalis amylosucrase by semi-rational design: effect of loop flexibility on the polymerization reaction. Int J Biol Macromol 130:177–185. https://doi.org/10.1016/j.ijbiomac.2019.02. 139 Shin HJ, Choi SJ, Park CS, Moon TW (2010) Preparation of starches with low glycaemic response using amylosucrase and their physicochemical properties. Carbohyd Polym 82(2):489–497. https://doi.org/10.1016/j.carbpol.2010.05.017 Skov LK, Mirza O, Henriksen A, De Montalk GP, Remaud-Simeon M, Sarcabal P, Willemot RM, Monsan P, Gajhede M (2000) Crystallization and preliminary X-ray studies of recombinant amylosucrase from Neisseria polysaccharea. Acta Crystallogr D 56:203–205. https://doi.org/10. 1107/S0907444999015887 Skov LK, Mirza O, Henriksen A, De Montalk GP, Remaud-Simeon M, Sarcabal P, Willemot RM, Monsan P, Gajhede M (2001) Amylosucrase, a glucan-synthesizing enzyme from the alphaamylase family. J Biol Chem 276(27):25273–25278. https://doi.org/10.1074/jbc.M010998200 Skov LK, Mirza O, Sprogoe D, Dar I, Remaud-Simeon M, Albenne C, Monsan P, Gajhede M (2002) Oligosaccharide and sucrose complexes of amylosucrase - structural implications for the polymerase activity. J Biol Chem 277(49):47741–47747. https://doi.org/10.1074/jbc. M207860200 Skov LK, Mirza O, Sprogøe D, van der Veen BA, Remaud-Simeon M, Albenne C, Monsan P, Gajhede M (2006) Crystal structure of the Glu328Gln mutant of Neisseria polysaccharea amylosucrase in complex with sucrose and maltoheptaose. Biocatal Biotransformation 24 (1–2):99–105. https://doi.org/10.1080/10242420500538100 Skov LK, Pizzut-Serin S, Remaud-Simeon M, Ernst HA, Gajhede M, Mirza O (2013) The structure of amylosucrase from Deinococcus radiodurans has an unusual open active-site topology. Acta Crystallograp Sect F Struct Biol Crystallization Commun 69:973–978. https://doi.org/10.1107/ s1744309113021714 Slámová K, Kapešová J, Valentová K (2018) “Sweet flavonoids”: glycosidase-catalyzed modifications. Int J Mol Sci 19(7):2126. https://doi.org/10.3390/ijms19072126 Stam MR, Danchin EGJ, Rancurel C, Coutinho PM, Henrissat B (2006) Dividing the large glycoside hydrolase family 13 into subfamilies: towards improved functional annotations of -amylase-related proteins. Protein Eng Des Sel 19(12):555–562. https://doi.org/10.1093/ protein/gzl044 Su L, Zhao Y, Wu D, Wu J (2020) Heterogeneous expression, molecular modification of amylosucrase from Neisseria polysaccharea, and its application in the preparation of turanose. Food Chem 314:126212. https://doi.org/10.1016/j.foodchem.2020.126212 Tian Y, Xu W, Zhang W, Zhang T, Guang C, Mu W (2018) Amylosucrase as a transglucosylation tool: from molecular features to bioengineering applications. Biotechnol Adv 36:1540–1552. https://doi.org/10.1016/j.biotechadv.2018.06.010 Tian Y, Deng Y, Zhang W, Mu W (2019a) Sucrose isomers as alternative sweeteners: properties, production, and applications. Appl Microbiol Biot 103(Suppl 12):1–11 Tian Y, Xu W, Guang C, Zhang W, Mu W (2019b) Thermostable amylosucrase from Calidithermus timidus DSM 17022: insight into its characteristics and tetrameric conformation. J Agr Food Chem 67(35):9868–9876. https://doi.org/10.1021/acs.jafc.9b04023 van der Veen BA, Potocki-Veronese G, Albenne C, Joucla G, Monsan P, Remaud-Simeon M (2004) Combinatorial engineering to enhance amylosucrase performance: construction, selection, and screening of variant libraries for increased activity. FEBS Lett 560(1–3):91–97. https:// doi.org/10.1016/S0014-5793(04)00077-8 van der Veen BA, Skov LK, Potocki-Veronese G, Gajhede M, Monsan P, Remaud-Simeon M (2006) Increased amylosucrase activity and specificity, and identification of regions important

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for activity, specificity and stability through molecular evolution. FEBS J 273(4):673–681. https://doi.org/10.1111/j.1742-4658.2005.05076.x Vergès A, Cambon E, Barbe S, Salamone S, Le Guen Y, Moulis C, Mulard LA, RemaudSiméon M, André I (2015) Computer-aided engineering of a transglycosylase for the glucosylation of an unnatural disaccharide of relevance for bacterial antigen synthesis. ACS Catal 5(2):1186–1198. https://doi.org/10.1021/cs501288r Vergès A, Barbe S, Cambon E, Moulis C, Tranier S, Remaud-Siméon M, André I (2017a) Engineering of anp efficient mutant of Neisseria polysaccharea amylosucrase for the synthesis of controlled size maltooligosaccharides. Carbohyd Polym 173:403–411. https://doi.org/10. 1016/j.carbpol.2017.06.011 Vergès A, Cambon E, Barbe S, Moulis C, Remaud-Siméon M, André I (2017b) Novel product specificity toward erlose and panose exhibited by multisite engineered mutants of amylosucrase. Protein Sci 26(3):566–577. https://doi.org/10.1002/pro.3106 Wang R, Bae JS, Kim JH, Kim BS, Yoon SH, Park CS, Yoo SH (2012) Development of an efficient bioprocess for turanose production by sucrose isomerisation reaction of amylosucrase. Food Chem 132(2):773–779. https://doi.org/10.1016/j.foodchem.2011.11.035 Wang Y, Xu W, Bai Y, Zhang T, Jiang B, Mu W (2017) Identification of an alpha-(1,4)-glucansynthesizing amylosucrase from Cellulomonas carboniz T26. J Agric Food Chem 65 (10):2110–2119. https://doi.org/10.1021/acs.jafc.6b05667 Xiao J, Muzashvili TS, Georgiev MI (2014) Advances in the biotechnological glycosylation of valuable flavonoids. Biotechnol Adv 32(6):1145–1156. https://doi.org/10.1016/j.biotechadv. 2014.04.006 Yang C, Fan W, Zhang R, Shi J, Knežević-Jugović Z, Zhang B (2019) Study on transglucosylation properties of amylosucrase from Xanthomonas campestris pv. Campestris and its application in the production of α-arbutin. Catalysts 9(1):5. https://doi.org/10.3390/catal9010005 Yu S, Wang Y, Tian Y, Xu W, Bai Y, Zhang T, Mu W (2018) Highly efficient biosynthesis of α-arbutin from hydroquinone by an amylosucrase from Cellulomonas carboniz. Process Biochem 68:93–99. https://doi.org/10.1016/j.procbio.2018.02.012 Zhang H, Chen Z, Zhou X, He J, Wang T, Luo X, Wang L, Wang R (2018) Anti-digestion properties of amylosucrase modified waxy corn starch. Int J Biol Macromol 109:383–388 Zhang H, Wang R, Chen Z, Zhong Q (2019) Enzymatically modified starch with low digestibility produced from amylopectin by sequential amylosucrase and pullulanase treatments. Food Hydrocoll 95:195–202. https://doi.org/10.1016/j.foodhyd.2019.04.036 Zhang H, Wang R, Chen Z, Zhong Q (2020) Amylopectin-sodium palmitate complexes as sustainable nanohydrogels with tunable size and fractal dimensions. J Agr Food Chem 68 (12):3796–3805. https://doi.org/10.1021/acs.jafc.9b06248 Zhu X, Tian Y, Xu W, Bai Y, Zhang T, Mu W (2018a) Biochemical characterization of a highly thermostable amylosucrase from Truepera radiovictrix DSM 17093. Int J Biol Macromol 116:744–752. https://doi.org/10.1016/j.ijbiomac.2018.05.096 Zhu X, Tian Y, Xu W, Guang C, Zhang W, Zhang T, Mu W (2018b) Bioconversion of sucrose to maltooligosaccharides by the synergistic action of amylosucrase and α-amylase. Process Biochem 74:71–76. https://doi.org/10.1016/j.procbio.2018.08.026 Zhu X, Tian Y, Zhang W, Zhang T, Guang C, Mu W (2018c) Recent progress on biological production of α-arbutin. Appl Microbiol Biot 102(19):8145–8152. https://doi.org/10.1007/ s00253-018-9241-9 Zhu L, Xu M, Lu C, Chen L, Xu A, Fang J, Chen H, Lu Y, Fan Y, Chen X (2019) Optimization of whole-cell biotransformation for scale-up production of α-arbutin from hydroquinone by the use of recombinant Escherichia coli. AMB Express 9(1). https://doi.org/10.1186/s13568-019-08207

Chapter 12

Glucansucrases Derived from Lactic Acid Bacteria to Synthesize Multitudinous α-Glucans Ziwei Chen, Dawei Ni, and Wanmeng Mu

12.1

Introduction

Lactic acid bacteria (LAB) are widely applied in the industrial production of fermented foods, such as pickles, yoghurt, and soy sauce, due to their accredited “GRAS” (generally regarded as safe) status and also well known to produce exopolysaccharides (EPS) (Patel and Prajapat 2013). α-Glucan is one kind of exopolysaccharides, consisting of various numbers of α-linked glucosyl units. The α-glucans derived from various LAB strains display a variety of linkage compositions, branching degrees, and sizes. Their linkage types include (α1 ! 6), (α1 ! 3), (α1 ! 4), and/or (α1 ! 2) in main chain or branching orientation (Leemhuis et al. 2013; Meng et al. 2016b). These α-glucans can be divided into four categories: dextran, mutan, alternan, and reuteran, on the basis of their main linkage types and structural organizations (Fig. 12.1). Dextran is a water-soluble α-glucan constituted by D-glucose residues with more than 50% of consecutive (α1 ! 6) linkages in the main chain and a small amount of (α1 ! 4), (α1 ! 3) and/or (α1 ! 2) linkages in branching orientations. In contrast, mutan is a water-insoluble α-glucan because it predominantly contains (α1 ! 3) linkages in the linear main chain. The watersoluble reuteran is composed of (α1 ! 4) linear fragments with various sizes interconnected by single (α1 ! 6) bridges. Particularly, alternan has a unique main chain with alternative (α1 ! 3) and (α1 ! 6) linkages and thus shows a considerable water solubility. Their structural diversity makes that they show miscellaneous physico-chemical properties and an enormous application potential in Z. Chen · D. Ni State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China W. Mu (*) State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_12

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Fig. 12.1 Typical structures of α-glucans synthesized by various glucansucrases

different fields, including food, cosmetic, medicine, biotechnology, and environmental governance. Dextran and its derivatives have been extensively exploited as blood plasma expander, anticoagulant, lubricant, drug carrier, thickener, stabilizer, texturizer, emulsifier, gels, flocculants, and chromatography matrices (Naessens et al. 2005; Bhavani and Nisha 2010; Zannini et al. 2016; Heinze et al. 2006). In 2013, dextran was consented to be used as an ingredient added directly into foodstuffs (21CFR186.1275) with a “generally regarded as safe” (GRAS) status by the US Food and Drug Administration (FDA) (Zannini et al. 2016). Additionally, the low molecular weight dextran70 (70 kDa) is referenced in the 19th list of essential medicines by the World Health Organization (Claverie et al. 2017). Reuteran can be potentially used as a food ingredient to prompt health, a dietary fiber to bring about satiety in humans, and a food texturizer to improve bread mouthfeel (Xiao et al. 2016; Ekhart et al. 2006). Alternan has been described as a low-calorie food additive due to its anti-digestible property and as a texturizer to substitute oil or fat in cosmetic productions (Frohberg and Pilling 2019). However, mutan polysaccharide detrimentally facilitates the adhesion of oral flora microorganisms on the tooth surface due to its water-insolubility, resulting in plaque formation and dental crisis (Fernandes et al. 2018; Pleszczynska et al. 2015).

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Glucansucrases (GSs), also known as glucosyltransferases (Gtfs), are classified into glycoside hydrolase 70 family (GH70) according to the CAZy classification system (http://www.cazy.org/). GS enzymes can split sucrose and polymerize the D-glucose moieties to synthesize α-glucan. GSs are mainly divided into dextransucrase (EC 2.4.1.5), alternansucrase (EC 2.4.1.-), reuteransucrase (EC 2.4.1.140), and mutansucrase (EC 2.4.1.125) based on their synthesized products. Among of them, dextransucrase is the most widely studied GS enzyme and has been applied in different fields. Compared to Leloir glycosyltransferase enzymes, GSs are lower-cost tools used for the biotechnological production of α-glucan polysaccharides from sucrose, without the requirement of expensive nucleotideactivated sugars (e.g., UDP-glucose). To date, GSs are exclusively found to exist in LAB genera, including Lactobacillus, Streptococcus, Weissella, Leuconostoc, and Oenococcus (Gangoiti et al. 2018). Most of them can only produce one GS enzyme, but some of them can produce multiple different GS enzymes. For example, the LAB strain Leuconostoc citreum NRRL B-1299 (previously also known as Leuconostoc mesenteroides NRRL B-1299) can encode six GSs, including DSR-A, DSR-B, DSR-M, DSR-DP, DSR-E, and BRS-A, which display different product specificities (Passerini et al. 2015). Additionally, the pathogenic Streptococcus mutans can also produce three different GSs, namely, GtfB, GtfC, and GtfD (Ooshima et al. 2001; Hanada and Kuramitsu 1989). GSs are high molecular weight (approximately ranging from 120 to 200 kDa) extracellular enzymes, which significantly hinders their expression, purification, characterization, and crystallization. According to the CAZy database, although 751 GS enzymes have been sequenced, only 58 of them have been identified (by Aug 2020). In the past few decades, researchers have put a lot of efforts to obtain the GS crystal structures, but until 2010, the first crystal structure of GS enzyme, dextransucrase Gtf180-ΔN from Lactobacillus reuteri 180, has been solved by a N-terminal truncation method (Vujičić-Žagar et al. 2010). Subsequently, other types of GS enzymes, including reuteransucrase GtfAΔN from Lactobacillus reuteri 121 (Pijning et al. 2012), mutansucrase Gtf-SI from Streptococcus mutans (Ito et al. 2011), branching sucrase DSR-E Δ123-GBD-CD2 from L. citreum NRRL B-1299 (Brison et al. 2012b), dextransucrase DSR-MΔ2 from L. citreum NRRL B-1299 (Claverie et al. 2017), and alternansucrase ASRΔ2 from Leuconostoc citreum NRRL B-1355 (Molina et al. 2019), were also successfully determined with various truncation forms in variable region but retaining full activity. Although these solved crystal structures deepen our understanding on the structure-function relationship of GS enzymes, the synthesis mechanism of α-glucan, especially in product specificity (including linkage specificity and branching specificity), has not been thoroughly elucidated yet. This chapter mainly focuses on the characterization, 3D structure, catalytic mechanism, and product specificity of glucansucrase.

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Characterization of Glucansucrases

12.2.1 Dextransucrase In 1861, Pasteur reported that a slime substance derived from unknown microorganisms resulted in the mysterious thickening of beet and cane sugar juices (Pasteur 1861). In 1874, Scheibler determined that this slime was a carbohydrate with an empirical formula (C6H10O6)n and a positive rotatory power and thus named it as “dextran” (Scheibler 1874). Later, this dextran-producing bacterium and corresponding extracellular enzyme were named Leuconostoc mesenteroides and dextransucrase, respectively (Van Tieghem 1878; Hestrin et al. 1943). Nowadays, the dextransucrase DSR-S from Leuconostoc mesenteroides NRR B-512F has been industrially used to produce a high molecular weight dextran (up to 1 MDa) composed of 95% (α1 ! 6) and 5% (α1 ! 3) linkages in the main chains and branching directions, respectively, which has been commercialized in many developed countries around the world (Gangoiti et al. 2018; Monchois et al. 1997). Dextransucrase has been extensively characterized from others LAB stains, including Lactobacillus, Streptococcus, Weissella, and Oenococcus genera. Another extensively studied dextransucrase Gtf180 from L. reuteri 180 also produce a high molecular weight dextran (36 MDa) composed of 31% (α1 ! 3) and 69% (α1 ! 6) linkages (Kralj et al. 2004a; van Leeuwen et al. 2008b). Two dextransucrases DSR-A and DSR-B from L. citreum NRRL B-1299 are characterized to produce two dextrans both composed of a high proportion of (α1 ! 6) linkages (85% and 95%, respectively) and a small amount of (α1 ! 3) linkages (15% and 5%, respectively) (Monchois et al. 1996, 1998a). Particularly, the other two dextransucrases DSR-M and DSR-DP from L. citreum NRRL B-1299 both produce the dextrans, exclusively composed of (α1 ! 6) linkages (Passerini et al. 2015). Similar situations have also been found in other LAB strains. Three dextransucrases, including GtfK from Streptococcus salivarius ATCC 25975, DSRWC from Weissella cibaria CMU, and DSRBCB4 from L. mesenteroides NRRL B-1299 CB4, can also produce the peculiar dextrans with absolute (α1 ! 6) linkages (Kang et al. 2008, 2009; Simpson et al. 1995). Notably, DSR-M is the first characterized dextransucrase to naturally produce a low molecular weight dextran (27 kDa) from sucrose. Subsequently, the dextran synthesis is verified to adopt a nonprocessive polymerization mechanism (Claverie et al. 2017). On the contrary, DSR-OK from Oenococcus kitaharae DSM 17330 is the dextransucrase that is biochemically characterized to produce the largest dextran to date, with an ultrahigh molecular weight (up to 1100 MDa) containing 98% (α1 ! 6) and 2% (α1 ! 3) linkages (Vuillemin et al. 2018). Recently, the dextransucrase Gtf-DSM from Lactobacillus ingluviei DSM 14792 has been characterized to have a unique product specificity, producing a dextran composed of 1% (α1 ! 2), 6% (α1 ! 3), 24% (α1 ! 4), and 69% (α1 ! 6) linkages, which is the first α-glucan containing four known linkages and directly synthesized from sucrose by a GS enzyme having only one independent catalytic domain (Chen et al. 2019). In

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addition, other characterized dextransucrases, including DSR-F from Leuconostoc citreum B/110-1-2 (Fraga et al. 2011), GtfB-2 from Leuconostoc citreum B-2 (Feng et al. 2018), DSRC39-2 from Weissella confusa 39-2 (Amari et al. 2013), GtfR from Streptococcus oralis ATCC10557 (Hendrik et al. 2008), GtfM from S. salivarius ATCC 25975 (Simpson et al. 1995), Gtf-S form Streptococcus downei Mfe 28 (Gilmore et al. 1990), GtfD from Streptococcus mutans GS5 (Hanada and Kuramitsu 1989), GtfKg3 from Lactobacillus fermentum Kg3, Gtf33 from Lactobacillus parabuchneri 33, and GtfKg15 from Lactobacillus sakei Kg15 (Kralj et al. 2004a), produce the dextrans containing a high percentage ( 70%) of (α1 ! 6) linkages (Table 12.1).

12.2.2 Mutansucrase Mutansucrases are mainly derived from pathogenic Streptococcus strains and also rarely found in Lactobacillus and Leuconostoc genera (Kralj et al. 2004a; Côté and Skory 2012). To date, all characterized mutansucrases synthesize the mutans only containing (α1 ! 3) and (α1 ! 6) linkages (Table 12.1). Two mutansucrases from S. mutans GS5, namely, GtfB and GtfC, are characterized to synthesize waterinsoluble mutans from sucrose and display similar linkage specificity (Tamesada et al. 2004). GtfB and GtfC can produce the mutans containing 88% and 85% (α1 ! 3) and 12% and 15% (α1 ! 6) linkages, respectively (Shiroza et al. 1987; Ueda et al. 1988). In contrast, two mutansucrases GtfJ and GtfL from S. salivarius ATCC 25975 show different product specificity. GtfJ produces a mutan composed of 90% (α1 ! 3) and 10% (α1 ! 6) linkages, but GtfL produces a mutan composed of equal (α1 ! 3) and (α1 ! 6) linkages (Simpson et al. 1995). The mutansucrase DSR-I from Leuconostoc mesenteroides NRRL B-1118 displays the same linkage specificity as GtfL, which also produces a mutan containing approximately equal amounts of (α1 ! 3) and (α1 ! 6) linkages (Côté and Skory 2012). The mutansucrase GtfML1 from Lactobacillus reuteri ML1 synthesizes a high molecular weight mutan (8 MDa) comprising 65% (α1 ! 3) and 35% (α1 ! 6) linkages (Kralj et al. 2004a). The mutansucrase Gtf-I from S. downei Mfe 28 produces a mutan containing 94% (α1 ! 3) and 6% (α1 ! 6) linkages (Monchois et al. 1999).

12.2.3 Reuteransucrase To date, only three reuteransucrases have been biochemically characterized and both of them are produced by L. reuteri strains (Table 12.1). The reuteransucrase GtfA from probiotic strain L. reuteri 121 has been studied most and synthesizes a high molecular weight reuteran (40 MDa) with 58% (α1 ! 4) and 42% (α1 ! 6) linkages (Kralj et al. 2004b; Meng et al. 2016b). Structural analysis of reuteran derived from GtfA indicated that it has a heterogeneous structure without repeat modules and it is

Glucansucrase Dextransucrase

GtfKg15 GtfKg3 Gtf33 DSR-F

GtfM

DSRWC GtfK

DSRBCB4

DSR-OK

DSR-M

DSR-DP

DSR-B

Gtf180 DSR-A

Name DSR-S

LAB strain L. mesenteroides NRRL B-512F L. reuteri 180 L. citreum NRRL B-1299 L. citreum NRRL B-1299 L. citreum NRRL B-1299 L. citreum NRRL B-1299 O. kitaharae DSM 17330 L. mesenteroides B-1299 CB4 W. cibaria CMU S. salivarius ATCC 25975 S. salivarius ATCC 25975 L. sakei Kg15 L. fermentum Kg3 L. parabuchneri 33 L. citreum B/110–1-2 98

1.1  109

95 90 92 81 93

2.7  107 2.4  107 0.2  106 2  106

100 100

NR

NR NR

100

100

2.7  104

NR

100

95

69 85

1

10 8 19 6

5

2

5

31 15

Linkage composition (%) (α1 ! 6) (α1 ! 4) (α1 ! 3) 95 5

2  106

NR

3.6  107 NR

Molecular weight (Mw, Da) 1  106

Table 12.1 Characterized GS enzymes from various LAB strains (α1 ! 2)

Kralj et al. (2004a) Kralj et al. (2004a) Kralj et al. (2004a) Fraga et al. (2011)

Simpson et al. (1995)

Kang et al. (2009) Simpson et al. (1995)

Kang et al. (2008)

Vuillemin et al. (2018)

Passerini et al. (2015)

Monchois et al. (1998a) Passerini et al. (2015)

Kralj et al. (2004a) Monchois et al. (1996)

Reference Monchois et al. (1997)

256 Z. Chen et al.

Alternansucrase

Mutansucrase

Reuteransucrase

S. oralis ATCC10557 S. mutans GS5

S. downei Mfe 28 W. confusa 39–2 L. citreum B-2 L. reuteri 121 L. reuteri ATCC 55730 L. reuteri SK24.003 L. reuteri ML1 L. mesenteroides NRRL B-1118 S. mutans GS5 S. mutans GS5 S. downei Mfe 28 S. salivarius ATCC 25975 S. salivarius ATCC 25975 L. mesenteroides NRRL B-1355 L. citreum SK24.002 Leuconostoc citreum ABK-1

GtfR GtfD

Gtf-S DSRC39–2 GtfB-2 GtfA GtfO Gtf-SK3 GtfML1 DSRI

GtfB-SK2 LcALT

ASR

GtfL

GtfB GtfC Gtf-I GtfJ

L. ingluviei DSM 14792

Gtf-DSM

69

50 56 58 60

2  106 4.6  107 NR

12 15 6 10

90 97 75 42 21 20 35 50

86 70

NR

NR NR NR NR

NR 2  106 3.8  106 4  107 4.2  107 4.3  107 8  106 NR

NR NR

5.5  107, 5.3  105

58 79 80

6

24

42 40

44

50

88 85 94 90

65 50

10 3 19

14 30

6

1

Chen et al. (2019)

(continued)

Côté and Robyt (1982) Song et al. (2016) Wangpaiboon et al. (2018)

Simpson et al. (1995)

Shiroza et al. (1987) Ueda et al. (1988) Monchois et al. (1999) Simpson et al. (1995)

Hendrik et al. (2008) Hanada and Kuramitsu (1989) Gilmore et al. (1990) Amari et al. (2013) Feng et al. (2018) Kralj et al. (2004b) Kralj et al. (2005a) Miao et al. (2014b) Kralj et al. (2004a) Côté and Skory (2012)

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NR Not reported

Glucansucrase Branching sucrase

BRS-B GtfZ-CD2

BRS-A

Name DSR-E

Table 12.1 (continued)

LAB strain L. citreum NRRL B-1299 L. citreum NRRL B-1299 L. citreum NRRL B-742 L. kunkeei DSM 12361 NR NR

NR

Molecular weight (Mw, Da) NR

60

50 40

Linkage composition (%) (α1 ! 6) (α1 ! 4) (α1 ! 3) 81 3 10 34

(α1 ! 2) 5

Vuillemin et al. (2016) Meng et al. (2018)

Passerini et al. (2015)

Reference Fabre et al. (2005)

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built of maltose, maltotriose, and maltotetraose residues connected with single (α1 ! 6) bridges (van Leeuwen et al. 2008a). The reuteransucrase GtfO from Lactobacillus reuteri ATCC 55730 is a highly hydrolytic GS enzyme and synthesizes a high molecular weight reuteran (42 MDa) with large amounts of (α1 ! 4) linkages (79%) and minor (α1 ! 6) linkages (21%) (Kralj et al. 2005a). The reuteransucrase Gtf-SK3 from Lactobacillus reuteri SK24.003 which is isolated from a traditional Chinese fermented dairy product shows a product specificity similar to that of GtfO, which also synthesizes a high molecular weight reuteran (43 MDa) with 80% (α1 ! 4) and 20% (α1 ! 6) linkages (Miao et al. 2014b; Ming et al. 2015).

12.2.4 Alternansucrase The alternansucrase ASR from L. mesenteroides NRRL B-1355 is characterized to produce a peculiar α-glucan with alternate (α1 ! 3) and (α1 ! 6) linkages (Table 12.1), and thus this α-glucan is referred to alternan (Côté and Robyt 1982). Biochemical characterization showed that the alternan produced by ASR is composed of 44% (α1 ! 3) and 56% (α1 ! 6) linkages and has a high water solubility and strong resistance to enzymatic degradation, due to its alternating structural characteristic (Joucla et al. 2006). The alternansucrase Gtf-SK2 from Leuconostoc citreum SK24.002 which is isolated from Chinese traditional pickled vegetables synthesizes a high molecular weight alternan (46 MDa) with 42% (α1 ! 3) and 58% (α1 ! 6) linkages (Miao et al. 2014a; Song et al. 2016). Recently, a novel alternansucrase LcALT from Leuconostoc citreum ABK-1 has been biochemically characterized to produce an alternan-like α-glucan with 40% (α1 ! 3) and 60% (α1 ! 6) linkages interconnected in an irregular alternating form (Wangpaiboon et al. 2018).

12.2.5 Branching Sucrases Branching sucrases are atypical glucansucrases of GH70 family and different from typical glucansucrases in substrate specificity. Using sucrose as single substrate, branching sucrases only catalyze sucrose hydrolysis and cannot polymerize the released D-glucose to synthesize α-glucan polysaccharides. However, when supplementing additional dextran as acceptor substrate, branching sucrases can graft (α1 ! 3) or (α1 ! 2) linkages in the branching orientations of dextran acceptor, synthesizing highly branched dextran products (Moulis et al. 2016). DSR-E from L. citreum NRRL B-1299 is a peculiar GS enzyme and contains two full active catalytic domains (DSR-E-CD1 and DSR-E-CD1) insulated by an intermediate glucan-binding domain (GBD) (Bozonnet et al. 2002). Native DSR-E can produce a dextran with 5% (α1 ! 2), 3% (α1 ! 3), 10% (α1 ! 4), and 81%

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(α1 ! 6) linkages from sucrose (Fabre et al. 2005). Truncation experiment and biochemical characterization indicated that both CD1 and CD2 of DSR-E display high regiospecificity. DSR-E-CD1 is characterized to be a dextransucrase and converts sucrose to the dextran long chain with dominant (α1 ! 6) linkages. Nevertheless, DSR-E-CD2 stems (α1 ! 6) main chain synthesized by DSR-ECD1 with (α1 ! 2) linkages, producing highly branched dextran, and thus is classified as a branching sucrase (Brison et al. 2012a, 2016). The branching sucrase BRS-A from L. citreum NRRL B-1299 displays a high sequence identity with DSR-E-CD2 and introduces 37% (α1 ! 2) linkages in the branching orientations of dextran acceptor (Passerini et al. 2015). Recently, a novel GS enzyme GtfZ from Lactobacillus kunkeei DSM 12361 similar to DSR-E has been also characterized to harbor two catalytic domains, namely, GtfZ-CD1 and GtfZ-CD2, interconnected by a GBD (Meng et al. 2018). However, biochemical studies showed that branching sucrase GtfZ-CD2 synthesizes (α1 ! 3)-linked branches onto the linear dextran chain, forming highly branched comb-like dextran (Meng et al. 2018). The branching sucrase BRS-B from Leuconostoc citreum NRRL B-742 catalyzes the formation of 50% (α1 ! 3) branching linkages on the dextran main chain with an appropriate ratio of dextran acceptor and sucrose (Vuillemin et al. 2016). Additionally, the other two branching sucrases BRS-D from Lactobacillus kunkeei EFB6 and BRS-C from Leuconostoc fallax KCTC3537 also have been characterized to synthesize (α1 ! 2) and (α1 ! 3) branching linkages, respectively (Vuillemin et al. 2016). Overall, these emerging branching sucrase promise a great application potential to synthesize tailor-made α-glucans.

12.3

Structures of Glucansucrases

12.3.1 Structural and Functional Organization Before the available crystal structures, the structural organization of GSs was mainly deduced by the sequence alignments with GH13 enzymes, which are evolutionarily relevant to GH70 family enzymes. Their primary sequence alignments indicated that full-length GSs contain four regions, including signal peptide (SP), N-terminal variable region (VR), catalytic domain (CD), and C-terminal glucan-binding domain (GBD) (Moulis et al. 2006; Monchois et al. 1998b). Until 2010, the crystal structure of Gtf180-ΔN from L. reuteri 180 was first solved by truncating its N-terminal variable region (Vujičić-Žagar et al. 2010). Later on, other crystal structures of various truncated GSs, including GtfA-ΔN, Gtf-SI, DSR-E ΔN123-GBD-CD2, ASRΔ2, and DSR-MΔ2, have been determined in succession. Unexpectedly, their three-dimensional structures showed that GSs employ a novel structural organization different from the previously predicted structures according to amino acid sequence alignments. The polypeptide chains of resolved GSs pose a U-shape route harboring five structural domains called after A, B, C, IV, and V (Vujičić-Žagar et al. 2010). In terms of A, B, IV, and V domains, each of them is composed

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Fig. 12.2 Crystal structures and domain organization of representative GH13 amylase from Bacillus licheniformis (a; PDB: 1BLI) and GH70 dextransucrase Gtf180-ΔN from L. reuteri 180 (b; PDB: 3KLK)

of two discrete polypeptides chains located in N- and C-termini, respectively. However, domain C is a consecutive polypeptides chain and connects the N- and C-terminal polypeptides chains. Interestingly, three central domains A, B, and C are also found in GH13 family enzymes, but domains IV and V are unique to GH70 family enzymes (Fig. 12.2). Domain A harbors the (β/α)8 barrel catalytic core, employing a predicted circular permutation relative to that of GH13 family enzymes (MacGregor et al. 1996). Four conserved amino acid sequence motifs I, II, III, and IV of GH13 family enzymes have been also found in GS enzymes (Fig. 12.2). Three proposed catalytic residues nucleophile (aspartate), general acid/base catalyst (glutamate), as well as transitionstate stabilizer (aspartate) of GSs were verified to be located in the loops following β-strands β4, β5, and β7, in the C-terminal conserved motifs II, III, and IV of domain A, respectively. The catalytic pocket and groove are embraced by a few flexible loops of domain A and B (Leemhuis et al. 2013; Meng et al. 2016b). Domain B is adjacent to catalytic domain A and composed of five or six highly distorted antiparallel β-strands connected by several random loops (Fig. 12.2b). The residues L981, A987, L940, and L938 (Gtf180-ΔN numbering) located in the of loops domain B form the substrate or acceptor binds sites, having a significant effect on the product specificity (Vujičić-Žagar et al. 2010). The only continuous domain C, located at the bottom of the U-shape, comprises an eight-stranded

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β-sheet with a conserved Greek key motif, similar to that of domain C in GH13 family enzymes (Vujičić-Žagar et al. 2010; Uitdehaag et al. 1999). Although this domain is conserved in the GH70 and GH13 families, its accurate function has not been verified yet to date. In addition to catalytic core (domains A, B, and C), GSs have two unique domains (IV and V) of GH70 family enzymes, following the domain B. Domain IV, positioned between domain B and domain V, has partial interactions with domain B, but almost no with domain V (Vujičić-Žagar et al. 2010). Domain IV has no similarity to any known protein structures, and its function excepting as a linker remains unclear. It was proposed that domain IV may play a role similar to a “hinge,” which prompts domain IV with bound α-glucans to approach or stay away from the active sites (Ito et al. 2011). Domain V is attached to domain IV and harbors several ~20 amino acid sequence repeats, which have been verified to anchor α-glucan chains (Giffard and Jacques 1994; Janeček et al. 2000; Kingston et al. 2002; Lis et al. 1995). Interestingly, the solved crystal structures of various GSs showed that domain V has a considerable variability. For instance, compared with Gtf180-ΔN, domain V of GtfA-ΔN displays a deviation of approximately 20 Å relative to other domains (Pijning et al. 2012). Domain V of DSR-E Δ123-GBD-CD2 close to the catalytic core is a significantly different position from that of Gtf180-ΔN (Brison et al. 2012b). The B-factor values of domain V of various GS crystal structures are analyzed to be higher than the other domains, indicating that domain V is more flexible (Pijning et al. 2013). Recently, the crystal structure of whole domain V of the inactive mutant DSR-MΔ2 E715Q, complexed with isomaltotetraose (I4), has been determined. The DSR-MΔ2 enzyme presents a horseshoe-like shape, and the apical residues of domain V is very close to the helix α6 and α7 (5 Å) located within the central (β/α)8 barrel (Claverie et al. 2017). More recently, the crystal structure of ASRΔ2 has been also determined to employ an overall horseshoe-like shape, and its domain V heavily bends toward catalytic sites (Molina et al. 2019). All these results manifest that the domain V of GSs has a considerable positional variability. Additionally, truncation of Gtf180-ΔN domain V has no distinct effects on linkage specificity but results in the production of higher amounts of oligosaccharides and lower amounts of polysaccharides, indicating that domain V of GSs is certainly involved in polysaccharides synthesis (Meng et al. 2015b). To date, the crystal structures of full-length GSs have not been successfully determined yet. Small angle X-ray scattering (SAXS) experiments validate that the N-terminal variable region approximately comprising 700 amino acids further stretches away from domain V, resulting in that the overall molecule of Gtf180 presents a boomerang-like conformation with the corner located in joint point of domain IV and domain V (Pijning et al. 2013).

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12.3.2 Catalytic Mechanism GH70 family GSs have been confirmed to adopt the α-retaining double-replacement reaction mechanism, involving in three previously mentioned catalytic residues, namely, the nucleophile, acid/base catalyst, as well as transition-state stabilizer (van Hijum et al. 2006; Moulis et al. 2006; Uitdehaag et al. 1999). In the first reaction, the (α1-β2) glycosidic linkage of the sucrose substrate is split by the attack of nucleophile, generating a covalent β-D glucosyl-enzyme intermediate. This covalent intermediate is stabilized by transition-state stabilizer. The fructosyl moiety is protonated by acid/base catalyst, synchronously releasing a fructose molecule. In the second reaction, the glucosyl moiety is transported to the non-reducing terminal of an acceptor with retention of the α-anomeric configuration. The α-glucan polysaccharides or oligosaccharides are synthesized from sucrose by the cycle of these two reaction steps. The crystal structure of Gtf180-ΔN D1025N (an inactive mutant) complexed with sucrose substrate showed that seven absolutely conserved residues (Q1059, Y1456, E1063, D1136, H1135, D1025, and R1023) located in the active site, of which six are also conserved in GH13 family enzymes, have interactions with the 1 subsite glucosyl moiety of donor sucrose (Fig. 12.3). The residues R1023, D1025, H1135, D1136, and Q1509 form direct hydrogen bonds with the hydroxyl groups of the glucosyl moiety. The side chains of residues D1025 and E1063 face toward the anomeric C1 atom of the glucosyl moiety and the glycosidic oxygen of sucrose, respectively. The nucleophilic residue D1025 attacks the anomeric C1 carbon of the glucosyl moiety, forming a covalent β-D glucosyl-enzyme intermediate, stabilized by D1136. The acid/base catalyst E1063 protonates glycosidic oxygen, releasing a Fig. 12.3 Sucrose binding at 1 and + 1 subsites in the Gtf180-ΔN mutant D1025N crystal structure in complex with sucrose (PDB: 3KLL). Residues from domain A and domain B are shown as blue and green stick models, respectively. Hydrogen bonds are drawn as dashed lines

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fructose molecule. Residue Y1465 located at the bottom of 1 subsite has hydrophobic interactions with the glucosyl moiety of sucrose. The residues Q1140 and N1141 located on the side of 1 subsite form indirect hydrogen bonds with the C3 hydroxyl groups of the glucosyl moiety mediated by a water molecule, resulting in a pocket-like active site which only contains one glucosyl moiety. This result manifested that GH70 family GSs can only transfer one glucosyl moiety in a reaction cycle. By contrast, GH13 family α-amylases have a cleft-like active site with a series of consecutive subsites. At the +1 subsite of Gtf180-ΔN, the residues E1063, W1065, and D1136, combining with the C3 hydroxyl group of the fructosyl moiety, form a hydrogen bond network (Fig. 12.3). Additionally, the residues Q1140 and N1029 form the hydrogen bonds with the C6 and C1, respectively. However, two hydrophobic residues L981 and L982 from domain B have Van der waals interactions with the fructosyl moiety (Vujičić-Žagar et al. 2010; Meng et al. 2016b). Although the crystal structures of GSs are resoundingly solved, their product specificity, including chain length (α-glucan size) determinants, branch formation, and linkage specificity, has not been thoroughly understood yet.

12.4

Product Specificity of Glucansucrases

12.4.1 Elongation of α-Glucans Sucrose has been found to act as a primer in the initial stage of α-glucan polysaccharide synthesis of GtfA (Dobruchowska et al. 2013) and also as a terminator attached to the non-reducing end of low molecular weight α-glucan (DP 20–30) formed by Gtf-S3 from Streptococcus sobrinus (Cheetham et al. 1991). Additionally, sucrose as an promoter has been also reported in the synthesis of α-glucan polysaccharides of DSR-S from L. mesenteroides NRR B-512F and ASR from L. mesenteroides NRRL B-1355 (Moulis et al. 2006). However, DSR-S from L. mesenteroides NRR B-512F also produces isomalto-oligosaccharides (DP  25), starting from glucose moiety, while oligosaccharides (DP  12) with the non-reducing end of sucrose moiety (Moulis et al. 2006). In DSR-S, both glucose and sucrose are speculated to be initial acceptor for α-glucan polysaccharide synthesis, but glucose is more accepted (Moulis et al. 2006). However, it is still unknown whether other GSs employ this initial acceptor. Concerning GH70 family GSs, there was a controversy over whether the elongation mechanism of α-glucan chains was processive or non-processive. Previously, GSs were deemed to adopt a processive polymerization mechanism based on the detected high molecular weight α-glucans and without intermediate oligosaccharides. Subsequently, oligosaccharides were also detected using more sensitive high performance anion exchange chromatography (HPAEC), suggesting that the non-processive polymerization mechanism is also practicable. Size-exclusive chromatography (SEC) analysis demonstrated that Gtf180-ΔN incubated with sucrose produces a mixed population of high molecular weight α-glucan polysaccharides

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and low molecular weight α-glucan oligosaccharides, but no intermediate size α-glucans (Meng et al. 2015c). Similar situations are also found in DSR-S and GtfA (Meng et al. 2015a; Dobruchowska et al. 2013). Therefore, a semi-processive polymerization mechanism for GSs proposed by Moulis et al. is generally accepted (Moulis et al. 2006). According to this semi-processive mechanism, GSs synthesize α-glucan oligosaccharides using a non-processive action mode in the initial stage of the reaction. When the synthesized oligosaccharides reach a certain size, α-glucan polysaccharides are synthesized using a processive action mode. Particularly, the crystal structure of DSR-MΔ2 from L. citreum NRRL B-1299 complexed with sucrose and isomaltotetraose, which naturally synthesizes low molecular weight dextrans from sucrose, revealed that DSR-MΔ2 employs a clear non-processive mode, caused by a preferred sucrose as a acceptor in the initial reaction, the ability to elongate the growing dextran chains irrespective of their sizes, and a weak binding between domain V and dextran chains (Claverie et al. 2017). Whether processive or non-processive mechanism, their structural basis is proposed to be involved in both the acceptor substrate binding sites and domain V containing several repeat units. In DSR-S, the truncation of A repeats results in a less ability to synthesize polysaccharides compared to the native enzyme (Moulis et al. 2006). In Gtf180-ΔN, truncation of whole domain V obviously impairs the polysaccharide formation and increases the amounts of oligosaccharides (Meng et al. 2015b). However, mutation of the residues located within the acceptor binding site can partly repair the polysaccharides formation of Gtf180-ΔNΔV (Meng et al. 2015b). Additionally, the mutations around the acceptor binding sites, reducing or abrogating polysaccharide formation, have also been reported in GtfR, DSR-S, and ASR (Hendrik et al. 2008; Moulis et al. 2006). These results indicate that the structural basis for processive and non-processive of GSs is located in both domain V and the acceptor binding sites, representing far and near binding sites of growing α-glucan chains, respectively. However, the effects of intermediate binding sites on the polysaccharide synthesis has not been revealed to date, due to lacking the crystal structures of GSs complexed with the larger size α-glucan acceptor substrates. Additionally, the reaction conditions, including sucrose concentrations, enzyme concentrations, pH, and temperatures, have been proven to have distinct effects on the product distribution of GSs (Kim et al. 2003; Falconer et al. 2011). Notably, the molecular size determinants of α-glucan produced by GSs have not been revealed so far.

12.4.2 Branching Specificity The majority of GSs display their respective branching specificity, producing branched α-glucans with various degrees. However, the formation mechanism of branch points is still unknown. As early as 1976, it was proposed that the acceptor reactions of GSs resulted in the branched structures of α-glucans (Robyt and Taniguchi 1976). Mutation of three consecutive residues (S1137, N1138, A1139)

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Fig. 12.4 Maltose binding at +1 and + 2 subsites in the Gtf180-ΔN crystal structure in complex with maltose (PDB: 3HZ3). Residues from domain A and domain B are shown as blue and green stick models, respectively. Hydrogen bonds are drawn as dashed lines

of the crystal structure of Gtf180-ΔN complexed with maltose, which are adjacent to the transition state stabilizer D1136 and close to +2 subsite of maltose (Fig. 12.4), resulted in forming higher degree branches (van Leeuwen et al. 2009). Single and combined mutations of three residues (D1085, R1088, and N1089), which are located in the other side of +2 subsite and have indirect hydrogen bond with acceptor substrate maltose by a water molecule, showed that D1085 and R1088, but not N1089 have remarkable effects on the branch formation (Meng et al. 2016a). Mutation of these three corresponding residues in DSR-S (D460, H463, and T464) increased the ratio of (α1 ! 3) branching linkages (Irague et al. 2013). Mutating the residues A978 and D1028 of Gtf180 to the residues with larger side chains to partly block the groove in the front of +1 subsite decreased the amounts of branching linkages (Fig. 12.4) (Meng et al. 2015c). Together, these results suggested that the formation of branching linkages may be caused by acceptor reaction of GS enzymes. In other words, the orientation changes of recombination, following dissociation of growing α-glucan chain with the acceptor binding subsites, bring the formation of branch points. Although these mutational investigations partly revealed the branching specificity of GSs, the time and location of branch points remain unclear (Meng et al. 2016b).

12.4.3 Linkage Specificity Different LAB-derived GS enzymes synthesize α-glucans with different linkage compositions, which is the major determinant of α-glucan physicochemical properties, such as viscosity and solubility. Recently, a multitude of mutagenesis studies have aimed at the linkage specificity of GSs, but it has not been thoroughly revealed to date. These mutagenesis studies showed that the linkage specificity of GSs is determined by the binding way between acceptor substrates and acceptor binding

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subsites (Moulis et al. 2006; Kralj et al. 2005b; Leemhuis et al. 2012). Before the crystal structures of GSs are available, the regions involved in acceptor binding site have been speculated by sequence alignment with GH13 family enzymes. Three putative regions (3 ~ 5 consecutive amino acids), following three catalytic residues D1025 (Gtf180 numbering, nucleophile in conserved motif II), E1063 (acid/base catalyst in motif III), and D1136 (transition-state stabilizer in motif IV), respectively, display a variability in different GSs and are confirmed to be important to product specificity. Particularly, five consecutive residues (N1134, N1135, S1136, Q1137, and D1138, GtfA numbering), following the transition-state stabilizer D1133, have been proven to significantly influence linkage specificity in several GSs (Kralj et al. 2005b; Moulis et al. 2006; Hendrik et al. 2008; Meng et al. 2014). Combined mutations of the residues N1134, N1135, and S1136 in GtfA altered the linkage specificity from dominating (α1 ! 4) to (α1 ! 6) linkages, and the residue N1134 was subsequently confirmed to play a key part in this change of linkage specificity (Kralj et al. 2005b, 2006). Similar to GtfA, mutations in the three corresponding residues to transform the linkage specificity are also found in DSR-S from L. mesenteroides NRR B-512F, Gtf180 from L. reuteri 180, and GtfR from S. oralis ATCC10557 (Hendrik et al. 2008; van Leeuwen et al. 2009; Moulis et al. 2006). Mutations of the fourth residue Q1140 at C-terminal transition-state stabilizer of Gtf180 resulted in increasing the proportion of (α1 ! 6) linkages or introducing (α1 ! 3) linkages (van Leeuwen et al. 2009). Mutations of the fifth residue D569 at C-terminal transition-state stabilizer of DSR-I from L. mesenteroides NRRL B-1118 increased the percentage of (α1 ! 3) linkages (Cote and Skory 2014). In summary, these results indicated that the residues located C-terminal to the transition-state stabilizer affect linkage specificity, which is a general feature of GS enzymes. The solved crystal structure of Gtf180-ΔN complexing with acceptor substrate maltose structurally illustrated the effects of mutations in acceptor binding site previously observed on linkage specificity (Vujičić-Žagar et al. 2010). This complexed crystal structure revealed that maltose binds at +1 and + 2 acceptor binding sites (Fig. 12.4). The C6 hydroxyl group of non-reducing end of maltose in the direction of the sucrose binding pocket is activated by acid/base catalyst E1063 and then attacks the C1 of the glucosyl-enzymes intermediate, forming an (α1 ! 6) linkage. This binding mode deciphers how an (α1 ! 6) linkage is formed using maltose acceptor substrate. At +1 subsite (Fig. 12.4), the residue N1029 from domain A forms indirect and direct hydrogen bonds with the +1 C3 and C4 hydroxyl groups, respectively, and the residue D1028 from domain A has an indirect hydrogen bond with the +1 C4 hydroxyl group. Four residues (L938, L940, A978, and L981) from domain B constitute a cavity close to the + subsite. Before the available crystal structures, these residues at +1 subsite are rarely studied by site-directed mutagenesis. Recently, these residues, especially from domain B, have been revealed to play a critical role in linkage specificity (Meng et al. 2014, 2015c). At +2 subsite (Fig. 12.4), the residues S1137, N1138, A1139, Q1140, and D1141 are located at one side of the +2 glucosyl moiety, which have been proven to be important for linkage specificity as well. Notably, the residue S1137 forming a direct hydrogen bond with the +2 C1 hydroxyl group has been proven to be the

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main determinant in linkage specificity (Kralj et al. 2005b, 2006; Hendrik et al. 2008). The residues D1085, R1088, and N1089 are located at the other side of the +2 glucosyl moiety, and each of them has an indirect hydrogen bond with the +2 C2 hydroxyl group by the same water molecule. Site-directed mutagenesis studies indicated that these residues also participate in the determination of linkage specificity. In conclusion, the mutation studies at +1 and +2 acceptor binding sites showed that the linkage specificity of GS enzymes is determined by not a single residue but an interaction of various residues from both domain A and domain B, shaping the acceptor binding sites. To date, only the residues located at +1 and +2 acceptor binding sites have been mapped out, and the crystal structures complexing larger acceptor substrates lying in the catalytic pocket have not been obtained yet. Thus, the effects of the residues located at further acceptor binding sites on linkage specificity remain unclear.

12.5

Conclusions and Outlook

With the development of sequencing technology, more and more putative genome sequences of GH70 family GS enzymes are deposited in the gene bank. However, only a small amount of them have been biochemically characterized to date. Intriguingly, these putative GSs are exclusively found in LAB strains and the reason is still unclear. In recent years, the solved crystal structures of GSs have greatly advanced our understanding on the structure–function relationship, especially in product specificity, including sizes, branching formation, and linkage specificity. However, the size determinants, the time and place of branching formation points, and the effects of further acceptor binding sites on linkage specificity remain unknown. Contemplating the future, thoroughly pinpointing the product specificity of GSs would make it possible to synthesize tailor-made α-glucans with desired properties, which can facilitate the industrial applications of α-glucans. The directed evolution combined with high-throughput NMR screening and the protein computational design approach based on the available crystal structures of GSs promises a great potential to further investigate their product specificity. In addition, in order to meet the requirements of industrial applications, the thermal stability of GS enzymes needs to be improved by protein engineering.

References Amari M, Arango LFG, Gabriel V, Robert H, Morel S, Moulis C, Gabriel B, Remaud-Simeon M, Fontagne-Faucher C (2013) Characterization of a novel dextransucrase from Weissella confusa isolated from sourdough. Appl Microbiol Biotechnol 97(12):5413–5422. https://doi.org/10. 1007/s00253-012-4447-8

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Chapter 13

Trends in Enzymology for Functional Carbohydrate Production Qiuming Chen and Wanmeng Mu

13.1

Enzyme Engineering for Industrial Enzymes

Enzyme engineering or protein engineering is one of the most exciting developments in enzymology following various genetic engineering approaches. This technology revolutionized the discovery method for better biocatalysts. As a biocatalyst, natural enzymes generally have poor stereo /regioselectivities, narrow substrate specificities, low catalytic efficiency, poor stability, or high-level reaction product inhibition, which severely hinder the wide application of biocatalysts. Instead of screening the desirable enzymes from natural biocatalysts, enzyme engineering focuses on modifying the sequence of existing enzymes to tailor the biocatalysts with improved properties. Enzyme engineering strategies include directed evolution, rational enzyme design, semi-rational enzyme design, and de novo design. There is no clear definition of these strategies, which might partially overlap (Fig. 13.1). The sections discussed below are not intended to define the strategies but to help the readers better understand the overall framework of enzyme engineering.

13.1.1 Directed Evolution There is a universal phenomenon of evolution in life on earth. Natural evolution improves the fitness of enzymes in new environments. For most of this time without Q. Chen (*) State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China W. Mu State Key Laboratory of Food Science and Technology, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2021 W. Mu et al. (eds.), Novel enzymes for functional carbohydrates production, https://doi.org/10.1007/978-981-33-6021-1_13

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Fig. 13.1 Schematic diagram of the conceptual framework for enzyme engineering strategies

Fig. 13.2 The momentous development of methodology for library generation

even knowing they were doing it, humans harness the evolvability of the natural world to discover better enzymes. In 1984, Eigen and Gardiner proposed a theoretical work flow for improvement of enzymes (Eigen and Gardiner 1984). A decade later, the so-called directed evolution strategy was brought into the laboratory by Frances H Arnold. The strategy is defined as an iterative two-step process involving the generation of mutational library and the screening of this library. An enzyme variant which was active in a highly unnatural environment was obtained by Arnold (1993). Comparing to the natural evolution, directed evolution of enzymes also requires two basic elements: mutation and screening. The mutation procedure is done through intended variation of protein sequences at a defined level of randomness. The screening and selection will be performed to identify those mutants that exhibit better properties. Directed evolution of enzymes relies on genetic engineering to induce targeted changes in the structure and function of the biological molecules, so that it takes place in the laboratory in a short time.

13.1.1.1

Pioneering Technologies for Library Generation

The technical development of the methodology for directed evolution is a continual process (Fig. 13.2). In 1978, Michael Smith and his teammates first proposed sitespecific mutagenesis, which opened the door to protein design (Hutchison et al. 1978). For this contribution, he won the Nobel Prize in Chemistry in 1993. In 1985, The James A. Wells team developed the oligonucleotide-based saturation mutagenesis (OSM) technology to achieve single-point saturation mutations at specific sites of gene sequence (Wells et al. 1985). The team of David W. Leung firstly proposed

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the concept of error-prone Polymerase Chain Reaction (epPCR) (Leung 1989), which was subsequently applied to in vitro modification of antibodies (Hawkins et al. 1992). The principle of epPCR is to increase the random mismatch rate of PCR bases by changing the conditions of the reaction system or using low-fidelity DNA polymerase, generating a mutation library with sequence diversity. This technique does not require the awareness of the structural information of proteins and is simple to operate, which is widely used by researchers. Another prominent contributor to directed evolution methodology was William Stemmer. His team developed a DNA recombination strategy termed “DNA shuffling,” which is a good tool for propagating beneficial mutations while increasing the size of a DNA library. This technology is accomplished by random fragmentation and re-assembly of genes. It is easy to obtain more variation than many other methods (Stemmer 1994a, b). However, one of the disadvantages is that it requires at least 70% identity between gene sequences. DNA shuffling was successfully implemented to improve the activity of β-lactamase (Stemmer 1994a, b). Professor Manfred T. Reetz first applied directed evolution technology in the field of asymmetric catalytic transformation of enzymes in 1997. The enantioselectivity of lipase derived from Pseudomonas aeruginosa was increased from 2% to 81% (Reetz et al. 1997). He found that the mutation sites that are beneficial for chiral selection are mainly concentrated in the substrate binding pocket region. Therefore, the Combinatorial Active-site Saturation Test (CAST) (Reetz et al. 2005) and Iterative Saturation Mutagenesis (ISM) (Reetz and Carballeira 2007) strategies were introduced, which greatly simplified the construction scale of the mutation library, thereby improving the efficiency of library generation.

13.1.1.2

Selection and Screening Techniques

The selection criteria and screening techniques usually need be adapted for each type of enzymatic reaction. Selection may be coupled to a cellular survival function or a spectroscopic enzyme assay. An imprecise approach could result in variants with the desired properties undetected. Therefore, this step is almost the most difficult step of the directed evolution experiment. The selections used in directed evolution are typically limited to enzymes involved in cell growth. Selection of the other enzymes can be performed by fusing the target gene to that of an enzyme with the appropriate properties. When implementing directed evolution to select enzyme activity under non-native conditions, the conditions may have to be considered as selection pressures. The selection pressures usually lead to a direct correlation between cell survival and the desired enzyme property. The selection may be performed with agar plates on which only cells containing a desired variant will survive. Selection is typically followed up by an activity-based screen that involves an enzyme assay (Porter et al. 2016). Microtiter plate screening is widely used in directed evolution strategy. It is often more laborious than selection-based assays but can provide more information. Enzyme variants are usually expressed in individual wells and are assayed through

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reactions that can be monitored with a microplate spectrophotometer. Agar plate screening can also serve as a measure method of the quality of the variants. This method requires a visual signal to identify colonies that express a variant with the desired property. It is usually simpler than microtiter plate screening, but the result is not very precise. Recently microfluidic screening was studied as a screening method for directed evolution. Individual samples or single cells can be compartmentalized into microdroplets and can be screened through fluorescence-activated cell sorting, which is not always possible to employ (Kintses et al. 2010). In conclusion, currently the high manual labor of the screening methods considerably limited the development of directed evolution. Generic high-throughput screen or selection methods available for a variety of enzymes are still needed to be developed for directed evolution.

13.1.2 Rational and Semi-Rational Enzyme Design A clear line between rational and semi-rational enzyme design is difficult to draw. A crucial requirement in rational enzyme design is the awareness of the enzyme’s structural information (Korendovych 2018). For rational design, structure of enzyme is evaluated to propose mutations, which are usually conducted by site-specific mutagenesis. Semi-rational design strategy combines advantages of rational design and directed evolution. Generally, the screening methods with lower throughputs are competent to the ranking task in semi-rational design strategy (Lutz 2010). It utilizes information on sequence and structure of enzymes usually in combination with some computational predictive algorithms to creating smaller and smarter mutation libraries. Advances in computational protein design algorithms have gradually made computers the indispensable working tools for rational and semi-rational enzyme design. The computer-aided enzyme design, also known as in silico enzyme design, is the hot trend in the field of enzyme design. Computational enzyme design to decrease the library size is achieved by screening of virtual libraries computationally and eliminating mutations predicted to be undesirable. Development of computational predictive algorithms is generally based on physical energy or statistical fitness functions. Many application-oriented tools for enzyme engineering have been developed in recent years. Some representative rational design techniques and their general uses are listed in Table 13.1. Note that this list of the rational approaches for enzyme design is by no means exhaustive. We refer the reader to the reviews on the computational design (Chowdhury and Maranas 2020; Ebert and Pelletier 2017; Goldenzweig and Fleishman 2018; Romero-Rivera et al. 2016).

13.1.3 De Novo Design Designing new proteins from scratch are referred to as de novo protein design. Recently, great progress has been made by computational biologists in de novo

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Table 13.1 Representative computational methods for enzyme design Strategy or method Web server or software tool Sequence-based bioinformatics • BLAST National Center for biotechnology information (NCBI) and UniProt • MSA ClustalW and MUSCLE • Ancestral PhyloBot, FastML sequence reconstruction • Consensus HotSpot wizard method Structure-based bioinformatics • B-factorMAP(2.0)3D, PredyFlexy, based method PredBF, and HotSpot wizard • Disulphide Disulfide by design, and bond design SSBOND • Structural modelling

HHPred, RaptorX, SWISSMODEL, Rosetta, Robetta, and Sparks-X Pymol and VMD

• Structural inspection Static systems calculations • Stability FoldX, Rosetta, and changes I-mutant predictor • Molecular Glide, AutoDock Vina, and docking AutoDock Dynamics simulations QM calculaGaussian, Orca, and tion and GAMESS QM/MM MD Amber, CHARMM, simulations Gromacs, and NAMD

General usage

Reference

Sequence searching

McGinnis and Madden (2004) and Madden (2013)

Multiple sequence alignment Ancestral sequence searching Conserved residues searching

Thompson et al. (1994) and Larkin et al. (2007) Hanson-Smith and Johnson (2016) Porebski and Buckle (2016) and Bendl et al. (2016)

Predicting flexibility of residues

Verma et al. (2012) and de Brevern et al. (2012)

Introduction of novel disulfide bonds Protein structure prediction

Craig and Dombkowski (2013) Rohl et al. (2004) and Schaarschmidt et al. (2018)

Structural visualization

Humphrey et al. (1996)

Predicting free energy changes upon mutations Predicting binding modes of protein and ligand

Schymkowitz et al. (2005)

Research on the bond breaking and forming Research on the dynamic behavior of molecules

Morris et al. (2009) and Friesner et al. (2004)

Neese (2012) and Senn and Thiel (2010) Phillips et al. (2005), Van Der Spoel et al. (2005), Brooks et al. (2009)

protein design. This technology is to build a protein that does not exist in nature and to accurately predict how designer proteins will fold to the form with expected functions. De novo protein design plays a role in the field of developing new virus vaccines (Azoitei et al. 2011) and tumor treatment (Procko et al. 2014). In 1997, Dahiyat and Mayo (1997) reported the first case of de novo protein design. A computational design algorithm based on potential functions and stereochemical

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constraints was used to predict optimal side chain conformation for a given backbone pose. Using the module of the zinc finger Zif268 as a template, they successfully designed a ββα protein motif composed of 28 amino acids. This motif was in good agreement with the experimental structure determined by nuclear magnetic resonance (NMR). In 2003, David Baker and colleagues from University of Washington designed and constructed a non-natural 93-residue protein called Top7 using a general computational strategy, which established a novel computational methodology for de novo design. The Rosetta software they used has now been developed into a software suite includes algorithms for computational modeling and design of protein structures (Leaver-Fay et al. 2011). The process of Rosettabased design is generally operated according to the “inside-out” protocol (Kiss et al. 2013). In the first step of the protocol, quantum mechanics calculations are carried out to determine the transition state conformation and to design the theozyme (short for theoretical enzyme). The theozyme is defined as the three-dimensional arrangements of the active center formed by the key catalytic residues of the theoretical enzyme (Dahiyat and Mayo 1997); RosettaMatch is then used to search the database of the native active sites of existing protein structures for selecting the protein backbone structure that can maintain theozyme conformation (Liu and Kuhlman 2006); Rosetta Design or SABER is tasked with generating amino acids located in the active center but not directly involved in catalysis. Rotamer sampling by Monte Carlo simulated annealing is then used to perform multiple rounds of sampling to refine the side chains, backbone, and substrate conformation near the active center. Finally, the designs will be ranked based on empirical criteria, such as Rosetta energy, to evaluate their ability to stabilize the key catalytic residues. The Rosetta energy function is the cornerstone of the Rosetta biomolecular modeling suite. MD simulations are valuable to assess the structural integrity of the theozyme and to expose the design flaws that are intractable from static prediction (Kiss et al. 2010).

13.2

Enzyme Immobilization

13.2.1 Introduction The enzyme immobilization has been a topic of research in the field of biocatalysis for more than 60 years (Robinson 2015). The first industrial application of immobilized enzymes was to produce amino acid. In the early twentieth century, immobilized glucose isomerase was successfully used in the production of high fructose syrup. The technology combines the enzyme with certain carriers in a specialized formulation to maximize the physical stability and enzymatic property of the biocatalyst. It makes the biocatalyst possess completely different characteristics from its free state. Enzyme immobilization can improve the economics of an enzymatic process in industry by improving the enzyme’s reusability. Usually the thermostability of the immobilized enzymes can also be improved compared to the soluble enzyme. Moreover, the microenvironment such as PH surrounding enzymes

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is easier to be controlled through immobilization than that of the free state of enzymes (Novick and Rozzell 2005). The first immobilized form of enzyme was discovered in naturally occurring membrane system, such as enzymes on the rocks, human teeth, and water pipes. The modification effect of immobilization technology on biological enzymes is affected by many factors, such as immobilization carrier, reaction medium, preparation conditions, enzyme molecular properties and organic solvents, etc. There is currently no universal strategy for different immobilized objects. The appropriate immobilization method needs to be selected according to the characteristics of the enzyme and the requirements of application (Yang et al. 2012).

13.2.2 Overview of Traditional Immobilization Techniques After decades of development, traditional immobilization technologies are mainly divided into five categories, namely, adsorption, covalent bonding, entrapment, encapsulation, and cross-linking (Xie et al. 2009). The types of entrapment and encapsulation are sometimes considered as one category (Bornscheuer 2003).

13.2.2.1

Adsorption

The adsorption method is the simplest method of immobilization. It utilizes the weak force between the enzyme and the carrier, such as van der Waals force, hydrophobic force, and surface tension. Therefore, the desorption of enzymes from the carriers is very easy to occur. Materials such as macroporous resin, porous silica glass, and molecular sieves are good carriers for this traditional immobilization technology. They were often used in the application of the first-generation industrial immobilized enzymes. The advantages of the enzyme immobilization by adsorption method includes simple to operate, high recovery rate of enzyme activity, easy to recover carrier, low cost, time-efficient, and no need for chemical modification. However, it is also accompanied by some disadvantages such as unstable form of immobilization, easy to lose the enzymes, and product adsorption to the carriers (Trevan 2014).

13.2.2.2

Covalent Binding

The covalent attachment method is a stable immobilization strategy to form covalent bonds between the amino acid residues on the surface of enzymes and the active groups on the surface of carriers. This method usually requires the carrier to contain many chemical groups or easy to produce chemical bond coupling with enzyme molecules. The covalent attachment method is more rigid and less susceptible to denaturation compared to the one mentioned above (Novick and Rozzell 2005). However, covalently immobilized enzymes are also more expensive and complex

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because of the higher costs of the carriers. In general, the increased stability and less enzyme leaching could compensate the cost. Several functional groups such as the amino of lysine (-NH2), the carboxylate of glutamic acid, and aspartic acid (-COOH), and the thiol of cysteine (-SH), on the enzyme surface, are the covalently binding target of the carriers. The carriers can be either natural polymers, such as modified cellulose, starch, agal polysaccharides, and gelatin, or the synthetic, such as polystyrene, polyacrylates, and polyamides. The materials such as porous glass, metals, sand, and porous ceramics with variety of chemistries can also be chosen to attach the enzymes. Multi-step immobilization is a good technology to enhance the stability of covalent bonding.

13.2.2.3

Entrapment

Entrapment method refers to the use of a gel material with the lattice of polymer matrixes to cage enzyme molecules in a specific structure, such as carrageenan, polyethyleneimine, and polyacrylamide. This immobilization method has the advantages of high immobilization rate and can be used for the co-immobilization of various purpose molecules. However, if the catalytic reaction occurs quickly, the accumulated reaction products is difficult to quickly release through the carriers to the reaction solvent, thereby reducing the reaction rate. The commonly used carrier is alginate, which is widely distributed in the cell walls of brown algae. A judicious selection of precursors, modifiers, and polymerization conditions can control the degree of porosity appropriately (Pizarro et al. 1997). The sol-gel technique is also widely used to entrap enzymes. This gelation process of orthosilicate, such as methyltrimethoxysilane and tetraethyl orthosilicate, can form a cage network, where the enzymes can be trapped into. The entrapment method by nanostructured carriers such as electrospun nanofibers have revolutionized the immobilization technology of enzymes due to its widespread application (Wang et al. 2009; Wen et al. 2011).

13.2.2.4

Cross-Linking

Cross-linking or copolymerization is a carrier-free immobilization strategy that uses bifunctional or multifunctional cross-linking agents, such as glutaraldehyde, dicarboxylic acid, dimethyl adipimidate, etc., to chemically link enzyme molecules. Such a strategy can form a large complex three-dimensional structure with hydrophobicity, through which immobilized enzyme can be separated from the solution. Due to the disorder of the cross-linking reaction, cross-linking may occur at the active center of the enzyme, thereby reducing or inactivating the enzyme activity, which will greatly reduce the recovery rate of the immobilized enzyme. Moreover, the immobilized enzyme cross-linked body formed by simple cross-linking usually has poor mechanical properties. Therefore, the cross-linking method is rarely used

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for enzyme immobilization alone. This technique is usually combined with other immobilization methods to consolidate or improve the effect of immobilization.

13.2.3 New Technology for Enzyme Immobilization The exploration and research of new immobilization technologies never stop. The immobilized carriers possessing excellent physical and chemical properties, such as high degree of porosity and good physical and chemical stability, need to be explored. Appropriate immobilizing carriers can effectively improve the immobilization rate and catalytic efficiency of enzymes. Therefore, lots of researches on novel immobilization technologies on carrier materials have been carried out. In the meantime, to maximize the advantages of the new carrier and make up for its shortcomings, researchers have also conducted a lot of innovative research on the methods and strategies of immobilization techniques. The immobilization technologies on relatively new carriers and formulation methods are discussed below.

13.2.3.1

New Carrier Materials

With the development of interdisciplinary research of biotechnology, materials, and chemistry, new materials emerge constantly. The new carrier materials with the characteristics of large surface area, porous structure, etc. can be mainly divided into nanomaterial, magnetic material, and the carriers modified from traditional materials. The nanocarrier refers to a material with a nanoscale structure. This type of material has a large surface area and good dispersibility, which can greatly improve the immobilization rate and reaction catalysis efficiency of enzymes. The nanosized scaffolds such as spheres, tubes, and fibers have been used for enzyme immobilization. For instance, gold nanoparticles can interact with amino and cysteine groups of proteins as strong as that of the commonly used thiols and thus are excellent biocompatible surfaces for the immobilization. Gold nanoparticles may immobilize the enzymes directly without any modification (Li et al. 2010). Magnetic fields have been utilized in enzyme immobilization. Magnetic material is usually prepared by iron, manganese, cobalt, or their corresponding oxide compounds. The biggest advantage of magnetic material for enzyme immobilization is that it can quickly separate the immobilized enzyme through magnetic attraction. The method is simple and can effectively reduce the cost. Single magnetic particles are generally not directly used for enzyme immobilization. They are usually used in combination with other organic polymers or inorganic porous materials to obtain a high immobilization and recovery rate. Some traditional carriers, such as porous silicon materials, macroporous resins, and molecular sieves, are of excellent performance. However, some defects of them, such as poor stability and difficulty in recycling,

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would hinder their extensive application. The research on modification of such defects is one of the current research hotspots in the field of immobilization.

13.2.3.2

Synthesis of Single Enzyme Nanoparticles

The protocol for the synthesis of single enzyme nanoparticles (SENs) was established by Kim and Grate (2003). This technique can encapsulate each enzyme molecule with a hybrid polymer network to stabilize the enzyme activity. The encapsulating layer is very thin and has a nanometer scale structure, so it can be well dispersed in the solvent. Therefore, the mass transfer resistance between the single enzyme nanoparticle and the reaction substrate is very small, thereby improving the catalytic activity of the immobilized enzyme. Single-enzyme nanoparticles have good catalytic stability and resistance to environmental interference and have a variety of reaction forms, which can be immobilized by multiple enzymes with synergistic effects. Comparing to larger pore sizes, smaller pore sizes of SENs are more favorable for enzyme stabilization. However, the size of SENs equal to or slightly larger than the enzyme dimensions will lead to a significant mass transfer limitation on the diffusion of substrates and products, which might reduce the catalytic efficiency (Kim et al. 2006). Moreover, because of its small size, it is not easy to be recycled and separated. SENs can be used in combination with other porous materials in applications, such as magnetic particles (Yang et al. 2008).

13.2.3.3

Immobilization Assisted by Microwave Irradiation

It is reported that controlled microwave irradiation can dramatically accelerate the process rate of enzyme immobilization. Since different carriers and enzymes have different physical and chemical properties such as hydrophilicity and solubility, there are certain dispersion and contact barriers in the immobilization process. Microwave irradiation can accelerate the mass transfer during both the immobilization process and the catalytic reaction by providing an additional driving force (Singh et al. 2013). The microwave-assisted processes are usually faster and present much better yields comparing to those performed under conventional conditions. However, on the other hand, microwave radiation will affect the binding of enzymes and carriers to a certain extent, which might cause shedding of immobilized enzymes. Therefore, this technology should be carefully applied in the preparation of immobilized enzymes.

Enzymatic Immobilization of Enzyme One of the main goals in enzyme industries is to use green chemistry rather than harsh chemicals to avoid the enzyme denaturation. Therefore, an alternative strategy of enzyme immobilization assisted by enzymes was developed to fabricate solid

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protein formulations (Tanaka et al. 2007; Wong et al. 2008). In the study of Tanaka et al. (2007), microbial transglutaminase (MTG) was utilized to fulfil enzymatic protein immobilization. The enhanced green fluorescent protein and glutathione S-transferase were successfully immobilized onto the casein-coated surface with the help of MTG. This strategy has been also applied for the immobilization of a model acyl carrier protein (Wong et al. 2008) and immobilization of glycosyltransferases onto solid supports (Ito et al. 2010).

13.2.4 Application of Enzyme Immobilization in Food Industry Immobilized enzymes have varied applications in food industry. The advancement of immobilized enzymes lies in the fact that they are economical, environmentally friendly, and easy to be used. In the food industry, the enzyme immobilization technology is commonly used for raw material processing. Lactose intolerance is quite prevalent in people of African, Latino, or Asian ancestry. In a certain amount of areas, large population can only consume lactosefree milk. β-galactosidase can be used to hydrolyze the β-glycosidic bond in lactose in either soluble or immobilized forms. Immobilized β-galactosidase has the advantage of being used both in batch processing and in continuous operation. Lactosefree milk can be produced continuously by immobilized β-galactosidase. Moderate stability of the soluble β-galactosidase is one of the limitations that hinder its application, which can be improved by the immobilization technology (Panesar et al. 2010). Utilizing the immobilized glucose isomerase to produce the highfructose corn syrup is one of the most important application of immobilization technology in food industry (Dicosimo et al. 2013). Pectin is the major component present in most primary cell walls of terrestrial plants. In the fruit juice industry, continuous processes utilizing immobilized pectin lyase were usually implemented for pectin degradation (Li et al. 2008). Up to date, there are few industrial applications of enzyme immobilization for functional carbohydrates production. This might be due to the small market share of functional carbohydrates. With the development of people’s quality of life and increasing attention to health, significant progress in enzyme immobilization will be seen in the field of functional carbohydrates production.

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Prospect of the Enzymatic Production for Functional Carbohydrates

In this chapter, we have shown some technologies of enzyme engineering and enzymes immobilization to solve the current barriers and problems on the way forward large-scale production of functional carbohydrates. At present, to implement the enzymatic progress economically, it is not easy because of the current limitation of enzyme technologies. Solving the technical problems for the enzymatic production of function carbohydrate is a good challenge to be addressed. The high value of the produced function carbohydrate will give commercial feedback to biotechnologists and stimulate the research process on such enzymes. In 2018, the award of one-half the Nobel Prize for Chemistry was awarded to Frances Arnold for the directed evolution of enzymes. This is a landmark in the biocatalysis research field. It proves that improvements can be done to break through the bottlenecks in enzyme application. Although the process is long and arduous, but the future of functional carbohydrates production is bright and will be realized with more and more technological breakthroughs.

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