Industrial Starch Debranching Enzymes 9811970254, 9789811970252

Table of contents :
Preface
Contents
Contributors
Chapter 1: An Overview on Starch Processing and Key Enzymes
1.1 Introduction
1.2 Enzymes Involved in Starch Conversion
1.2.1 Endo- and Exoamylases
1.2.1.1 α-Amylases
1.2.1.2 β-Amylases
1.2.1.3 Glucoamylases
1.2.1.4 α-Glucosidases
1.2.2 Debranching Enzymes
1.2.2.1 Pullulanases
1.2.2.2 Isoamylases
1.2.3 Transferase
1.2.3.1 Cyclodextrin Glucanotransferases (CGTAs, EC 2.4.1.19)
1.3 Conclusion and Future Perspectives
References
Chapter 2: Classification and Enzyme Properties of Starch Debranching Enzymes
2.1 Introduction
2.2 Classification of Microbial SDBEs
2.2.1 Glucoamylases
2.2.2 Pullulanases
2.2.3 Isoamylases
2.3 Sources and Biochemical Properties
2.3.1 Temperature Optimum and Thermostability
2.3.2 pH Optimum
2.3.3 Specific Activity
2.4 Commercial SDBEs
References
Chapter 3: Sequence, Structure, and Engineering of Microbial Starch Debranching Enzymes
3.1 Introduction
3.2 Sequence Classifications
3.3 Structural Features
3.4 Protein Engineering
References
Chapter 4: Production and the Applications in Preparation of Branched Sugar Products of Starch Debranching Enzymes
4.1 Introduction
4.2 Production of SDBEs
4.2.1 Heterologous Expression of SDBEs in E. coli Strains
4.2.2 Heterologous Expression of SDBEs in Bacillus Strains
4.2.3 Heterologous Expression of SDBEs in Yeast Strains
4.3 Conclusions
References
Chapter 5: Recombinant Expression of Starch Debranching Enzymes in Escherichia coli
5.1 Introduction
5.2 Effect of Fermentation Conditions on Soluble Expression of Recombinant Pullulanase
5.2.1 Recombinant B. deramificans Pullulanase Easily Formed Active Protein Aggregates
5.2.2 Effect of Induction Temperature on Fermentation of the Recombinant Strain
5.2.3 Effect of Inducer Concentration on Fermentation of the Recombinant Strain
5.2.4 Effect of Osmotic Pressure Regulator on Fermentation of Recombinant Strain
5.2.4.1 Effects of Osmotic Pressure Regulators on the Fermentation of the Recombinant Strain
5.2.4.2 Effects of Betaine Concentration and Addition Time on Fermentation of the Recombinant Strain
5.2.5 Optimization of High-Density Fermentation Conditions of the Recombinant Strain
5.2.5.1 Effect of Fermentation Temperature on High-Density Fermentation of the Recombinant Strain
5.2.5.2 Effect of Betaine on High-Density Fermentation of the Recombinant Strain
5.2.5.3 Comparison of Fermentation Parameters of Recombinant Strain in Shake Flask and 3-L Fermenter
5.3 Effect of N-Terminal Domain Excision on the Thermostability and Secretory Efficiency of Pullulanase
5.3.1 Construction, Recombinant Expression, and Purification of the N-Terminal Truncated Mutant of Pullulanase
5.3.1.1 Construction and Recombinant Expression of the N-Terminal Truncated Mutant
5.3.1.2 Isolation and Purification of the N-Terminal Truncated Mutant
5.3.2 Construction, Expression, and Purification of Superimposed Mutants
5.3.2.1 Construction of Superimposed Mutants
5.3.2.2 Isolation and Purification of Superimposed Mutants
5.3.3 Analysis of Enzymatic Properties of Mutants
5.3.3.1 Determination of Optimum pH for Mutants
5.3.3.2 Optimum Temperature and Temperature Stability
5.3.3.3 Determination of the Kinetic Parameters of Mutants
5.3.3.4 Substrate Specificity Analysis of Mutants
5.3.4 Application of Mutants in Starch Saccharification
5.4 Surfactant Promotes the Soluble Secretory Expression of Pullulanase in E. coli
5.4.1 Depolymerization of Pullulanase Active Aggregates by Surfactants
5.4.2 The Influence of Surfactant Species on the Growth and Enzyme Production of E. coli
5.4.3 Effect of Triton X-100 Concentration on Growth and Enzyme Production of E. coli
5.4.4 Effect of Triton X-100 Addition Time on the Growth and Enzyme Production of E. coli
5.4.5 Effect of Triton X-100 on High-Density Fermentation of E. coli in a 3-L Fermenter
5.5 Optimization of the Induction Method Combined with the Glycine Feeding Strategy to Promote the Extracellular Expression of...
5.5.1 Effect of Inducer Type and Concentration on Growth and Enzyme Production of E. coli
5.5.2 Effect of Glycine Concentration on Growth and Enzyme Production of E. coli
5.5.3 Effect of Lactose Flow Acceleration on Pullulanase Production by High-Density Fermentation of E. coli in a 3-L Fermenter
5.5.4 Effect of Induction Time on Pullulanase Production by High-Density Fermentation of E. coli in a 3-L Fermenter
5.5.5 Effect of Glycine Feeding Strategy on Pullulanase Production by High-Density Fermentation of E. coli in a 3-L Fermenter
5.5.6 ``Mixed Conformation´´ Model of Pullulanase Expression in E. coli
5.6 Recombinant Expression of Thermobifida fusca Isoamylase in E. coli BL21(DE3)
5.6.1 Cloning and Sequence Analysis of the T. fusca Isoamylase Gene
5.6.2 Construction of Recombinant Plasmid and Strain
5.6.3 Shake Flask Fermentation of the Recombinant Strain and Purification
5.6.4 High-Density Fermentation of the Recombinant Strain in a 3-L Fermenter
5.7 Recombinant Expression of T. Fusca Isoamylase in E. coli MDS42
5.7.1 Construction of the Recombinant Strain E. coli MDS42/Tfu_1891-pSX2
5.7.2 Expression of Recombinant Strain
5.7.3 Optimization of Fermentation Conditions for Recombinant Strain E. coli MDS42/Tfu_1891-pSX2 by Shake Flask Cultivations
5.7.3.1 Effect of Induction Temperature on the Growth and Enzyme Production of the Recombinant Strain
5.7.3.2 Effect of Inducer Type and Concentration on the Growth and Enzyme Production of the Recombinant Strain
5.7.4 Optimization of Fermentation Conditions for Recombinant Strain E. coli MDS42/Tfu_1891-pSX2 by 3-L Fermenter Cultivation
5.7.4.1 Effect of Induction Temperature on the Growth and Enzyme Production of the Recombinant Strain
5.7.4.2 Effect of Induction Time on the Growth and Enzyme Production of the Recombinant Strain
5.7.4.3 Effect of Inducer Concentration on the Growth and Enzyme Production of the Recombinant Strain
5.7.5 Comparison of Fermentation Parameters of Recombinant Isoamylase Production in Shake Flasks and 3-L Fermenters
References
Chapter 6: Production of Starch Debranching Enzymes in Bacillus Strains
6.1 Introduction
6.2 Recombinant Expression of Pullulanase in B. choshinensis
6.2.1 Construction of the Recombinant Strain B. choshinensis (pNCMO2/pulA-d2)
6.2.2 Optimization of Shake-Flask Cultivations
6.2.2.1 Original Fermentation Medium Optimization
6.2.2.2 Carbon Source Optimization
6.2.2.3 Nitrogen Source Optimization
6.2.2.4 Metal Ion Optimization
6.2.2.5 Key Factor Optimization Using Response Surface Methodology
6.2.2.6 Seed Growth Curve and Seed Cultivation Time Optimization
6.2.2.7 Inoculation Ratio Optimization
6.2.2.8 Original pH Optimization
6.2.2.9 Cultivation Temperature Optimization
6.2.3 Optimization of 3-L Fermenter Cultivations
6.2.3.1 Batch Fermentation in a 3-L Fermenter
6.2.3.2 Effect of pH on Fermentation of Recombinant B. choshinensis
6.2.3.3 Effect of Dissolved Oxygen on Fermentation of Recombinant B. choshinensis
6.2.3.4 Effect of Inorganic Nitrogen Source on Fermentation of Recombinant B. choshinensis
6.2.3.5 Effect of Beef Extract Concentration on Fermentation of Recombinant B. choshinensis
6.2.4 The Mechanism by which Magnesium Ions Promote the Expression of ``High Activity´´ Pullulanase in Recombinant B. choshine...
6.2.5 Magnesium Ions Promote the Expression of ``High Activity´´ Pullulanase in Recombinant B. choshinensis
6.2.5.1 Effect of Magnesium Ions on the Expression of Other Heterologous Proteins in B. choshinensis
6.2.5.2 Effect of Magnesium Ions on the Specific Activity and Aggregation of Pullulanase Pure Enzyme Expressed by B. choshinen...
6.2.5.3 Heat Treatment Distinguishes the Active and Inactive Forms of Pullulanase
6.2.5.4 Effect of Magnesium Ions on the Secondary Structure of Pullulanase Expressed by B. choshinensis
6.2.5.5 Effect of Magnesium Ions on the Cell Morphology of B. choshinensis
6.2.5.6 Effect of Magnesium Ion on Transcription and Protein Expression of pulA-d2 and HWP Genes
6.2.6 ``Folding Master´´ Mechanism of Pullulanase Expression in B. choshinensis
6.3 Recombinant Expression of Pullulanase in B. subtilis
6.3.1 Construction of CRISPR/Cas9 Gene Editing System Disruption Plasmid
6.3.2 Disruption of the srfC Gene by the CRISPR/Cas9 Gene Editing System
6.3.3 Disruption of the spoIIAC Gene by the CRISPR/Cas9 Gene Editing System
6.3.4 Disruption of nprE and aprE Genes by the CRISPR/Cas9 Gene Editing System
6.3.5 Disruption of the amyE Gene by the CRISPR/Cas9 Gene Editing System
6.3.6 Transformation of Screened Wild-Type B. subtilis
6.3.7 Improved Pullulanase Expression Through Promoter Optimization
6.3.7.1 The Expression of β-CGTase Was Enhanced by Single Promoter Optimization
6.3.7.2 The Expression of β-CGTase Was Enhanced by Dual-Promoter Optimization
6.3.7.3 Expression of Pullulanase and α-CGTase Using the Dual Promoter PHpaII-PamyQ
6.3.7.4 Fermentation Culture of Recombinant Strain WS5PUL in a 3-L Fermenter
Protease Disruption and Optimization of Pullulanase Fermentation Conditions
Knockout of Six Extracellular Proteases
Expression of Pullulanase in WS11
Fermentation of Recombinant Strain WS11PUL in a 3-L Fermenter
6.3.7.5 Optimization of Fermentation Conditions of Recombinant Strain WS11PUL
6.3.8 The Effect of Protease on the Expression of Pullulanase and the Optimization of Fermentation Feed Solution
6.3.8.1 Effect of Protease on the Expression of Pullulanase in Shake Flask Fermentation
6.3.8.2 Effect of Protease on the Expression of Pullulanase in a 3-L Fermenter
6.3.8.3 Purification of Pullulanase Samples, Determination of Kinetic Parameters and Aggregation State under Different Proteas...
6.3.8.4 Thermal Stability of Pullulanase Samples at Different Protease Concentrations
6.3.8.5 Improve the Activity of Pullulanase by Optimizing the 3-L Fermenter Feeding Solution
6.3.9 Increasing Cell Wall Negative Charge and Optimizing Signal Peptide to Improve the Expression of Pullulanase
6.3.9.1 Effect of Chaperone Protein on the Expression of Pullulanase in WS11
6.3.9.2 Effect of hrcA Gene Knockout on the Expression of Pullulanase
6.3.9.3 The Chaperone Gene Was Coexpressed on Plasmid
6.3.9.4 The Chaperone Protein Gene Was Expressed on the Pullulanase Expression Plasmid
6.3.9.5 Effect of Chaperone GrpE on the Expression of Pullulanase in WS4, WS5, WS9, and WS10
The Chaperone Protein Gene Was Expressed on the Pullulanase Expression Plasmid
6.3.9.6 The Effect of Chaperone Gene Integration into the Genome on the Expression of Pullulanase
6.3.10 Effect of dltB Gene Knockout on the Expression of Pullulanase
6.3.11 The Effect of lytC Gene Knockout on the Expression of Pullulanase
6.3.12 Effect of Signal Peptide on the Expression of Pullulanase
6.3.12.1 Screening of Signal Peptides
6.3.12.2 Expression of Different Signal Peptides in WS9D and WS9DL
6.3.13 Fermentation of Recombinant Strains WS9DPUL/ywtF and WS9DLPUL/ywtF in a 3-L Fermenter
References
Chapter 7: Applications of Starch Debranching Enzymes in Starch Processing
7.1 Introduction
7.2 Glucose and Oligosaccharides
7.2.1 Glucose and Malto-Oligosaccharides
7.2.1.1 Two-Step Liquefaction-Saccharification
7.2.1.2 One-Step Liquefaction-Saccharification
7.2.2 Trehalose
7.2.3 Isomalto-oligosaccharides
7.3 Cyclodextrins
7.4 Cycloamyloses
7.5 Isomalto/Malto-Polysaccharides
7.6 Slowly Digesting Starch and Resistant Starch
7.6.1 Production of RS and SDS by Debranching Enzyme Treatment
7.6.1.1 Pullulanases
7.6.1.2 Isoamylase
7.6.1.3 Amylopullulanase
7.6.2 Production of RS and SDS in Combination with Debranching Enzymes and Other Treatments
7.6.2.1 Dual Treatment by Amylase and Pullulanase
7.6.2.2 Dual Treatment by Pullulanase and Amylosucrase
7.6.2.3 Dual Treatment by Pullulanase and Ultrasound
7.7 Complex Starch
7.7.1 Production of Starch Complexes by Debranching Enzyme Treatment
7.7.2 Production of Starch Complexes in Combination with Debranching Enzymes and Other Treatments
References

Citation preview

Jing Wu Wei Xia   Editors

Industrial Starch Debranching Enzymes

Industrial Starch Debranching Enzymes

Jing Wu • Wei Xia Editors

Industrial Starch Debranching Enzymes

Editors Jing Wu State Key Laboratory of Food Science and Resources Jiangnan University Wuxi, Jiangsu, China

Wei Xia State Key Laboratory of Food Science and Resources Jiangnan University Wuxi, Jiangsu, China

ISBN 978-981-19-7025-2 ISBN 978-981-19-7026-9 https://doi.org/10.1007/978-981-19-7026-9

(eBook)

© The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 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

Starch is a polysaccharide polymerized with glucose as the basic unit. It is one of the most abundant carbohydrates in nature. It is not only the main source of nutrition for humans and animals but also one of the most important industrial raw materials. The glycosidic bonds in starch mainly include α-1,4 bonds and α-1,6 bonds. Although the α-1,6-glycosidic bonds account for only approximately 6% of the total glycosidic bond, the starch chain forms branching structures. Most natural starches contain approximately 20–25% amylose and 75–80% amylopectin. The enzymes used in amylase hydrolysis are generally called starch-processing enzymes, which mainly include starch endonuclease, starch exonuclease, starch-debranching enzyme, and glycoside transferase. Because starch has very complex structures, it is difficult for a single enzyme to hydrolyze completely, so the synergism of many enzymes is usually needed in the process of enzymatic hydrolysis of starch to produce oligosaccharides or monosaccharides. Although the content of the branching α-1,6 bonds is low in starch, it limits the catalytic efficiency of α-1,4-glycosidic bond hydrolase and forms the limit dextrin that cannot be used. The utilization rate of raw materials and production efficiency can be effectively improved if α-1-line 6-glycoside hydrolase, that is, starch-debranching enzyme, is compounded during starch hydrolysis. Pullulanase and isoamylase are the most important starch-debranching enzymes, which can specifically and efficiently hydrolyze the branching bonds in amylopectin, so they are widely used in industry. The addition of pullulanase or isoamylase in the process of amylase hydrolysis can accelerate the speed of amylase hydrolysis, shorten the hydrolysis time, improve the conversion rate of products, and reduce the use of other enzyme preparations. Starch-processing products have been widely used in food, medicine, paper, and many other industries for a long time because of their wide consumption field and large consumption quantity. With the continuous emergence of new starchprocessing products and processes, the performance requirements of starchdebranching enzymes in the modern starch-processing industry are increasing. A comprehensive understanding of the structure–function relationship of SDBEs and the true needs of specific applications in which they are employed are necessary to v

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Preface

broaden their application prospects. The development of a series of starchdebranching enzymes with high catalytic activity, high specificity, and high yield will become a new trend in the research of starch-debranching enzymes. This book mainly summarizes the latest research progress of family starch debranching enzymes from microorganisms and expounds the characteristics, research status, practical application, and research trend of pullulanase and isoamylase. First, an overview of starch processing and key enzymes is introduced by Prof. Ranjana Das and Arvind M. Kayastha. Wei Xia, Lei Wang, and Prof. Jing Wu summarized the concept, the original sources, different properties, and substrate specificities of the major groups of starch-debranching enzymes and introduced the main features of the sequence and structure of microbial starch-debranching enzymes and relevant engineering studies. Then, the applications in starch processing and preparation of branched sugar products of starch-debranching enzymes are summarized by Wei Xia, Lingqia Su, and Sheng Chen. Moreover, considering industrial-scale applications, overexpression strategies for starchdebranching enzymes in Escherichia coli and Bacillus strains were introduced by Kang Zhang, Zhanzhi Liu, and Zhengfei Yan. We expect that this book will be used as an important reference for graduate students, researchers, and bioengineering experts who are engaged in starchprocessing research and industry. On behalf of my coeditor Dr. Wei Xia, I express our sincere appreciation to all authors for their contribution and dedication. Wuxi, Jiangsu, China

Jing Wu Wei Xia

Contents

1

An Overview on Starch Processing and Key Enzymes . . . . . . . . . . . . Ranjana Das and Arvind M. Kayastha

2

Classification and Enzyme Properties of Starch Debranching Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wei Xia and Jing Wu

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Sequence, Structure, and Engineering of Microbial Starch Debranching Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wei Xia, Lei Wang, and Jing Wu

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Production and the Applications in Preparation of Branched Sugar Products of Starch Debranching Enzymes . . . . . . . . . . . . . . . . . . . . Wei Xia, Sheng Chen, and Jing Wu

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Recombinant Expression of Starch Debranching Enzymes in Escherichia coli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kang Zhang, Zhanzhi Liu, and Jing Wu

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Production of Starch Debranching Enzymes in Bacillus Strains . . . . 139 Kang Zhang, Zhengfei Yan, and Jing Wu

7

Applications of Starch Debranching Enzymes in Starch Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Lingqia Su and Jing Wu

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Contributors

Sheng Chen State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China Ranjana Das School of Biotechnology, Institute of Science, Banaras Hindu University, Varanasi, India Arvind M. Kayastha School of Biotechnology, Institute of Science, Banaras Hindu University, Varanasi, India Zhanzhi Liu State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China Lingqia Su State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China Lei Wang State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China

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Contributors

Jing Wu State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China Wei Xia State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China Zhengfei Yan State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China Kang Zhang State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, Jiangnan University, Wuxi, China International Joint Laboratory on Food Safety, Jiangnan University, Wuxi, China

Chapter 1

An Overview on Starch Processing and Key Enzymes Ranjana Das and Arvind M. Kayastha

Abstract Starch makes an important component of human and cattle diets. A large portion of our food and fodder originates from them. In addition, chemically or enzymatically processed starch is used as a variety of products such as starch hydrolysates, glucose syrups, maltose, fructose, starch or maltodextrin derivatives or cyclodextrins. These enzymes for starch processing exist in natural resources and are categorized as (i) endoamylases, (ii) exoamylases, (iii) debranching enzymes, and (iv) transferases; on the type of reaction, they catalyze. The individual category and their corresponding enzymes are discussed in detail in this chapter. The aim of this chapter is a general study of these enzymes’ operation in various industries for the production of newly refined starch. Keywords Starch hydrolysis · α-Amylase · β-Amylase · Glucoamylase · α-Glucosidase · Pullulanase · Isoamylase

1.1

Introduction

Starch, an amylum, is a polymeric carbohydrate consisting of several glucose units held together by glycosidic bonds. Having its origin from plants, cellulose is the second most profuse compound synthesized by plants (cellulose is the first) and found in granular form. It is one of the important food products and bioresources used globally for varied purposes. Despite maintaining its traditional use in the food industry, established as an important starting point of energy for humans, technological advancement has led to its increased application in many other sectors, such as health, medicine, paper, textile, chemicals, petroleum engineering, construction engineering, and agriculture. The food industry utilizes starch either as food items or thickening agents, preservation or aimed at improving quality in baked foods, confectioneries, pastas, soups, sauces, mayonnaises, etc. Pure starch is a white powder, tasteless with no odor, that is, insoluble in cold water or alcohol. Depending

R. Das · A. M. Kayastha (✉) School of Biotechnology, Institute of Science, Banaras Hindu University, Varanasi, India © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 J. Wu, W. Xia (eds.), Industrial Starch Debranching Enzymes, https://doi.org/10.1007/978-981-19-7026-9_1

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R. Das and A. M. Kayastha

Fig. 1.1 Chemical structure of amylose and amylopectin

on the species, starch granules produced by each plant have specific structures and compositions, such as the amylose/amylopectin unit constituting the glucose chain, and the ratio of fat and protein content of the storage organ also varies considerably. Therefore, starch stands out depending on the source. This innate functional heterogeneity due to the varied biological sources broadens its scope for application in industries [1]. Naturally occurring starch is made up of two types of α-D-glucan chains: 80% amylopectin and 20% amylose. Amylose is the linear polymer of glucose with α-1,4 bonds weighing approximately 106 Dalton; on the other hand, amylopectin has 5% α-1,6 linkages in addition to the aforesaid bonds and weighs approximately 108 Dalton. Accordingly, we can see that amylopectin has many microbranches, but amylose is composed of a few long branches [2, 3]. The composition and arrangement of starch are given in Fig. 1.1. The build-up, molecular weight, and ratio of amylopectin to amylose are different in starches extracted from various sources. This has led to diversification in the characteristics of starches obtained from diverse sources [4]. Starches are classified into various types, such as type A or B starch (based on crystallinity pattern), waxy or hylon starch (based on amylose/amylopectin ratio), etc. Waxy starch consists of a good proportion of amylopectin, but the hylon type comprises far more than 50% amylose [5]. The composition and structure disparities in starches have varied origins, which determines their properties and mode of cooperation with other elements of foods

1

An Overview on Starch Processing and Key Enzymes

3

that give the unique final product, desired texture, and taste. In the agricultural food sector, starch is applied as a supplement that checks the consistency, durability, and texture of soups and sauces to combat gel breakdown during treatment to extend the shelf life of various food items [1]. Starch is comparatively easy to extract and does not require tedious downhill purification processes. It is synthesized in large amounts in major plant sources such as cereal grains (rice, corn, wheat, barley, sorghum, etc.), tubers (potato), and roots (tapioca). These sources are cost-effective, easily available, and imitate raw materials for large-scale industrial manufacturing [6]. Corn starch solely contributes 80% of the production of starch in the world market. It is one of the major ingredients in the manufacturing of various food products and has been extensively used as a thickening agent, stabilizer, colloidal gelling agent, water holding, and sticking agent due to its adaptable physicochemical attributes [7]. Starches from tubers such as potatoes are considered nonconventional sources and have gained importance in furnishing alternatives for expanding the spectrum of wanted functional assets, which are requisites for the development of value-added food products. The consistency of native granular starch under changed physiological values of pH and temperature varies adversely. For example, native starch granules are insoluble in water below 60 °C and extremely resistant to hydrolysis by starchdebranching enzymes such as amylase. Accordingly, native starch has confined functionality. Below 60 °C, starch granules are insoluble. Above this temperature, the granules retained water, leading to swelling. This temperature is characteristic of the type of starch and is known as the gelatinization temperature. As heating is continued, the granules open up, and starch aggregates are produced. This colloidal suspension of starch is called “starch paste”. Paste making by starch is its most important capability since enzymatic conversions are susceptible to this paste only [8]. To achieve enhanced properties and desirable capabilities, such as solubility, viscosity, texture, and thermostability, which are needed for the required product or persona in the industry, native starches are altered. Considering the panorama of utilization of starches with distinguishing properties, investigations of nonconventional starches and other native starches have become more obligatory [9, 10]. Current advances in the structure–function relationship of starches from distinct sources further provide valuable statistics for improving efficiency in industrial applications. Modified adaptation is achieved mostly by physical or chemical means. Biotechnological modification via an enzymatic approach is increasingly being explored [11]. Physical modification methods are inexpensive and simple, viz., dry heating at high temperature, continuous deep freezing and thawing, simultaneous increase or decrease of pressure, etc. Chemical modification introduces unique functional elements into the starch molecule through its hydroxyl groups, which results in important changes in its physicochemical properties (Fig. 1.2). The functional properties of chemically modified starch depend on various factors, such as its biological origin, reagents, reagent concentration, pH, reaction time, catalyst, type and degree of substitution, and the dissemination of the substituent in the modified starch molecule.

R. Das and A. M. Kayastha

4

Fig. 1.2 Various processes for starch modification

The enzymatic approach of starch modification helps in designing a starch with a fresh structure, in which the molecular mass, branch chain length, and amylose: amylopectin ratio can be amended by enzymatic reactions when the reaction of enzymes takes place with gelatinized starch. Various enzymes associated with the degradation of starch are shown in Fig. 1.3. They normally produce starches with changed physicochemical properties and structural characteristics for various food and nonfood applications. The current scenario shows that starch processing enzymes have yearly 30% of the global enzyme market share. The most common commercial application of starch debranching enzymes in the food industry for the production of branched-chain sugar products is shown in Table 1.1. Herein, we will discuss certain starch debranching enzymes in the production of branched-chain sugar products.

1.2

Enzymes Involved in Starch Conversion

Starch-converting enzymes can be categorized as (i) endoamylases, (ii) exoamylases, (iii) debranching enzymes, and (iv) transferases on the basis of the type of reaction being catalyzed.

1

An Overview on Starch Processing and Key Enzymes

a,1-4 + a,1-6 hydrolysis

a,1-6 hydrolysis

5

isoamylase/ amylopullulanase

a,1-6 transferase glucoamylase/ a-glucosidase

glucan branching enzyme

a,1-4 hydrolysis a,1-4 transferase cyclodextrin glycosyltransferase

b-amylase/ maltogenic amylase

amylomaltase b-amylase

Fig. 1.3 Various enzymes are involved in the shortening of starch chains. The open chain structure epitomizes the reducing end of a polyglucose molecule (Reused with permission from Van Der Maarel MJ et al., Journal of Biotechnology 2002, 94:137–155 [12]) Table 1.1 The most common trading application of starch debranching enzymes in the agri-food industry for producing branched-chain sugar products Enzyme(s) α-Amylase, β-amylase, glucoamylase, pullulanase, isoamylase, maltogenic amylase α-Amylase, β-amylase α-Amylase, β-amylase α-Amylase, β-amylase, pullulanase, debranched enzymes, branching enzymes, maltogenic amylase, glucoamylase, cyclodextrin glucanotransferases α-Glucanotransferases

1.2.1

Application Starch saccharification Starch liquefaction Baking industry Anti-staling

Cycloamylose production

Endo- and Exoamylases

The group endoamylases cleave α-1,4 glycosidic linkages present in the inner part (endo-) of starch (the amylose or amylopectin chain). According to EC, α-amylase (EC3.2.1.1) is commonly called endoamylase and is extensively distributed among microorganisms, belonging to archaea as well as bacteria, plants, and animals [13]. The final products of α-amylase action comprise oligosaccharides of diverse lengths with an α-configuration and α-limit dextrins, which constitute branched

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chain oligosaccharides. The second class of enzymes, exoamylases, either exclusively cleave α-1,4 glycosidic linkages, such as β-amylase (EC 3.2.1.2), or cleave both α-1,4 and α-1,6 glycosidic linkages, such as amyloglucosidase or glucoamylase (EC 3.2.1.3) and α-glucosidase (EC 3.2.1.20). Exoamylases act inward on the external glucose residues of amylopectin or amylose in the starch polysaccharide and thus produce either only glucose (glucoamylase and α-glucosidase) or maltose and β-limit dextrin (β-amylase). The European food enzyme market is expected to trundle at the rate of 4%, with revenues already reaching €700 million in 2019, and amylases contributed 25% of the enzymes. β-Amylase and glucoamylase are unique in the sense that they change the anomeric configuration of the liberated maltose from α to β. α-Glucosidase works excellently on short-length maltooligosaccharides and produces glucose with an α-configuration, while glucoamylase hydrolyzes longchain polysaccharides best. Glucoamylases and β-amylases were isolated for the first time from microorganisms. enzymes are cyclodextrin Furthermore, exo-acting amylolytic glycosyltransferase (EC 2.4.1.19), an enzyme possessing transglycosylation activity, maltogenic α-amylase (glucan 1,4-α-glucanohydrolase, EC 3.2.1.133), an amylase from B. stearothermophilus releasing maltose [14], and maltooligosaccharide producing amylases, e.g., the maltotetraose fabricating enzyme from P. stutzeri (EC 3.2.1.60) [15] or the maltohexaose producing amylase (EC3.2.1.98) from K. pneumoniae [16]. Let us discuss some of the exo- and endo-acting enzymes in the degradation of starch for the synthesis of sugar products.

1.2.1.1

α-Amylases

These enzymes (EC 3.2.1.1), grouped as endoamylases, break α-1,4 glycosidic bonds that subsist in the interiors of amylose or amylopectin chains but resign hydrolysis when α-1,6 linkages are reached [17]. Accordingly, they form linear and branched oligosaccharides and α-limit dextrins [12, 18]. These enzymes are rich in bacterial and fungal sources for industrial procurement. The bacterial α-amylases are mostly thermoresistant and react at normal pH, and calcium ions are essential for their activity, while the fungal α-amylases lose activity at high temperatures. α-Amylases have been categorized into three glycoside hydrolase (GH) families (GH-13, GH-57, and GH-119) in the Carbohydrate-Active enZYmes (CAZy) database (http://www.cazy.org/). A large number of α-amylases (>10,000) are from the GH-13 family, and their (bacteria, archaea, and eukaryotes) 3-D structures have been deciphered previously [19]. Despite the fact that the overall sequence identity within the GH-13 family is extremely low [20], it is well documented that several GH-13 family individuals comprise a conserved (β/α)8 barrel that is tucked into their 3-D structure. The structures of these α-amylases generally consist of three domains: a conserved (β/α)8-barrel domain (domain A), an additional domain inserted within domain A (domain B), and the C-terminal β-sheet domain (domain C). In addition, 3 unchangeable catalytic residues (Asp-Glu-Asp) are located in domain A, and calcium ions that enhance stabilization many-folds are found in its structure

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[21]. In addition, α-amylase from B. thermooleovorans contains an extra chloride ion binding site in its active site that has led to the enhanced catalytic efficiency of the enzyme, possibly due to an increase in the pKa of the hydrogen-donating residue in the active site [22, 23]. α-Amylase is an important enzyme for all starch-based industries, completely replacing chemical hydrolysis in starch-based industries. It inhabits applications in all industrial processes involving paper, sugar syrup, brewing, ethanol production, textiles, and the detergent industry for starch hydrolysis. It assists in curing digestive disorders. Pretreatment of animal feed with α-amylase leads to better digestion. In molecular biology, α-amylase plays a role in the selection of an advantageous inclusion of a reporter constructs along with antibiotic resistance. Since reporter genes are fringed by homologous regions of the structural regions for α-amylase, successful inclusion will break the α-amylase gene and suppress starch degradation, which can be easily tested using iodine staining. α-Amylase and pullulanases combined are applied for productive anti-staling property, which prevents the gumminess arising from the sole action of α-amylase on maltodextrins. This drawback is overcome by pullulanases, which hydrolyze α-1,6 glycosidic bonds of amylopectin. The primary market for α-amylase is the generation of starch hydrolysates, viz. dextrins, glucose, and fructose. This uses a liquefaction point where starch is solubilized primarily at high temperatures and then fractionally hydrolyzed with thermostable α-amylase. The hydrolyzed products are once again distressed by the next lot of hydrolyzing enzymes to yield high-fructose corn syrup (HFCS). This enzyme has been purified and characterized from various plants, microorganisms and fungal sources. α-Amylase from wheat [24], ungerminated soybean [25], germinated soybean [26] mung bean [27], etc., and various bacterial sources have been purified. In any enzyme development process, the enzyme itself is the limiting element to be wasted in each process. Thus, to overcome this problem, immobilization is necessary. Various conventional and nanomaterials are currently employed for enzyme immobilization. Since immobilized enzymes offer better kinetic parameters, they are frequently used in enzyme technology. α-Amylase from various sources has been immobilized on DEAE-cellulose, Amberlite MB, chitosan, and graphene from our laboratory [28–30].

1.2.1.2

β-Amylases

Another important enzyme of the starch and food industry is β-amylase (EC 3.2.1.2). An exo-hydrolase, operating two glucose units inward from the nonreducing end, breaks the α-1,4 glycosidic bond in the polysaccharide chain, yielding maltose, which gives up at branched sites since it cannot circumvent branched α-1,6 linkages [31, 32]. This exo-acting enzyme cannot distinguish α-1, 6 branches like α-amylases and cannot pass the barrier and thus end in high molecular weight dextrin, leading to β-limit dextrins [33]. The uniqueness of the ultimate product maltose lies in its being less sweet and companying better with flour in contrast to sucrose. Agri-food business benefits from the enzyme, since it prolongs the shelf life and finds

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application in mashing and brewing [34, 35]. These enzymes together with debranching enzymes are extensively in demand for the production of syrups in industry. The maltose syrups are moisture absorbing and have a more invariable color compared to glucose syrups. In addition, they find utility in the confectionery industry and in frozen desserts due to their lower crystallization and adhesion properties [36, 37]. The potential of β-amylase to cleave the exterior arms of starch lowers the retrogradation of starch-based foods [38, 39]. The prevention of starch retrogradation is brought about by the enzyme by shortening α-1,4 linkages in the linear chain of starch, taking advantage of its exo-type activity, thereby overcoming the intermolecular association of the linear portion of starch, i.e., amylose. Furthermore, the softness of starch-containing food is achieved by retaining moisture in maltose, produced by the action of enzymes. Hence, the enzyme is exploited in arresting retrogradation of rice cake. β-Amylase found in wheat flour effectively obstructs retrogradation during the course of baking, although barley-originated enzyme is used occasionally; its application is limited due to thermostability [40, 41]. Commercially, β-limit dextrins are produced by completely dissolving/dispersing starch in water by boiling (to gelatinize) and then treating it with β-amylase (with highly purified enzyme). The hydrolysis process is performed under optimal conditions of the enzyme until it interrupts the conversion of external chains of amylopectin maltose as a consequence of branch points. In general, the experimentation involves a starch content of 10% in acetate buffer (0.02 M, pH 4.8) and β-amylase added at 200 U/g of the starch while incubating the system at 37 °C. Waxy starches (e.g., waxy maize) are favored due to their high amylopectin content. After hydrolysis is complete, the enzyme is inactivated by boiling the mixture, and the precipitated protein is removed by centrifugation. Subsequently, β-limit dextrins are gathered and dried by spray or lyophilization. There are very few reports on the applications of β-limit dextrins. Applications include carriers (nutrients), bulking agents, texture providers, spray-drying aids, fat replacers, film formers, and freeze-control agents to arrest crystallization and to provide nutritional value as calories [42]. They also find applications as bioadhesive [43]. The functionality of β-limit dextrin is imparted by its unique structures (heavily branched α-glucans) and eventually its physical properties in solution, such as high molecular weight amorphous viscosifying α-glucan that resists retrogradation because of its branched structure. The dextrins do not make clear solutions like sugars, but they form colloidal suspensions (31% by weight). Despite being high molecular weight, they do not precipitate from dispersion/solution due to their highly branched structure, making them logical as clouding agents in drinks. β-Amylase isolated from ramie leaf by He et al. [44] and peanut by [38] had moderate thermostable enzyme properties. Their optimum temperatures were 65 °C and 60 °C, respectively, which are ideal for the baking industry. Processing baked foods with enzymes retarded their staling without any gumminess or unfavorably affected the palatability traits of the baked goods. In the food and biobased industries, plant-isolated β-amylases are preferred due to their higher specificity and safety. However, their utility is limited, as the enzymes obtained from cereal grains

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are human staple foods. In addition, their thermostability properties are not ideal for application as an intermediary temperature-stable enzyme. The identification of fresh plant-based enzymes that are highly stable and prove to be a potential alternative to traditional plant β-amylases still needs extensive study. The T50 for ramie leaf β-amylase is 66 °C, which is superior to that of β-amylase from wheat flour (50 ° C), soybean (63.2 °C), and barley (56.8 °C). Additionally, the thermal inactivation of β-amylase from peanut at 65 °C resulted in first-order kinetics with a rate constant (k) of 0.0126 min-1 and t½ 55 min. Thus, ramie leaf and peanut β-amylases have superb thermostability compared to other reported plant β-amylases. The presence of additives enhances the thermostability of wheat β-amylase [45]. In continuation, β-amylase from ungerminated and germinated fenugreek has also been purified in our laboratory [46, 47] and further immobilized onto nanomaterials for a better kinetic approach [48–50]. Examining the effect of partial β-amylolysis on the retrogradation of rice starch by [51], inferred the feasible use of β-amylase in rice product preparations, with an extended shelf life. The results show that partial hydrolysis using β-amylase slows amylopectin retrogradation by shortening the branch chain length of amylopectin. Furthermore, the inhibitory effect of retrogradation is attenuated by maltose produced in β-amylolysis.

1.2.1.3

Glucoamylases

These enzymes, commonly known as amyloglucosidase or sugar-making amylase, distinguish α-1,4 bonds and, to a lesser extent, α-1,6 bonds from the nonreducing end, similar to pullulanase type II [52]. They have been isolated from various microorganisms and fungal and plant sources. The most common sources are Aspergillus niger, A. japonicas, Clostridium thermosaccharolyticum, Fusarium solani, maize leaves, etc. The major commercial application of this enzyme is the production of high-glucose syrups from starch. This is achieved by using many enzymes, which may lead to unfavorable side reactions, such as back reaction and production of isomaltose and maltose, and concomitantly may decrease the final product. Glucoamylases are usually acidic in nature and sensitive to temperature. They are used for saccharification of dextrins to glucose-rich syrups in the industry [36, 53]. The syrups find application in fermentation, production of crystalline glucose, and as a starting material for fructose syrups [54, 55]. In addition, Glucoamylase has been exploited for the production of low-calorie beer in breweries, wherein maltodextrin in malted barley is broken down to simple sugars, which thereby undergo complete fermentation by brewer’s yeast. Glucoamylase stands second to proteases in worldwide sales and distribution, amidst industrial enzymes [56].

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1.2.1.4

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α-Glucosidases

α-Glucosidases also belong to the exo-acting enzyme family that hydrolyzes starch components. Amylose, amylopectin, and oligosaccharides with maltose from the nonreducing end and produce α-glucose as opposed to β-glucose [57]. The enzyme favors the transglycosylation reaction when the maltose concentration is high enough, and therefore, maltose is transformed to isomaltose (two glucose with 1, 6 linkages). Furthermore, isomaltose is glycosylated to produce isomaltotriose. This enzyme is also known to glycosylate branched oligosaccharides such as panose and isopanose. These branched chain sugar compounds are generally well known as isomalto-oligosaccharides (IMOs), which find application as prebiotic fibers in countries such as Japan and China [52].

1.2.2

Debranching Enzymes

The third category of starch-converting enzymes is the debranching enzymes, viz. isoamylase (EC 3.2.1.68) and pullulanase type I (EC 3.2.1.41), which exclusively cleave α-1,6 glycosidic bonds. Pullulanases and isoamylases differ mainly in their ability to hydrolyze pullulan by pullulanase, a polysaccharide with a repeating unit of maltotriose, which is 1-6 linked [58]. These enzymes solely cleave amylopectin, thus leaving long linear polysaccharides. The literature reports several other enzymes with pullulanase-type activity, i.e., hydrolyzing both α 1,4 and α 1,6 linkages. These belong to the type II pullulanase family and are called α-amylasepullulanase or amylopullulanase. The principal hydrolysis products are maltose and maltotriose. Another special enzyme belonging to this group of pullulanases is neopullulanase, which, apart from hydrolysis, can also perform transglycosylation with the generation of new α-1,4 or α-1,6 glycosidic bonds [59]. Let us discuss these debranching enzymes in brief.

1.2.2.1

Pullulanases

Pullulanases (EC 3.2.1.41) have gained attention recently in starch-based industries, especially those targeted for the production of glucose. Alternative names of pullulanases are α-dextrin 6-glucanohydrolase, amylopectin 6-glucanohydrolase, pullulan 6-glucanohydrolase, and limit dextrinase. Pullulanase, obtained from microbial sources, is a vital debranching enzyme widely utilized for hydrolyzing the α-1,6 glucosidic linkages in starch, amylopectin, pullulan (the repeated units contain three glucose residues with α-1,4 bonds that are linked together by one α-1,6 bond), and related oligosaccharides. This process allows a fully-fledged transformation of the branched polysaccharides into small sugars (fermentable) during saccharification. Industrial production of glucose involves two consecutive enzymatic steps:

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liquefaction, carried out after gelatinization by the application of α-amylase, and saccharification, which results in further conversion of maltodextrins into glucose. During the saccharification process, pullulanase increases the final glucose content along with a lower amount of glucoamylase. Thus, the retrogression reaction that involves resynthesis of saccharides from glucose is arrested. Five categories of pullulanase enzymes have been reported and are currently attributed to the features of the substrate and products generated: (i) pullulanase type I, (ii) amylopullulanase, (iii) neopullulanase, (iv) isopullulanase, and (v) pullulan hydrolase type III. Type I pullulanases cleave α-1,6 bonds in pullulan and branched polysaccharides. Type II pullulanases (amylopullulanases) cleave α-1,4 or α-1,6 linkages and are used mainly in the starch processing industry. Neither type of pullulanase reacts with cyclodextrins [60–62]. Neopullulanases (pullulan hydrolase type I) and isopullulanases (pullulan hydrolase type II) are reported to break α-1,4 bonds and act on cyclodextrins. They do not act on starch. Pullulan hydrolase type III stands out from other pullulan hydrolyzing enzymes in breaking α-1,4 and α-1,6 bonds in pullulan and producing a mixture of maltotriose, panose, maltose, and glucose [63]. They are also known for analyzing starch, amylose, and amylopectin and generating maltose and maltotriose [53]. Pullulanase is also utilized in the production of high-amylose starches, which have vast market demand [64]. Interestingly, high-amylose starches are transformed into “resistant starch,” which has nutritional advantages [65]. In contrast to normal starch, resistant starch is not easy to digest in the small intestine. Thus, it is fermented in the large intestine by bacteria present in the gut, producing short-chain fatty acids such as butyrates that are beneficial for colon health. Corrugated board and paper are produced by high-amylose starches, which also find application in adhesive production [66]. A marked increase in evidence points out that many of the chronic diseases in developed countries could be toned down by alterations in diet. The people of the United States rely mainly on white bread, cakes, noodles, etc., a large portion of which is easily digestible starch. This is bothersome because rapidly digesting starch is known to cause chronic disease in humans and cattle. Owing to these problems, starches resistant to digestive enzymes are experiencing a growing research highlight. Such starches, termed resistant starches [67], have been extensively reviewed in the context of their health and functional properties as a food ingredient [68] and their role in gut health, potentially through the production of butyrate [69]. The proposed daily dose of resistant starch by Americans is ~5 g/day, much lower than the prescribed intake of 6 g of resistant starch per meal recommended for health benefits [70]. Resistant starch-rich powdered ingredients are produced in three ways: physical, chemical, and enzymatic processes. The enzymatic process focuses on debranching the α-1,6 bonds in the starch polysaccharide amylopectin by pullulanase and isoamylase, which results in reorganization of the structure and eventually in the retrogradation process [71]. Other enzymes, such as α- and β-amylase, hydrolyze amorphous regions of the starch and leave a tightly packed crystalline structure. Therefore, the objective of using debranching enzymes is to hydrolyze amylopectin branch chains and provide more linear parts [72]. Subsequently, retrogradation will produce increased levels of resistant starch with double helical structures stabilized by hydrogen bonding.

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Resistant starch is a marvelous prebiotic with many additional health benefits. It also has technologically innovative effects on food products as a functional component. Nevertheless, processing conditions, especially heat treatment, can mitigate its content. Enzymatic debranching is more promising due to the production of resistant starch at high concentrations. It also emerges as an essential step when used with other methods, e.g., heat and cool cycles. There is a demand for a balanced environment for better enzyme activity and subsequent heat treatments. Let us discuss a few industrial applications of pullulanase. Commercial Practice of Pullulanase Hydrolysis of Starch Currently, starch hydrolysates on the market are enzyme-transformed products of higher dextrose equivalents (DEs). DE is a measure of reducing sugar on a dry basis, pure dextrose (glucose) being DE 100 and starch being close to zero. The production of glucose, maltose, and fructose from starch by enzymatic processes is used as food sweeteners, representing an important expansion area of industrial enzymes. The fundamental application of pullulanase is in starch saccharification, but its most valuable industrial application is the production of high-glucose (30%–50% glucose; 30%–40% maltose) or high-maltose (30%–50% maltose; 6%–10% glucose) syrups [12, 73]. In the saccharification process, glucoamylases or β-amylases are also used along with pullulanases [74, 75]. Generation of High-Maltose- and Fructose-Rich Corn Syrup Maltose (a reducing sugar)-rich syrups are known for their mild sweetness, low solution viscosity, low hygroscopicity, good thermostability, lack of color formation, etc., which account for its pharmaceutical distribution. It is perfectly adapted for various applications in food processing, one being the manufacturing of high-quality candy and ice cream [76]. In the past few years, there has been an elevated demand for maltose in pure form in the pharmaceutical industry. The disbursement of maltose in India and Southeast Asia has appreciated drastically, being the only source of carbon for the production of the Diphtheria Pertussis Tetanus (DPT) vaccine to immune the whole population [35]. Maltose can replace D-glucose for intravenous feeding, where maltose is injected at higher concentrations without shooting blood glucose levels. Perhaps pure maltose could also be used as a starting material for the production of maltitol and crystalline maltitol [77]. High-glucose syrup is used as a source of carbon in fermentation and feeds to make crystalline glucose and high-fructose syrups [73]. High-fructose syrup has been produced by the treatment of glucose syrup (DE 95-96) with immobilized glucose isomerase [12]. Fructose-rich syrup from corn is a high-quality caloric sweetener. Fructose-rich syrup with a high DE value is required in the production of crystalline glucose. It finds wide application in commodity description. They are inexpensive and low calorific to sucrose (but 1.2–1.8 times sweeter than the same). Similar to high-maltose syrup, 90% of fructose-rich syrups are used in food for diabetics because they can be easily metabolized without insulin.

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Detergent Some alkaline pullulanases provide a good alternative as additives in dishwashing and laundry detergents for the removal of food starches under alkaline conditions [78, 79]. Amylopullulanase used in combination with alkaline α-amylase improves the efficacy of these alkaline debranching enzymes in washing since a single amylopullulanase can catalyze both reactions, viz., debranching (α-1,6 hydrolytic) and liquefying (α-1,4 hydrolytic) [80]. Production of Cyclodextrins Pullulanase also boosts the yield of cyclodextrins by the reaction of cyclodextrin glucantransferase with gelatinized starches and maltodextrin syrups in the presence of cyclodextrin complexing agents. Cyclodextrins have a full spectrum of applications in complexing materials in foods, pharmaceuticals, environmental engineering, chemical industries, agriculture, etc. [81]. Pullulanases also claim production of low-calorie beer for health-conscious people [82] and in the baking industry as the anti-staling agent to improve qualities such as texture, flavor, and volume of bakery products. It is also used in dental plaque/mouthwashes [83].

1.2.2.2

Isoamylases

Isoamylases (EC 3.2.1.68, glycogen-6-glucanohydrolase) can hydrolyze α-1,6-glucosidic branched linkages in amylopectin to produce amylose and oligosaccharides. Glycogen is known to be completely debranched by isoamylases only. Isoamylases act via endo-mechanisms, and the branch linkages located in the interior parts of the amylopectin molecule are hydrolyzed to give linear maltodextrins, thus retaining the α-configuration. This enzyme was first isolated from broad beans and classified as an “R-enzyme” [84]. The enzyme cleaves some of the α-(1 → 6) linkages of amylopectin, amylopectin β-amylase limits dextrin, and different branched α-amylases limit dextrins. It was unable to act on glycogen or its β-amylase limit dextrin. Isoamylase from P. amylodermosa was crystallized, and its specificity was studied [85]. Like pullulanases, isoamylases also do not remove single-branch glucosyl units from branched maltodextrins and act extremely slowly on maltosyl units attached by α-(1 → 6) linkages. The distinction between the two, i.e., isoamylase and pullulanase is that the former is able to cleave the α-(1 → 6) linkages of amylopectin and glycogen completely and at rates of 7× and 124×, respectively, the rates of hydrolysis of these substrates by pullulanase [86]. The rate of hydrolysis of the α-(1 → 6) branch linkages of short chains is 10% or less than the rate of hydrolysis of the branch linkages of the longer chains. Pseudomonas isoamylase hydrolyzes the α-(1 → 6) branched linkages in maltotriosyl chains more rapidly. The hydrolysis of the α (1 → 6) linkage in pullalan by Pseudomonas isoamylase is extremely slow. Unlike pullulanase, which can hydrolyze only outer branch linkages, Pseudomonas isoamylase is known for catalyzing the hydrolysis of both the inner and outer

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branch linkages of amylopectin and glycogen. Pseudomonas isoamylase hydrolyzes all of the branch linkages, whereas pullulanase hydrolyzes only a few, with glycogen as a substrate. Thus, they appear to be true debranching enzymes for starch and glycogen, even though they do not release single glucosyl units but release maltosyl units, which occurs too slowly. Their distinctiveness for longer chain lengths and its ability to hydrolyze the interior α-(1 → 6) branch linkages observed that the active site of Pseudomonas isoamylase binds two chains (the A- and B- or the A- and C-chains). James et al. [87] cloned a starch debranching isoamylase obtained from sugary l (sul) maize, and this was found to have 32% sequence homology with the Pseudomonas enzyme. Many more isoamylases or starch debranching enzymes have been isolated from rye [88], sweet potato [89], Rhizopus oryzae [90], Dyella sp. strain [91], Corallococcus sp. strain EGB [92], etc.

1.2.3

Transferase

Finally, we have transferases that cleave the α-1,4 glycosidic bond of the donor molecule and carry part of the donor to a glycosidic acceptor, and a new glycosidic bond is generated. For instance, enzymes such as amylomaltase (EC 2.4.1.25) and cyclodextrin glycosyltransferase (EC 2.4.1.19) form a new α-1,4 glycosidic bond, while 1,4-alpha-glucan branching enzyme (EC 2.4.1.18) forms a new α-1,6 glycosidic bond. The low hydrolytic activity of cyclodextrin glycosyltransferases results in the formation of cyclic oligosaccharides resulting in 6, 7, or 8 glucose residues. They also form greatly branched, high molecular weight dextrins, and cyclodextrin glycosyltransferase limits dextrins. Cyclodextrins are released as a result of an intramolecular transglycosylation reaction leading to enzyme cleavage of the α-1,4 glycosidic bond and simultaneous joining of the reducing end to the nonreducing end. Amylomaltases resemble cyclodextrin glycosyltransferases in terms of the type of enzymatic reaction. The fundamental difference between the two is that amylomaltase carries out a transglycosylation reaction leading to a linear product, while cyclodextrin glycosyltransferase generates a cyclic product. Amylomaltases in microorganisms aid them in utilizing maltose or glycogen hydrolysis [93].

1.2.3.1

Cyclodextrin Glucanotransferases (CGTAs, EC 2.4.1.19)

These enzymes originating from bacteria and archaea are active at high temperatures. They are well known for catalyzing cyclization, coupling, disproportion, and hydrolysis reactions. β-CDs have fundamental applications in the bioremediation of chronically infected soils by increasing water solubility and decreasing the toxicity of fungicides, pushing them toward biodegradation. The main employment of this enzyme is the generation of industrial cyclodextrins. α-, β-, and γ-Cyclodextrins are believed to be gluco-oligosaccharides with cyclic α-1, 4 bonds and are called

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sacchardingers. The sequence of their glucose residues creates a hydrophobic inner part and hydrophilic outer part. Consequently, they can merge with compounds hydrophobic in nature and create a new physical and chemical properties. These compounds function as antiseptic factors [52]. They enable the removal of cholesterol from eggs and lactic products, phenolic compounds from fruits and vegetable juices and the detection of aflatoxin in food samples. In the food sector, α-, β-, and γ-cyclodextrins have broad applications in stabilizing aromatic and sensitive compounds, solubilizing hydrophobic compounds in water, and eliminating undesirable odors and tastes [94–96]. α-Cyclodextrins are considered diet fiber, which is indigestible. It is generated from the action of cyclodextrin glucanotransferase enzymes on liquefied starch. CGTAs also prevent staling of bread. Because of the structural similarity of this enzyme with an anti-staling enzyme, Novomyl, the structural change in the latter converts it to CGTAs. Reports from the literature have shown that mutant CGATs drastically improve the quality of bakery production [97, 98].

1.3

Conclusion and Future Perspectives

Thus far, we have discussed starch refining enzymes used in the modification and conversion of starch and the production of a variety of compounds. Enzyme technology is a promising tool for modifying starch in starch-based foods such as baking products, resistant starch production, and limiting dextrins. Because enzymes acting on carbohydrates can react in a substrate-specific manner toward the target starch in the complex network of food items. Microbial enzymes compromise their safety issues. Therefore, plant-derived enzymes are considered safe. However, their use is limited due to thermostability during starch processing. Thus, new plant-based enzymes need to be further investigated. As discussed in this chapter, retrogradation of starch-based food can be treated by using a combination of enzymes. The mechanistic enzyme reaction needs to be further explored for industrial usage. Despite extensive research and production of various compounds, all starch processing enzymes have not been thoroughly studied. Therefore, much more inspection in the area of recognition and application of these enzymes for novel products is being exercised so that the production of new refined and practical starch can be witnessed in the future.

References 1. Santana Á, Meireles M. New starches are the trend for industry applications: a review. Food Public Health. 2014;4:229–41. 2. Srichuwong S, Sunarti TC, Mishima T, Isono N, Hisamatsu M. Starches from different botanical sources I: contribution of amylopectin fine structure to thermal properties and enzyme digestibility. Carbohydr Polym. 2005;60:529–38.

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3. Yoo S-H, Jane J-l. Structural and physical characteristics of waxy and other wheat starches. Carbohydr Polym. 2002;49:297–305. 4. Syahariza Z, Li E, Hasjim J. Extraction and dissolution of starch from rice and sorghum grains for accurate structural analysis. Carbohydr Polym. 2010;82:14–20. 5. van der Maarel MJ, Leemhuis H. Starch modification with microbial alpha-glucanotransferase enzymes. Carbohydr Polym. 2013;93:116–21. 6. Daudt RM, Külkamp-Guerreiro IC, Cladera-Olivera F, Thys RCS, Marczak LDF. Determination of properties of pinhão starch: analysis of its applicability as pharmaceutical excipient. Ind Crop Prod. 2014;52:420–9. 7. Zhu F, Wang Y-J. Characterization of modified high-amylose maize starch-α-naphthol complexes and their influence on rheological properties of wheat starch. Food Chem. 2013;138: 256–62. 8. Woods L, Swinton S. Enzymes in the starch and sugar industries. In: Enzymes in food processing. Springer; 1995. p. 250–67. 9. Albano KM, Franco CM, Telis VR. Rheological behavior of Peruvian carrot starch gels as affected by temperature and concentration. Food Hydrocoll. 2014;40:30–43. 10. Rath SS, Sahoo H. A review on the application of starch as depressant in iron ore flotation. Miner Process Extr Metall Rev. 2020:1–14. 11. Neelam K, Vijay S, Lalit S. Various techniques for the modification of starch and the applications of its derivatives. Int Res J Pharm. 2012;3:25–31. 12. Van Der Maarel MJ, Van der Veen B, Uitdehaag JC, Leemhuis H, Dijkhuizen L. Properties and applications of starch-converting enzymes of the α-amylase family. J Biotechnol. 2002;94:137– 55. 13. Pandey A, Nigam P, Soccol CR, Soccol VT, Singh D, Mohan R. Advances in microbial amylases. Biotechnol Appl Biochem. 2000;31:135–52. 14. Diderichsen B, Christiansen L. Cloning of a maltogenic alpha-amylase from Bacillus stearothermophilus. FEMS Microbiol Lett. 1988;56:53–9. 15. Robyt JF, Ackerman RJ. Isolation, purification, and characterization of a maltotetraoseproducing amylase from Pseudomonas stutzeri. Arch Biochem Biophys. 1971;145:105–14. 16. Momma M. Cloning and sequencing of the maltohexaose-producing amylase gene of Klebsiella pneumoniae. Biosci Biotechnol Biochem. 2000;64:428–31. 17. Kumari A, Singh K, Kayastha M. α-Amylase: general properties, mechanism and biotechnological applications—a review. Curr Biotechnol. 2012;1:98–107. 18. Kammoun R, Naili B, Bejar S. Application of a statistical design to the optimization of parameters and culture medium for α-amylase production by Aspergillus oryzae CBS 819.72 grown on gruel (wheat grinding by-product). Bioresour Technol. 2008;99:5602–9. 19. Ochiai A, Sugai H, Harada K, Tanaka S, Ishiyama Y, Ito K, Tanaka T, Uchiumi T, Taniguchi M, Mitsui T. Crystal structure of α-amylase from Oryza sativa: molecular insights into enzyme activity and thermostability. Biosci Biotechnol Biochem. 2014;78:989–97. 20. Janeček Š. How many conserved sequence regions are there in the α-amylase family. Biologia. 2002;57:29–41. 21. Savchenko A, Vieille C, Kang S, Zeikus JG. Pyrococcus furiosus α-amylase is stabilized by calcium and zinc. Biochemistry. 2002;41:6193–201. 22. Prakash O, Jaiswal N. α-Amylase: an ideal representative of thermostable enzymes. Appl Biochem Biotechnol. 2010;160:2401–14. 23. Tiwari S, Srivastava R, Singh C, Shukla K, Singh R, Singh P, Singh R, Singh N, Sharma R. Amylases: an overview with special reference to alpha amylase. J Global Biosci. 2015;4: 1886–901. 24. Singh K, Kayastha AM. α-Amylase from wheat (Triticum aestivum) seeds: its purification, biochemical attributes and active site studies. Food Chem. 2014;162:1–9. 25. Tripathi P, Dwevedi A, Kayastha AM. Purification and partial characterization of α-amylase from soybean (Glycine max). Orient Pharm Exp Med. 2004;4:227–34.

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26. Kumari A, Singh VK, Fitter J, Polen T, Kayastha AM. α-Amylase from germinating soybean (Glycine max) seeds–purification, characterization and sequential similarity of conserved and catalytic amino acid residues. Phytochemistry. 2010;71:1657–66. 27. Tripathi P, Leggio LL, Mansfeld J, Ulbrich-Hofmann R, Kayastha AM. α-amylase from mung beans (Vigna radiata)–correlation of biochemical properties and tertiary structure by homology modelling. Phytochemistry. 2007;68:1623–31. 28. Singh K, Kayastha AM. Optimal immobilization of α-amylase from wheat (Triticum aestivum) onto DEAE-cellulose using response surface methodology and its characterization. J Mol Catal B Enzym. 2014;104:75–81. 29. Singh K, Srivastava G, Talat M, Srivastava ON, Kayastha AM. α-Amylase immobilization onto functionalized graphene nanosheets as scaffolds: its characterization, kinetics and potential applications in starch based industries. Biochem Biophys Rep. 2015;3:18–25. 30. Tripathi P, Kumari A, Rath P, Kayastha AM. Immobilization of α-amylase from mung beans (Vigna radiata) on Amberlite MB 150 and chitosan beads: a comparative study. J Mol Catal B Enzym. 2007;49:69–74. 31. Das R, Mishra H, Srivastava A, Kayastha AM. Covalent immobilization of β-amylase onto functionalized molybdenum sulfide nanosheets, its kinetics and stability studies: a gateway to boost enzyme application. Chem Eng J. 2017;328:215–27. 32. Das R, Talat M, Srivastava O, Kayastha AM. Covalent immobilization of peanut β-amylase for producing industrial nano-biocatalysts: a comparative study of kinetics, stability and reusability of the immobilized enzyme. Food Chem. 2018;245:488–99. 33. Li X, Yu H-Y. Extracellular production of beta-amylase by a halophilic isolate, Halobacillus sp. LY9. J Ind Microbiol Biotechnol. 2011;38:1837–43. 34. Das R, Kayastha AM. Enzymatic hydrolysis of native granular starches by a new β-amylase from peanut (Arachis hypogaea). Food Chem. 2019;276:583–90. 35. Das R, Ranjan R, Sinha N, Kayastha AM. Comparative characterization of peanut β-amylase immobilization onto graphene oxide and graphene oxide carbon nanotubes by solid-state NMR. J Phys Chem C. 2018;122:19259–65. 36. Tucker GA, Woods L. Enzymes in food processing. Springer Science & Business Media; 1995. 37. Das R, Kayastha AM. β-Amylase: general properties, mechanism and panorama of applications by immobilization on nano-structures. In: Biocatalysis. Cham: Springer; 2019. p. 17–38. 38. Das R, Kayastha AM. An antioxidant rich novel β-amylase from peanuts (arachis hypogaea): its purification, biochemical characterization and potential applications. Int J Biol Macromol. 2018;111:148–57. 39. Nguyen DHD, Tran PL, Ha HS, Lee JS, Hong WS, Le QT, Oh BC, Park SH. Presence of β-amylase in ramie leaf and its anti-staling effect on rice cake. Food Sci Biotechnol. 2015;24: 37–40. 40. Lundgard R, Svensson B. The four major forms of barley β-amylase. Purification, characterization and structural relationship. Carlsberg Res Commun. 1987;52:313. 41. Silano V, Bolognesi C, Castle L, Cravedi JP, Fowler P, Franz R, Grob K, Gürtler R, Husøy T. Safety evaluation of a β-amylase food enzyme obtained from wheat (Triticum spp.). EFSA J. 2017;15:e04754. 42. Tester RF. a chemical carrier based on a beta-limit dextrin. Google Patents. 2004; 43. Qi X, Tester RF. Bioadhesive properties of β-limit dextrin. J Pharm Pharm Sci. 2011;14:60–6. 44. He L, Park S-H, Dang NDH, Duong HX, Duong TPC, Tran PL, Park J-T, Ni L, Park K-H. Characterization and thermal inactivation kinetics of highly thermostable ramie leaf β-amylase. Enzyme Microb Technol. 2017;101:17–23. 45. Daba T, Kojima K, Inouye K. Characterization and solvent engineering of wheat β-amylase for enhancing its activity and stability. Enzyme Microb Technol. 2012;51:245–51. 46. Srivastava G, Kayastha AM. β-Amylase from starchless seeds of Trigonella foenum-graecum and its localization in germinating seeds. PLoS One. 2014;9:e88697. 47. Agrawal DC, Dwevedi A, Kayastha AM. Biochemical and thermodynamic characterization of de novo synthesized β-amylase from fenugreek. Int J Biol Macromol. 2019;130:786–97.

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48. Agrawal DC, Yadav A, Kesarwani R, Srivastava O, Kayastha AM. Immobilization of fenugreek β-amylase onto functionalized graphene quantum dots (GQDs) using Box-Behnken design: its biochemical, thermodynamic and kinetic studies. Int J Biol Macromol. 2020;144: 170–82. 49. Srivastava G, Singh K, Talat M, Srivastava ON, Kayastha AM. Functionalized graphene sheets as immobilization matrix for fenugreek β-amylase: enzyme kinetics and stability studies. PLoS One. 2014;9:e113408. 50. Agrawal DC, Yadav A, Singh VK, Srivastava A, Kayastha AM. Immobilization of fenugreek β-amylase onto functionalized tungsten disulfide nanoparticles using response surface methodology: its characterization and interaction with maltose and sucrose. Colloids Surf B: Biointerfaces. 2020;185:110600. 51. Yao Y, Zhang J, Ding X. Partial β-amylolysis retards starch retrogradation in rice products. J Agric Food Chem. 2003;51:4066–71. 52. Whitehurst RJ, Van Oort M. Enzymes in food technology. Wiley Online Library; 2010. 53. Hii SL, Tan JS, Ling TC, Ariff AB. Pullulanase: role in starch hydrolysis and potential industrial applications. Enzyme Res. 2012;2012 54. Bhatti HN, Rashid MH, Nawaz R, Asgher M, Perveen R, Jabbar A. Purification and characterization of a novel glucoamylase from Fusarium solani. Food Chem. 2007;103:338–43. 55. Marlida Y, Saari N, Hassan Z, Radu S, Bakar J. Purification and characterization of sago starchdegrading glucoamylase from Acremonium sp. endophytic fungus. Food Chem. 2000;71:221– 7. 56. Negi S, Gupta S, Banerjee R. Extraction and purification of glucoamylase and protease produced by Aspergillus awamori in a single-stage fermentation. Food Technol Biotechnol. 2011;49:310–5. 57. Okuyama M, Saburi W, Mori H, Kimura A. α-Glucosidases and α-1, 4-glucan lyases: structures, functions, and physiological actions. Cell Mol Life Sci. 2016;73:2727–51. 58. Israilides C, Smith A, Scanlon B, Barnett C. Pullulan from agro-industrial wastes. Biotechnol Genet Eng Rev. 1999;16:309–24. 59. Nisha M, Satyanarayana T. Recombinant bacterial amylopullulanases: developments and perspectives. Bioengineered. 2013;4:388–400. 60. Roy A, Messaoud EB, Bejar S. Isolation and purification of an acidic pullulanase type II from newly isolated Bacillus sp. US149. Enzyme Microb Technol. 2003;33:720–4. 61. Messaoud EB, Ammar YB, Mellouli L, Bejar S. Thermostable pullulanase type I from new isolated Bacillus thermoleovorans US105: cloning, sequencing and expression of the gene in E. coli. Enzym Microb Technol. 2002;31:827–32. 62. Duffner F, Bertoldo C, Andersen JT, Wagner K, Antranikian G. A new thermoactive pullulanase from Desulfurococcus mucosus: cloning, sequencing, purification, and characterization of the recombinant enzyme after expression in Bacillus subtilis. J Bacteriol. 2000;182: 6331–8. 63. Niehaus F, Bertoldo C, Kähler M, Antranikian G. Extremophiles as a source of novel enzymes for industrial application. Appl Microbiol Biotechnol. 1999;51:711–29. 64. Vorwerg W, Radosta S, Leibnitz E. Study of a preparative-scale process for the production of amylose. Carbohydr Polym. 2002;47:181–9. 65. Bird AR, Brown IL, Topping DL. Starches, resistant starches, the gut microflora and human health. Curr Issues Intest Microbiol. 2000;1:25–37. 66. Jobling S. Improving starch for food and industrial applications. Curr Opin Plant Biol. 2004;7: 210–8. 67. Englyst HN, Kingman SM, Cummings J. Classification and measurement of nutritionally important starch fractions. Eur J Clin Nutr. 1992;46:S33–50. 68. Fuentes-Zaragoza E, Sánchez-Zapata E, Sendra E, Sayas E, Navarro C, Fernández-López J, Pérez-Alvarez JA. Resistant starch as prebiotic: a review. Starch - Stärke. 2011;63:406–15. 69. Brouns F, Kettlitz B, Arrigoni E. Resistant starch and “the butyrate revolution”. Trends Food Sci Technol. 2002;13:251–61.

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70. Murphy MM, Douglass JS, Birkett A. Resistant starch intakes in the United States. J Am Diet Assoc. 2008;108:67–78. 71. Reddy CK, Suriya M, Haripriya S. Physico-chemical and functional properties of resistant starch prepared from red kidney beans (Phaseolus vulgaris. L) starch by enzymatic method. Carbohydr Polym. 2013;95:220–6. 72. Cai L, Shi Y-C. Structure and digestibility of crystalline short-chain amylose from debranched waxy wheat, waxy maize, and waxy potato starches. Carbohydr Polym. 2010;79:1117–23. 73. Gomes I, Gomes J, Steiner W. Highly thermostable amylase and pullulanase of the extreme thermophilic eubacterium Rhodothermus marinus: production and partial characterization. Bioresour Technol. 2003;90:207–14. 74. Haki G, Rakshit S. Developments in industrially important thermostable enzymes: a review. Bioresour Technol. 2003;89:17–34. 75. Chaplin M, Production of syrups containing maltose; 2002. 76. Shaw J-F, Sheu J-R. Production of high-maltose syrup and high-protein flour from rice by an enzymatic method. Biosci Biotechnol Biochem. 1992;56:1071–3. 77. Shirke S, Ludescher RD. Dynamic site heterogeneity in amorphous maltose and maltitol from spectral heterogeneity in erythrosin B phosphorescence. Carbohydr Res. 2005;340:2661–9. 78. Hatada Y, Saito K, Hagihara H, Ozaki K, Ito S. Nucleotide and deduced amino acid sequences of an alkaline pullulanase from the alkaliphilic bacterium Bacillus sp. KSM-1876. Biochim Biophys Acta. 2001;1545:367–71. 79. Schallmey M, Singh A, Ward OP. Developments in the use of Bacillus species for industrial production. Can J Microbiol. 2004;50:1–17. 80. Ara K, Saeki K, Igarashi K, Takaiwa M, Uemura T, Hagihara H, Kawai S, Ito S. Purification and characterization of an alkaline amylopullulanase with both α-1, 4 and α-1, 6 hydrolytic activity from alkalophilic Bacillus sp. KSM-1378. Biochim Biophys Acta. 1995;1243:315–24. 81. Kim Y-K, Robyt JF. Enzyme modification of starch granules: formation and retention of cyclomaltodextrins inside starch granules by reaction of cyclomaltodextrin glucanosyltransferase with solid granules. Carbohydr Res. 2000;328:509–15. 82. Olsen HS. Enzymes at work: a concise guide to industrial enzymes and their uses. Novo Nordisk A/S; 2000. 83. Marotta M, Martino A, De Rosa A, Farina E, Cartenı M, De Rosa M. Degradation of dental plaque glucans and prevention of glucan formation using commercial enzymes. Process Biochem. 2002;38:101–8. 84. Zhu Z-P, Hylton CM, Rössner U, Smith AM. Characterization of starch-debranching enzymes in pea embryos. Plant Physiol. 1998;118:581–90. 85. Robyt JF. Enzymes and their action on starch. In: Starch. Elsevier; 2009. p. 237–92. 86. Fujita N, Toyosawa Y, Utsumi Y, Higuchi T, Hanashiro I, Ikegami A, Akuzawa S, Yoshida M, Mori A, Inomata K. Characterization of pullulanase (PUL)-deficient mutants of rice (Oryza sativa L.) and the function of PUL on starch biosynthesis in the developing rice endosperm. J Exp Bot. 2009;60:1009–23. 87. James MG, Robertson DS, Myers AM. Characterization of the maize gene sugary1, a determinant of starch composition in kernels. Plant Cell. 1995;7:417–29. 88. Zheng K, Xu J, Jiang Q, Laroche A, Wei Y, Zheng Y, Lu Z. Isolation and characterization of an isoamylase gene from rye. Crop J. 2013;1:127–33. 89. Kim S-H, Hamada T, Otani M, Shimada T. Cloning and characterization of sweetpotato isoamylase gene (IbIsa1) isolated from tuberous root. Breed Sci. 2005;55:453–8. 90. Ghosh B, Lahiri D, Nag M, Dash S, Ray RR. Bio characterization of purified isoamylase from Rhizopus oryzae. Prep Biochem Biotechnol. 2020;50:453–9. 91. Silano V, Barat Baviera JM, Bolognesi C, Cocconcelli PS, Crebelli R, Gott DM, Grob K, Lambré C, Lampi E. Safety evaluation of the food enzyme isoamylase from a Dyella sp. strain. EFSA J. 2020;18:e06250. 92. Li Z, Ji K, Zhou J, Ye X, Wang T, Luo X, Huang Y, Cao H, Cui Z, Kong Y. A debranching enzyme IsoM of Corallococcus sp. strain EGB with potential in starch processing. Int J Biol Macromol. 2017;105:1300–9.

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93. Takaha T, Smith SM. The functions of 4-α-glucanotransferases and their use for the production of cyclic glucans. Biotechnol Genet Eng Rev. 1999;16:257–80. 94. Astray G, Gonzalez-Barreiro C, Mejuto JC, Rial-Otero R, Simal-Gandara J. A review on the use of cyclodextrins in foods. Food Hydrocoll. 2009;23:1631–40. 95. Szente L, Szejtli J. Cyclodextrins as food ingredients. Trends Food Sci Technol. 2004;15:137– 42. 96. Reineccius T, Reineccius G, Peppard T. Flavor release from cyclodextrin complexes: comparison of alpha, beta, and gamma types. J Food Sci. 2003;68:1234–9. 97. Shim J-H, Seo N-S, Roh S-A, Kim J-W, Cha H, Park K-H. Improved bread-baking process using Saccharomyces cerevisiae displayed with engineered cyclodextrin glucanotransferase. J Agric Food Chem. 2007;55:4735–40. 98. Kelly RM, Dijkhuizen L, Leemhuis H. The evolution of cyclodextrin glucanotransferase product specificity. Appl Microbiol Biotechnol. 2009;84:119–33.

Chapter 2

Classification and Enzyme Properties of Starch Debranching Enzymes Wei Xia and Jing Wu

Abstract The α-1,6 glycosidic linkages in polysaccharides, including amylose, straight-chain starch, pullulan, and glycogen, are hydrolyzed by starch debranching enzymes (SDBEs). The production of syrups, resistant starches, and cyclodextrins all requires the use of SDBEs. Since SDBEs and other starch-acting hydrolases synergistic catalysis can significantly increase raw material usage and productivity in starch processing processes, including saccharification and modification, they have drawn much attention from researchers in recent years. The substrate specificities of the two major members of SDBEs, pullulanase, and isoamylases, are very different. Pullulanase generally requires the presence of at least two α-1,4-linked glucose units on two sugar chains linked by α-1,6 bonds, whereas isoamylases require at least three α-1,4-linked glucose units. Keywords Enzyme classification · Enzyme properties · Substrate specificity · Original sources

2.1

Introduction

Starch is the second most renewable resource in nature and is synthesized mainly by photosynthetic plants, green algae, and cyanobacteria as a carbohydrate and energy store [1, 2]. It is quite abundant in staple foods such as potatoes, wheat, maize, rice, and cassava and has a crucial role in human society, being the most common food component in the human diet [3–6] and in various industries concerning food, chemicals, and pharmaceuticals [7–9]. In terms of chemical properties, starch is a macromolecular biomolecule composed of hundreds of glucose units that are linked W. Xia · J. Wu (✉) State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, 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. 2023 J. Wu, W. Xia (eds.), Industrial Starch Debranching Enzymes, https://doi.org/10.1007/978-981-19-7026-9_2

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by major α-1,4-D-glucosidic bonds (~94%) and lesser α-1,6-D-glucosidic bonds (~6%) formed at the C1 oxygen atom. The general chemical composition of starch is (C6H10O5)n. Starch of plant origin contains two types of molecules. (1) straightchain starch, consisting of α-1,4-linked linear units, and (2) straight-chain starch, consisting of α-1,4-linked linear chains and several α-1,6-linked branched chains [10]. Straight-chain starch is the main component of plant starch, accounting for 70–80% of the weight of starch granules [11]. In addition, glycogen and pullulan are two other natural branched-chain α-glucans. Glycogen (also known as animal starch) is the counterpart of straight-chain starch, which is more branched and is a form of energy storage in animals, fungi, and bacteria [12], while pullulan is a linear polysaccharide produced by the fungus Aureobasidium pullulans and consists mainly of maltotriose repeats linked longitudinally by α-1,6-bonds [13]. Starch is commonly converted into dextrins and minor sugars of different polymerization degrees, such as glucose and maltose oligosaccharides, for industrial applications. Today, enzymatic methods have become the main alternative to chemical methods such as acid hydrolysis for the safe and efficient saccharification and modification of starch [14]. A series of synergistic starch-degrading enzymes, viz. α-Amylase (EC 3.2.1.1, 1,4-α-D-glucan glucose hydrolase), β-amylase (EC 3.2.1.2, 1,4-α-D-glucan malic enzyme hydrolase), glucoamylase (EC 3.2.1.3, dextran 1,4-α-glucosidase), α-glucosidase (EC 3. 2.1. 20), and depolymerase, which is involved in the hydrolysis and depolymerization of starch molecules [15, 16]. In detail, α-amylase acts on the linear part of the starch molecule by catalyzing the breaking of α-1,4 glucosidic bonds at random positions, ultimately producing maltotriose, maltose, glucose, and short-branched chains from straight-chain starch. β-Amylase acts in an outward mode to release maltose from the nonreducing end of the linear chain, while glucosidase and α-glucosidase act on the nonreducing end of straight-chain starch, and straight-chain starch or maltooligosaccharide ends cleave the α-1,4 glycosidic bond to produce a single glucose [17]. However, hydrolysis by these enzymes is interrupted by α-1,6 linkages at the branching point of straightchain starch, and the remaining part is referred to as “extreme dextrin,” which refers to the core part of straight-chain starch with branches and a few α-1,4-linked glucose units. Further degradation of extreme dextrins requires hydrolysis of the α-1,6 branching bonds [18, 19]. Starch debranching enzymes (DBEs) are a class of starch hydrolases that act on α-1,6-glycosidic linkages at the straight-chain starch and glycogen branches and, unlike other starch-active enzymes, have an affinity mainly for α-1,4-glycosidic linkages [20]. It is of great value in the starch industry because of its ability to efficiently eliminate limiting dextrins during straight-chain starch processing, thus improving the utilization and conversion of starch raw materials. Therefore, it has attracted particular research attention in recent decades [21–27]. In this review, the classification, biochemical properties, sequence and structural characteristics, production, and current applications of the enzymes are systematically described in detail. The aim is to summarize and deepen the understanding of the existing research progress, trends, and bottlenecks and to look forward to the development of enzyme engineering and starch debranching enzymes.

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2.2

23

Classification of Microbial SDBEs

SDBEs are usually divided into four broad groups based on substrate specificity (Fig. 2.1): pullulanases (also named limit dextrinases or R-enzymes when sourced from plants), amylose-α-1,6-glucosidases, glucoamylase and isoamylases [20, 28, 29]. Among these four groups, amylose-1,6-glucosidases are of limited use in the starch industry because they employ an indirect debranching mode, where the substrate must be depolymerized prior to the debranching reaction so that the branching consists of a single α-1,6-linked glucose. Glucoamylase is an inverted exo-acting amylase that can act on both α-1,4 and α-1,6 t bonds in polysaccharides such as amylose and straight-chain starch. Conversely, pullulanases and isoamylases, which employ direct debranching action, are the preferred SDBEs for industrial starch processing.

Fig. 2.1 Schematic diagram showing the mode of starch dedifferentiation by starch debranching enzymes (SDBEs) for (SDBEs) on (a) pullulan and (b) amylopectin. The target sites of type I/II pullulanases and isoamylases are marked with blue, green, and red arrows, respectively. The hydrolysis products of the three types of SDBEs are shown. This figure was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [30]

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2.2.1

W. Xia and J. Wu

Glucoamylases

Glucoamylases, also known as amyloglucosidases, are a kind of exo-acting starch hydrolase that continuously removes glucose units from the nonreducing end of the polysaccharide substrate, releasing D-glucose molecules in the β conformation [31, 32]. Glucoamylases have been widely found in prokaryotes and eukaryotes, especially in filamentous fungi. Glucoamylases from Rhizopus oryzae [33], Hypocrea Jecorina [34], Aspergillus niger [35], and Aspergillus awamori [36] have been produced industrially and applied to the production of glucose syrup. In the early 1990s, the three-dimensional structure of glycolytic enzymes from Aspergillus fumigatus X100 was reexamined [37–39]. Glucoamylases mainly act on α-1,4 glucosidic bonds, while many glycosylases have also been reported to be able to cleave branching points in straight-chain starch by hydrolyzing α-1,6 bonds with lower catalytic efficiency than α-1,4 bonds [40–42]. Although the activity of glucoamylase on the α-1,6 bond is only 0.2% of that of the α-1,4 bond, it is sufficient to adversely affect the yield of industrial saccharification [43]. Several studies aimed at improving the substrate specificity of glycosylases have been reported. Arg305 was shown to be an important determinant in differentiating the α-1,4 to α-1,6 substrate specificity of the glucoamylase of Aspergillus niger [44]. The glucoamylase variants Sll9Y, Gl83K, and Sl84H of Aspergillus Awamori showed enhanced selectivity for α-1,4- to α-l,6-linked disaccharides by 2.3- to 3.5-fold [45]. Thus, despite its strong activity against α-1,6-linked branching sites, glucoamylase is used more as an economically important hydrolase in starch glycosylation than as an efficient debranching enzyme.

2.2.2

Pullulanases

Pullulanases can be classified into two types depending on whether the enzyme hydrolyzes only α-1,6 linkages or both α-1,4 and α-1,6 bonds [46]. Although type I pullulanase shows considerable specificity for the α-1,6 bond in polysaccharides, it has no effect on α-1,4 linkages. Type I pullulanase was first identified in a mesophilic bacterium called Bacillus aerogenes (now called Klebsiella aerogenes; the human pathogen K. pneumoniae belongs to the same genus) [47]. Subsequently, type I pullulanase was found to be widely distributed in a variety of microorganisms, including K. acidophilus [48, 49], B. flavocaldarius [50, 51], K. pneumoniae [52], Fervidobacterium pennavorans [53, 54], and Anoxybacillus LM18-11 [55]. Extreme dextrinase (EC 3.2.1.41/142), formerly known as R-enzyme, is almost identical to type I pullulanase but of plant origin and plays an important role in both the degradation of extreme dextrin and the last debranching step in the synthesis of straight-chain starch in barley[56]. Type I pullulanase usually converts pullulan completely to simple maltotriose. When cooperating with enzymes that act on the α-1,4 glucosidic bond, such as α-amylase and glucoamylase, type I pullulanase is

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capable of converting straight-chain starch into a small sugar consisting mainly of glucose and maltose [46]. All the early discovered pullulanases belonged to type I until the discovery of another type of pullulanase that acts bifunctionally on the α-1,4 and α-1,6 glucosidic bonds in both straight-chain starch and pullulan and causes some degree of hydrolysis by acting endocytically on straight-chain starch. The newly defined type II pullulanase, also known as straight-chain pullulanase, exhibits debranching activity at the branching point while exhibiting stochastic endocytic disassembly activity on linear α-1,4-linked polysaccharides [57]. Consequently, the complete conversion of straight-chain polysaccharides to oligosaccharides is accomplished without the synergistic effect of any other enzyme [58, 59]. Like type I pullulanase, maltotriose is the end product of the hydrolysis of pullulan by most starch pullulanases. However, neither type I pullulanases nor straight-chain amylases act on branches containing only one glucose residue, such as glucan, isomaltose, and isomaltotriose [57].

2.2.3

Isoamylases

Isoamylase, often considered a direct debranching enzyme, hydrolyzes α-1,6-D glycosidic linkages in straight-chain starches, glycogen, and some other glucose oligosaccharides with more than three glucose units in their α-1,6-linked branching sugar chains [60]. As another important SDBE along with lalanase, isoamylase has been studied for more than 60 years. A specific type of amylase was first identified in Saccharomyces cerevisiae in 1951. Isoamylase catalyzes the hydrolysis of glutinous rice straight-chain starch to produce linear straight-chain starch precipitates of lower molecular weight and to modify the performance of straight-chain starch substrates in the iodine-chromogenic reaction. The first intracellular isoamylase was isolated from the bacterium Pseudomonas amyloderamosa [61]. Soon thereafter, many other microbial isoamylases were discovered in bacteria such as Bacillus circulans Mir-137 [62], Flavobacterium odoratum KU [63], Lipomyces kononenkoae [64], and Pectobacterium chrysanthemi PY35 [65]. Most isoamylases in plants play an important role in correcting the pruning of misaligned branches that would otherwise prevent the binding of adjacent linear chains, leading to crystallization during straight-chain starch biosynthesis [66]. The two main members of SDBEs, pullulanase, and isopentosidase, have very different substrate specificities, although they both hydrolyze the α-1,6 bond (Fig. 2.1). Pullulanase generally requires the presence of at least two α-1,4-linked glucose units on two sugar chains joined by an α-1,6 bond. However, it has been reported that although the purulanases of Klebsiella pneumoniae efficiently hydrolyze purulan polysaccharides and branched chain dextrins, they show low activity against high molecular weight straight-chain starch (approximately 15% of purulan polysaccharide activity) and glycogen (only 1%) [20]. In contrast, isoamylases preferentially hydrolyze high molecular weight glycogen and straight-chain starch

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and require at least three units of α-1,4-linked glucose; they do not hydrolyze pullulan. Thus, the minimum active substrate units for pullulanase and isoamylase are 62-α-maltose and 63-α-maltotriose, respectively. This apparent difference guides their application in different types of starch conversion and modification processes for the manufacture of products such as glucose syrup [67], resistant starch [60, 68], and cyclodextrins [26, 69]. However, in this regard, little research has been done on isoamylases compared to studies on rattanoidases.

2.3

Sources and Biochemical Properties

SDBEs are present in most animals, plants, and microorganisms and are commonly used for the synthesis or degradation of branched-chain starch or glycogen. Isoamylases are the predominant forms in higher organisms such as animals, plants, yeasts, and molds, while pullulanases are found only in fungi and bacteria [46]. Due to their ease of production in pure form and their appropriate properties, all SDBEs currently used in starch-related industries are of microbial origin, including mesophilic [70], thermophilic [71], hyperthermophilic [72, 73] bacteria, archaea, and some thermophilic fungi [74]. They can be produced by local fermentation or recombinant expression. Depending on their origin, SDBEs have different biochemical properties (Table 2.1).

2.3.1

Temperature Optimum and Thermostability

Commercial glucoamylases such as glucoamylases GA-L NEW (Genencor) [135, 136] and glucoamylase AMG 300 L (Novozyme) [137, 138] are usually used in combination with a desubstituting enzyme during the starch glycosylation phase, where the optimal temperature is generally 55–60 °C. Therefore, the temperature optimization value for industrial desubstituted enzymes is also approximately 60 °C. Previous studies on prurulent enzymes have focused on enzymes with moderate thermal stability, whose temperature selection is most suitable for commercial use (~60 °C, Table 2.1). Thermophilic bacteria are the main producers of this pullulanase enzyme. Many of the pullulanase enzymes obtained from thermophilic bacteria exhibit both thermophilic temperature selection and excellent thermal stability. Gram-positive bacteria, especially Bacillus spp., particularly Bacillus acidophilus [49], B. flavocaldarius [50, 51], B. cereus [77], B. deramificans [78], B. megaterium [79], and B. stearothermophilus [89], are moderately thermophilic Gram-positive bacteria that produce thermostable type I pullulanases with temperature maxima between 50 °C and 70 °C. Numerous type II pullulanases have also been discovered in thermophilic bacteria, including B. circulans [58], Alkalilimnicola sp. NM-DCM-1 [107], Anoxybacillus sp. WB42 [108], Bacillus

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27

Table 2.1 The enzyme type, source, and biochemical properties of starch debranching enzymesa Type and original source GH13 type I pullulanase Anaerobranca gottschalkii Anoxybacillus LM18–11 Anoxybacillus sp. SK3–4 Bacillus acidopullulyticus Bacillus cereus Nws-bc5 Bacillus deramificans Bacillus flavocaldarius Bacillus megaterium Bacillus methanolicus PB1 Bacillus naganoensis Bacillus pseudofirmus 703 Bacillus sp. AN-7 Bacillus sp. CICIM 263 Bacillus stearothermophilus Bacillus subtilis DR8806 Desulfurococcus mucosus Exiguobacterium Acetylicum YH5 Fervidobacterium pennavorans Ven5 Fervidobacterium nodosum Rt17-B1 Geobacillus kaustophilus DSM7263 Geobacillus thermoleovorans NP33 Geobacillus thermoleovorans US105 Hermetia illucens Hordeum vulgare (Barley) Hypocrea jecorina QM9414 Klebsiella pneumoniae Klebsiella variicola SHN-1 Lactococcus lactis IBB 500 Paenibacillus barengoltzii Paenibacillus lautus DSM 3035 Paenibacillus polymyxa Nws-pp2 Shewanella arctica Thermotoga maritime Thermotoga neapolitana GH13 type II pullulanase Alkalilimnicola sp. NM-DCM-1 Anoxybacillus sp. WB42 Bacillus circulans F-2 Bacillus megaterium Y103 Bacillus sp. DSM 405

Optima T (°C)

pH

b

Reference

70 60 60 60 60 55 80 55 50 62 45 90 70 60 70 80 50 80 80 65 60 70 40 40 35–65 50 55 45 50 40 35 35 90 80

8.0 6.4 7.5 5.0 6.0 4.5 7.0 6.5 5.5 5.0 7.0 6.0 6.5 7.0 9.5 5.0 6.0 6.0 5.0 6.0 7.0 6.0 9.0 5.5 6.5 6.0 5.0 4.5 5.5 7.0 6.0 6.0 5.9 6.5

56 750 – 1096.5 44.7 469.7 54 83 292 700 270 154 73.5 49.6 25.6 26 22.1 78 25.93 64.75 – 3.7 228.4 14.2 – 1160 – 2551.42 68.3 13.56 16.17 33.8 130 28.7

[75] [55] [76] [48, 49] [77] [78] [50, 51] [79] [80] [81–83] [84] [85] [86] [87–89] [90] [91] [92] [53, 54] [93] [94] [95] [96] [97] [98] [74] [52] [99, 100] [101] [102] [103] [104] [105] [71] [106]

50 55–65 50 45 70

9.5 5.5 7.0 6.5 6.0

599 – 1400

[107] [108] [58] [109] [110]

Specific activity (U/mg)

11,000

(continued)

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Table 2.1 (continued) Type and original source Bacillus sp. KSM-1378 Exiguobacterium sp. SH3 Geobacillus stearothermophilus L l4 Geobacillus thermoleovorans NP33 Hermetia illucens Lactobacillus amylophilus GV6 Streptomyces erumpens MTCC 7317 Thermoanaerobacter ethanolicus 39E Thermoanaerobacter strain B6A T. thermosaccharolyticum DSM 571 Thermus thermophilus HB27 GH57 type II pullulanase Cohnella sp. A01 00831 Cohnella sp.A01 01133 Desulfurococcus amylolyticus JCM 9188 Pyrococcus furiousus Pyrococcus woesei Thermococcus siculi KOD1 Thermofilum pendens GH13 Isoamylase Bacillus circulans Mir-137 Bacillus lentus JNU3 Bacillus sp. Bacillus sp. CICIM 304 Flavobacterium odoratum KU Flavobacterium sp. Lipomyces kononenkoae Manihot esculenta Crantz Pectobacterium chrysanthemi PY35 Pseudomonas amyloderamosa Sulfolobus solfataricus ATCC 35092 Rhizopus oryzae PR7 MTCC 9642

Optima T (°C) 50 45 65 60 40 37 50 60 65 65 70

pH 8.5 4.5 5.5 7.0 9.0 6.5 7.0 5.5 5.0 5.0 6.5

83.1 – 967 795 228.4 7.142 98.84 480 215 14 280

Reference [111] [112] [113] [114] [97] [70] [115] [116] [117] [118] [119]

60 70 95

6.0 8.0 5.0

118.84 52.79 –

[120] [120] [121]

105 100 100 95–100

6.0 6.0 5
5 3.5

77.5 35 – 37.2

[72, 73] [122] [123, 124] [125]

60 70 55 70 45–50 40 30 37 40 52 75 55

5.0 6.5 9.0 6.0 6.0 6–7 6.5 7.0 7.0 3.5 5.0 5.5

– – 469 6535 76,400 54,600 52 – – 59,100 4600 52.41

[62] [126] [127] [128] [63] [129] [64] [130] [65] [131, 132] [133] [134]

Specific activity (U/mg) b

a This table was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [30] b The presented values are specific activities using pullulan or amylopectin as the substrate

sp. DSM 405 [110], Streptomyces erumpens MTCC 7317 [115], Thermus thermophilus HB27 [119], and Thermoanaerobacter ethanolicus [116]. In addition to an optimum intermediate temperature, SDBE suitable for industrial applications must exhibit good thermal stability. The saccharification phase of starch

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29

processing usually takes 48–60 h, during which the enzyme in question must remain effectively active. Furthermore, an idealized simultaneous saccharification/liquefaction process requires the use of both α-amylase and pullulanase at 95–115 °C and a pH of 4.5–6.5. Although this combined process is more efficient, the temperature selection and thermal stability of the SDBEs involved must be much higher than those used at the conventional temperature conditions of 55–60 °C. Therefore, the moderately thermally stable pullulanase obtained from the above sources does not meet the thermal stability requirements of the enzyme. The current commercial pullulanase enzymes used for starch processing are Promozyme (Novozyme) from Bacillus acidopullulyticus (Novozyme) [48] and OptimaxL (Genencor) from B. licheniformis (Genencor) [139]. Although these enzymes may be able to solve the relatively low feedstock utilization problem currently prevalent in the starch industry, studies focusing on increasing the optimal temperature of superheatresistant Platanase are not sufficient. The development of improving the optimal temperature and thermal stability of pullulanase would be quite valuable and necessary to improve stepwise or simultaneous starch saccharification. In recent years, various thermophilic and thermophilic microorganisms have been found to produce heat-stable pullulanases, particularly type II pullulanases (Table 2.1). Type I pullulanases from Bacillus sp. AN-7 and Thermotoga maritime showed prominent high-temperature selectivity (90 °C) [71, 85], and type I pullulanases from Desulfurococcus mucosus [91], Fervidobacterium pennavorans Ven5 [53, 54], and Thermotoga neapolitana [106] showed the best activity of type I pullulanases at 80 °C. Amylopullulanases are more common in extremely thermophilic microorganisms than type I pullulanases. Pyrococcus furiousus [72, 73], Pyrococcus woesei [122], Thermococcus siculi KOD1 [123, 124], and Thermofilum pendens [125] provided the most thermally stable amylopullulanases, which were selected for a temperature range of 95–105 °C. The straight-chain mannanase from the extremophilic archaeon P. furiosus is one of the most thermotolerant mannanases to date, exhibiting maximum hydrolysis at 105 °C and maintaining activity at 130–140 °C under conditions of 5 mM Ca2+ [72]. Despite this exemplary activity, superthermostable amylopullulanase has not been produced commercially due to its low yield and limited activity under acidic conditions [57]. Other major sources of pullulanase are mesophilic microorganisms such as Hermetia illucens [97], Shewanella arctica [105], Klebsiella pneumoniae [52], Lactococcus lactis IBB 500 [101], Paenibacillus barengoltzii [102], Paenibacillus lautus DSM 3035 [103], and Paenibacillus polymyxa [104]. The pullulanases produced by them exhibited low heat resistance with a temperature adaptation range of 30–45 °C. Most mesothermal or chillophilic (cold-adapted) pullulanases belong to type I. Exceptions of low-temperature amylopullulanases also exist, such as cases from Hermetia illucens [97] and Exiguobacterium sp. SH3 [140]. P. polymyxa Nws-pp2 PulN [104] and Exiguobacterium sp. SH3 Pul-SH3 [140] are representative cold-adapted pullulanases of the enzymes that have been discovered. PulN showed optimal activity at 35 °C and maintained 23% activity at 10 °C when expressed under low-temperature induction in E. coli. Similarly, at a moderate temperature of 30 °C, the amylopullulanase Pul-SH3 showed significantly

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high activity, maintaining 23% activity at 0 °C. Although cardiophilic pullulanases are not suitable for high-temperature industrial processes such as conventional starch saccharification, they are candidates for specific applications requiring enzymatic treatment at low temperatures. The original microbial sources of most isoamylases are mesophilic and thermophilic bacteria. Bacillus circulans Mir-137 [62], B. lentus JNU3 [126], Bacillus sp. CICIM 304 [128], Pseudomonas amyloderamosa [132], and Sulfolobus solfataricus ATCC 35092 [133] produce temperature-stable isoamylases with temperature optima of 50 °C–75 °C. Psychrophilic isoamylases have optimal activity at low temperatures. For instance, Pectobacterium chrysanthemi PY35 expressed an intracellular mesophilic isoamylase showing high activity at pH 7.0 and 40 °C [65]. Other sources of mesophilic isoamylases are eukaryotic microorganisms such as Lipomyces kononenkoae [64]. Unlike pullulanase, which is localized primarily outside the surface membrane, isoamylase is an intracellular enzyme that is usually involved in glycogen hydrolysis for energy utilization. This distinctive feature may account for the low- or medium-temperature adaptation of isoamylases. To date, no extremely thermostable and heat-resistant isoamylases have been identified, which limits the application of isoamylases in processes requiring high temperatures.

2.3.2

pH Optimum

The optimal pH for microbial SDBEs is in the acidic to neutral range (4.0–7.0). The slightly acidic SDBEs found in different species of microorganisms are favored in the food industry. The genus Bacillus is an important producer of such SDBEs. For example, type I pullulanases from B. flavocaldarius [48, 49], B. megaterium [79], B. naganoensis [81–83], and B. acidopullulyticus [48, 49], amylopullulanases from B. circulans F-2, Bacillus sp. DSM 405 and Bacillus sp. KSM-1378 [58, 110, 111], and isoamylases from B. circulans Mir-137 [62], B. lentus JNU3 [126], Bacillus sp. CICIM 304 [128] showed a pH range of 5.0–7.0 for maximum activity. Acid- and basophilic SDBEs have also been identified in bacteria. Of these, due to its valuable thermal stability and good activity under acidic environments, B. acidopullulyticus amylopullulanase has been applied in commercial usage [48]. Pullulanase from Clostridium thermothiophilum DSM 3783 showed maximum activity at 75 °C and pH 3.0–5.5 [141]. The isoamylase from Pseudomonas amyloliquefaciens had a very high specific activity with straight-chain starch as substrate at pH 3.5, up to 59,100 U/mg, as reported [131, 132]. Some SDBEs also preferentially functioned at high pH values ranging from 8.0 to 9.5. Reported sources of these alkaline SDBEs included Anaerobranca gottschalkii [75], Alkalilimnicola sp. NM-DCM-1 [107], and B. subtilis DR8806 [90]. Among all reported alkaline cases, pullulanases from Alkalilimnicola sp. NM-DCM-1 exhibited optimal activity at pH 9.5 and 50–70 °C, representing the most alkaline-tolerant SDBEs [107]. However, neither of these extreme alkalophilic bacteria could provide the required specific activity or thermal stability.

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Classification and Enzyme Properties of Starch Debranching Enzymes

2.3.3

31

Specific Activity

The ability to debranch when employing substrates containing α-1,6-glycosidic linkages, such as amylopectin or pullulan, is the most common way that the particular activity of SDBE is expressed [141]. The specific activity of enzymes from various sources ranged from 3.7 to 76,400 U/mg (Table 2–1). As enzymes from closely similar species, such as Bacillus spp., had extremely substantial variances in specific activity (e.g., 3.7–11,000 U/mg of pullulanase), no clear pattern was found to explain these significant differences in a particular activity. In general, isoamylases have much higher specific activities and hydrolyze a wider range of substrates than pullulanases. The most active pullulanase is the straight-chain pullulanase from Bacillus sp. DSM 405, which shows a specific activity of 11,000 U/mg against straight-chain starch at 70 °C and pH 6.0 [110]. Flavobacterium odoratum KU [63], Flavobacterium sp. [129], and Pseudomonas amyloderamosa [131, 132] produced isoamylases with higher specific activities (76,400 U/mg, 54,600 U/mg, and 59,100 U/mg, respectively). However, these highly active isoamylases prefer a much lower temperature (~50 °C) than the temperature preferred by pullulanase.

2.4

Commercial SDBEs

To our data, the foremost widely used industrial SDBE area unit is the Promozyme series from Novozymes A/S and Optimax L-1000 from Genencor Int. Promozyme could be a lalanase made by B. acidopullulyticus and was developed by Novozyme in Scandinavian nations within the 20 developed as an advert catalyst preparation within the early 1980s [142–144]. This family of pullulanases includes Promozyme 200 L (200 PUN/g), Promozyme 600 L (600 PUN/ml), and Promozyme D2. It is acid and heat resistant and exhibits temperature and pH optimums of 60 °C and pH 4.5, respectively. Novozymes’ Promozyme has been sold in the Japanese and European markets since 1983 and has a large market share in China and worldwide. Optimax L-1000 is a pullulanase obtained from the fermentation broth of Bacillus licheniformis and is widely used in the beer, starch, and alcohol industries [145]. It also has optimal values for temperature and pH of 60 °C and 4.5, respectively, but works effectively at pH 4.0–6.5 and 40–65 °C. These commercial SDBEs generally require cooperation with other amylases to form complex enzymes for specific processing. For example, SDBEs can be used in combination with β-amylase for high maltose syrup production and with α-amylase and β-glucanase for beer brewing. Acknowledgments This chapter was modified from the paper published by our group in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [30]. The related contents are reused with permission.

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98. Vester-Christensen MB, Hachem MA, Naested H, Svensson B. Secretory expression of functional barley limit dextrinase by Pichia pastoris using high cell-density fermentation. Protein Expr Purif. 2010;69:112–9. 99. Chen WB, Nie Y, Xu Y. Signal peptide-independent secretory expression and characterization of Pullulanase from a newly isolated Klebsiella variicola SHN-1 in Escherichia coli. Appl Biochem Biotechnol. 2013;169:41–54. 100. Mu GC, Nie Y, Mu XQ, Xu Y, Xiao R. Single amino acid substitution in the Pullulanase of Klebsiella variicola for enhancing Thermostability and catalytic efficiency. Appl Biochem Biotechnol. 2015;176:1736–45. 101. Wasko A, Polak-Berecka M, Targonski Z. Purification and characterization of Pullulanase from Lactococcus Lactis. Prep Biochem Biotechnol. 2011;41:252–61. 102. Liu J, Liu Y, Yan F, Jiang Z, Yang S, Yan Q. Gene cloning, functional expression and characterisation of a novel type I pullulanase from Paenibacillus barengoltzii and its application in resistant starch production. Protein Expr Purif. 2016;121:22–30. 103. Chen S-Q, Cai X-H, Xie J-L, Wei W, Wei D-Z. Structural and biochemical properties of a novel pullulanase of Paenibacillus lautus DSM 3035. Starch-Starke. 2017;69:1500333. 104. Wei W, Ma J, Chen SQ, Cai XH, Wei DZ. A novel cold-adapted type I pullulanase of Paenibacillus polymyxa Nws-pp2: in vivo functional expression and biochemical characterization of glucans hydrolyzates analysis. BMC Biotechnol. 2015;15:96. 105. Elleuche S, Qoura FM, Lorenz U, Rehn T, Brück T, Antranikian G. Cloning, expression and characterization of the recombinant cold-active type-I pullulanase from Shewanella arctica. J Mol Catal B Enzym. 2015;116:70–7. 106. Kang J, Park KM, Choi KH, Park CS, Kim GE, Kim D, Cha J. Molecular cloning and biochemical characterization of a heat-stable type I pullulanase from Thermotoga neapolitana. Enzym Microb Technol. 2011;48:260–6. 107. Mesbah NM, Wiegel J. Biochemical characterization of halophilic, alkali thermophilic amylopullulanase PulD7 and truncated amylopullulanases PulD7DeltaN and PulD7DeltaC. Int J Biol Macromol. 2018;111:632–8. 108. Wang J, Liu Z, Zhou Z. Cloning and characterization of a novel thermophilic Amylopullulanase with a type I Pullulanase structure from Anoxybacillus sp. WB42. Starch-Starke. 2018;70:1700265. 109. Liu X, Chen H, Tao HY, Chen Z, Liang XB, Han P, Tao JH. Cloning and characterization of a novel amylopullulanase from Bacillus megaterium Y103 with transglycosylation activity. Biotechnol Lett. 2020;42:1719–26. 110. Brunswick JM, Kelly CT, Fogarty WM. The amylopullulanase of Bacillus sp. DSM 405. Appl Microbiol Biotechnol. 1999;51:170–5. 111. Ara K, Igarashi K, Saeki K, Ito S. An Alkaline Amylopullulanase from Alkalophilic Bacillus sp. KSM-1378; Kinetic evidence for two independent active sites for theα-1,4 andα-1,6 hydrolytic reactions. Biosci Biotechnol Biochem. 1995;59:662–6. 112. Rajaei S, Noghabi KA, Sadeghizadeh M, Zahiri HS. Characterization of a pH and detergenttolerant, cold-adapted type I pullulanase from Exiguobacterium sp. SH3. Extremophiles. 2015;19:1145–55. 113. Zareian S, Khajeh K, Ranjbar B, Dabirmanesh B, Ghollasi M, Mollania N. Purification and characterization of a novel amylopullulanase that converts pullulan to glucose, maltose, and maltotriose and starch to glucose and maltose. Enzym Microb Technol. 2010;46:57–63. 114. Nisha M, Satyanarayana T. The role of N1 domain on the activity, stability, substrate specificity and raw starch binding of amylopullulanase of the extreme thermophile Geobacillus thermoleovorans. Appl Microbiol Biotechnol. 2015;99:5461–74. 115. Kar S, Ray RC, Mohapatra UB. Purification, characterization and application of thermostable amylopullulanase from Streptomyces erumpens MTCC 7317 under submerged fermentation. Ann Microbiol. 2011;62:931–7.

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Chapter 3

Sequence, Structure, and Engineering of Microbial Starch Debranching Enzymes Wei Xia, Lei Wang, and Jing Wu

Abstract Starch dedifferentiating enzymes (SDBEs) belong mainly to the glycoside hydrolase (GH) family 13 and 57, except for type II pullulanases of GH57, GH13 pullulanase, and isoamylases, which share many similarities in the sequence and structure of the core catalytic domain. However, the N-terminal structural domain, which may be one of the determinants contributing to the substrate binding of SDBEs, is different in the different enzymes. To overcome the present deficiencies of SDBEs in terms of chemical action potency, thermal stability, and expression levels, great efforts have been made to develop effective accelerator engineering and fermentation methods. Herein, we summarize the different biochemical properties of pullulanase and isopentosidase from different sources as well as the distinctive features in sequence and structure. Recent developments in the enzymatic engineering, heterologous production, and industrial applications of SDBEs are also reviewed. Finally, research perspectives that contribute to the understanding and broadening of SDBE applications are provided. Keywords Pullulanases · Isoamylases · Enzyme structure · Enzyme engineering

3.1

Introduction

SDBEs have attracted tidy analysis interest as necessary biocatalysts for starch processing and preparation of downstream merchandize. Over the past decades, studies associated with the identification and characterization of enzymes have provided an upscale resource for a range of applications with totally different desires. The assorted SDBEs discovered to this point show outstanding similarities and W. Xia · L. Wang · J. Wu (✉) State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, 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. 2023 J. Wu, W. Xia (eds.), Industrial Starch Debranching Enzymes, https://doi.org/10.1007/978-981-19-7026-9_3

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important variations in several aspects of their chemical change properties. Structures solved in recent years have also helped to decipher the bond type specificity between α-1,6 and α-1,4 bonds and to identify sequence motifs and structural bases to functionally distinguish various types of SDBEs. These substructures and sequence motifs may serve as inspiration for stimulating enzyme engineering in previously reported or upcoming studies. Despite these advances, more data on the structures of various SDBEs, as well as complementary enzymatic properties, are still needed to provide sufficient details of structure–function relationships to enable rational engineering of potentially industrially relevant enzymes. Another important consideration for the practical application of SDBEs is that the yield of heterologous production is usually kept relatively low, except for a few prominent cases, especially for thermally stable enzymes with high potential from extremely hydrophilic bacteria. Although considerable efforts have been made to enhance the traditional applications of SDBEs and to develop new ones, their application requirements have not been fully met due to the increasing demands on productivity and cost. In addition, starch from different sources varies greatly in structure and branching ratios. Currently, most researchers have given only limited attention to the application of SDBEs in the processing of starch from traditional sources, such as corn, potato, and wheat. However, the utilization of some specific sources of polysaccharides, such as pea starch, taro starch, and even animal glycogen, has been little explored. This has led to a lack of awareness of the application needs of these polysaccharides, as well as a lack of suitable enzymes, including SDBEs.

3.2

Sequence Classifications

According to the Carbohydrate-Active Enzymes Database (CAZy, http://www.cazy. org/), the two major classes of SDBEs, namely, pullulanases and isoamylases, are mainly distributed in glycoside hydrolase (GH) families 13 and 57, based on amino acid sequence similarities[1, 2]. In detail, all SDBEs included in GH57 are type II pullulanases or pentyl pullulanases. Type I pullulanases, some mesophilic or heatstable pentyl pullulanases, and all isoprenylases are included in GH13 [3, 4]. Phylogenetic analysis of the characterized or putative SDBEs showed that they are distributed in three clusters: type I and type II pullulanases in GH13, isoamylases in GH13 and type II pullulanases in GH57 (Fig. 3.1). Enzymes of both GH13 and GH57 are considered to be part of the broad α-amylase part of the larger family. The difference is that the GH13 family represents the major α-amylase family, whereas GH57 is a smaller group established later and has some degree of similarity and dissimilarity to the GH13 family [5]. Most mesophilic amylopullulanases belong to the GH13 family, while most heat-stable amylopullulanases are present in GH57 [6, 7]. The organic process tree was performed by ClustalW and MEGA7 for multiple sequence alignment of organic compound sequences of SDBEs within the NCBI information victimization the neighbor-joining methodology, followed by rendering

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Fig. 3.1 Evolutionary analysis of starch debranching enzymes (SDBEs)

victimization the iTOL online tool (https://itol.embl.de/) [8]. The GenBank accession range of every SDBE is indicated before the species name. The biological group representing GH13 pullulanases contains the GH13_12–14 taxon and is shown in red. Clades representing GH13 isoamylases and GH57 amylocellulases are shown in blue and inexperienced, respectively. This figure was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9]. GH13 has a large number of members and is rapidly increasing. This large number of sequences has led to the definition of subfamilies within GH13 that are defined based on the higher sequence similarity in the catalytic TIM barrel domain [10, 11]. To date, 42 subfamilies have been designated in GH13. Their members

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have different catalytic activities and substrate specificities and are named α-amylase, amylase, cyclodextrin glucosyltransferase, α-glucanase, and 1,4-α-glucan branched enzymes. sDBEs in GH13 have been distributed in several different subfamilies. Isoamylases or glycogen fractionases from bacteria, archaea, and eukaryotes (EC 3.2.1.68) mainly belong to GH13_11. Pullulanase type I from Bastinia and pullulanase type II with two catalytic structural domains belong to GH13_12, while GH13_14 is composed of the bacterial counterparts of these enzymes. GH13_13 is mainly composed of pullulanases or glycogen fractionases from bacteria and eukaryotes. Pullulanases or limiting dextranases from organisms [3, 11]. Some type II pullulanases with one or two catalytic structural domains have recently been assigned to the GH13_39 and GH13_41 subfamilies, respectively. In addition, other debranching enzymes named oligosaccharide- or glucose1,6-α-glucosidases that hydrolyze the α-1,6-linked branches of linear isomaltooligosaccharides, pantothenes, and glucans have been classified as members of subfamilies 31 and 23 of GH13 [3, 12]. The SDBEs in GH57 are mainly straight-chain amylases with a single catalytic structural domain that act on the α-1,4-linked backbone and α-1,6 branches in starch. It is worth mentioning that the enzymes in GH57 are mainly from polarophilic microorganisms. As described in Chap. 1, GH57 contains amylases produced by thermophilic and archaeal bacteria, such as Pyrococcus furiousus [13, 14], Pyrococcus woesei [15], Thermococcus siculi KOD1 [16, 17], and Thermofilum pendens [18], which exhibit extreme thermophilic temperature optimization of 95–105 °C. Another example is the GH57 straight-chain amylase from Staphylothermus marinus (SMApu). This enzyme shows maximum pullulan degradation activity at 105 °C, excellent thermal stability at 100 °C with a half-life (t1/2) of 50 min, and degrades cyclodextrins [19].

3.3

Structural Features

Until now, crystal structures of ten pullulanases belonging to GH13 have been resolved (as presented in Table 3.1). As shown in Fig. 3.2, ost GH13 pullulanases usually adopt a multidomain structure containing one or several carbohydrate-binding domains (CBM) and an X domain of unknown function at the N-terminus, and the typical catalytic domain of the GH13 family consists of domains A and B, as well as a C-terminal domain [20–24]. Structural domain A adopts the classical (β/α)8 TIM barrel structure and contains the basic catalytic triad of β4-Asp, β5-Glu, and β7-Asp. For example, in the hydrolysis of BaPul13A (PDB ID: 2 WAN) of B. acidopullulyticus, the nucleophile, general acid/base, and transition state stabilizer are β4-Asp622, β5-Glu651, and β7Asp736, respectively, in structural domain A [21]. In pullulanase PulA from Anoxybacillus sp. LM18–11, the three corresponding residues were identified as Asp413 [20]. In addition, residue Trp708, the +2 binding subsite of pneumococcal pulanase, was found to be located in this structural domain and induces the

GH13 GH13

Bacillus acidopullulyticus Paenibacillus Barengoltzii

Anoxybacillus sp. LM18-11

Bacillus subtilis str. 168

Klebsiella oxytoca Nostoc punctiforme

Klebsiella pneumoniae

PulA

AmyX

PulA NPDE

Pul

GH13

GH13 GH13

GH13

GH13

GH Family

Source

Enzyme Pullulanase BaPulA PbPulA

1083

1078 488

718

710

921 698

Length (aa)

Table 3.1 Solved crystal structures of starch debranching enzymes

1.65 1.71 2.03 1.98 1.8 1.932

– – D350A – – – D470A – – – – – – – – – – – – – – –

2WAN 6JHF 6JHH 6JFX 6JEQ 6JFJ 6JHI 6JHG 3WDH 3WDI 3WDJ 2E8Y 2E8Z 2E9B 2YOC 2WC7 2WCS 2WKG 2FGZ 2FH6 2FH8 2FHB

2.32 1.89 1.75 2.2 2.22 2.11 2.2 2.3 2.88 2.37 2.8 3.0 1.75 1.8 1.9 1.8

Resolution (Å)

Mutant

PDB entry None None Maltotriose Maltotriose, maltopentaose β-Cyclodextrin α-Cyclodextrin, maltotriose Maltose None None Maltotriose Maltopentaose None α-Cyclodextrin Maltose, maltotriose None Acarbose None None None Glucose Isomaltose Maltose

Ligands

Sequence, Structure, and Engineering of Microbial Starch Debranching Enzymes (continued)

[26]

[24] [25]

[23]

[22]

[20] [21]

Reference

3 45

Source

Klebsiella pneumoniae

Klebsiella pneumoniae

Hordeum vulgare

Enzyme

PulA

PulA

HvLD

Table 3.1 (continued)

GH13

GH13

GH13

GH Family

905

884

1102

1053

Length (aa)

Resolution (Å) 1.85 1.65 1.30 1.50 1.84 1.64 2.30 2.59 2.32 2.20 2.23 2.05 1.96 2.1 2.49 2.67 1.9 1.75 1.6 1.7 1.45 1.67

Mutant – – – – G68L G68L – – – – – – – – – – – – – – – E510A

5YN2 5YN7 5YNC 5YNE 5YND 5YNH 5YNA 2Y4S 2Y5E 4CVW 4AIO 4J3S 4J3T 4J3U 4J3V 4J3W

PDB entry 2FHC 2FHF 6 J33 6j34 6 J35 6J4H Ligands Maltotriose Maltotetraose None Maltotriose G680L mutant G680L mutantmaltotriose None β-Cyclodextrin β-Cyclodextrin α-Cyclodextrin γ-Cyclodextrin γ-Cyclodextrin α-Cyclodextrin α-Cyclodextrin β-Cyclodextrin Endogenous inhibitor (LDI) None Maltotriose, maltotetraose Maltotetraose MaltosylS-β-cyclodextrin G2SG24 G3G13 [31] [32]

[30]

[29]

[28]

[27]

Reference

46 W. Xia et al.

GH13

GH13 GH13

GH13

GH13

GH13

750

657 840

718

714

877

2VNC 2VR5 2VUY 2WSK 4J7R 4OKD 1BF2

– – – – – – –

E510A – – – – – 3.0 2.8 3.0 2.25 2.3 2.4 2.0

1.75 2.1 2.4 1.85 2.25 2.37 None Acarbose None None None Maltoheptaose None

GG23-G23 None Maltotetraose None Maltotetraose G-moranoline

This table was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9].

Pseudomonas amyloderamosa SB-15

IAM

Sulfolobus solfataricus P2

Isoamylase TreX

Escherichia coli str. K-12 Chlamydomonas reinhardtii CC425

Streptococcus pneumoniae

SAP

GlgX ISA1

Streptococcus agalactiae

SpuA

4J3X 3FAW 3FAX 2YA0 2YA1 2YA2

[38]

[36] [37]

[35]

[34]

[33]

3 Sequence, Structure, and Engineering of Microbial Starch Debranching Enzymes 47

48

W. Xia et al.

Fig. 3.2 Domain features of starch debranching enzymes (SDBEs). Domain arrangements of GH13 SDBEs with resolved structure and a GH57 amylopullulanase (GenBank: AF113969) with unknown structure data. The domain information is illustrated by the DOG tool [27]. This figure was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9]

coordinated movement of the acid/base catalytic residue Glu706, which is quite important for the catalytic process [25]. Domain B is a smaller module that varies considerably among the different pullulanases and is located between the third β-strand and the third α-helix of the TIM barrel. Domain C has a Greek key topology and consists of a variable number (5–10) of β-strands [20, 26]. Furthermore, GH13 pullulanases differ greatly in the number and type of N-terminal structural domains but share a common feature of containing the same glycogen/starch binding CBM48 structural domain near the catalytic structural domain (Fig. 3.3) [28]. BaPul13A from B. acidopullulyticus (PDB ID: 2 WAN) has three additional modules in sequence at its N-terminus in addition to CBM48, namely, CBM41-X45-X25 [21], and PulA from Klebsiella oxytoca (PDB ID: 2YOC) contains CBM41 and an N2 structural domain of unknown function (residues 161–265) [24, 29]. The CBM41 module of GH13 pullulanase from T. maritima

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49

Fig. 3.3 Structural options of starch debranching enzymes (SDBEs)

was shown to have binding capacity for straight-chain amylose, straight-chain starch, and pullulan [30]. PulA from Anoxybacillus sp. LM18-11 and the limit dextrinase HvLD from Hordeum vulgare have only one such as CBM21 and other N-terminal structural domains of CBM68 [20, 31, 32]. Recently, residues Phe553/ Trp512 and Phe620/Asp621, located in the N-terminal CBM21-like structural domain of HvLD, were reported to affect the catalytic activity and determine the preference for branched substrates due to possible long-range effects on the active site or heterologous stimulation of the activity [33]. Janecek et al. noted that, based on extensive amylolytic enzyme studies, there is a general consensus that CBM 20, 21, 41, 48, and 68 are able to bind to protostarch [34]. In addition, the structures of four isoamylases, namely, the isoamylases ISA1 from Chlamydomonas reinhardtii CC425 (PDB ID: 4J7R) [35] and IAM from Pseudomonas amyloderamosa SB-15 (PDB ID: 1BF2) [36] and the glycogendebranching enzymes TreX from Sulfolobus solfataricus P2 (PDB ID: 2VNC) [37] and GlgX from Escherichia coli str. K-12 (PDB ID: 2 WSK) [38], have been identified. The structure of isoamylases is similar to that of pullulanase in GH13 and contains typical parts. The N-terminal structural domain, the catalytic structural

50

W. Xia et al.

domain, and the C-structural domain. The catalytic triad of isoamylase is also located on CSR II-III-IV, and its spatial arrangement is similar to that of pullulanase. The three catalytic residues Asp336, Glu371, and Asp443 of GlgX from E. coli K12 are located at the bottom of the active site cleft. Residues Trp219, His442, Asp336, Tyr221, and Leu337 form deeply buried -1 subsites, which are essential for the interaction with the substrate [38]. However, unlike the multiple N-terminal structural domains present in pullulanases, isoamylases usually contain only one CBM48 structural domain at their N-terminus (Fig. 3.2). This structural difference may be related to their different substrate specificities, resulting in less spatial hindrance during substrate binding for isoamylases than for pullulanases, making it easier to target polysaccharides with dense branching, such as the branching point of glycogen. In contrast, pullulanases typically have more multiple N-terminal structural domains than isoamylases, which may be associated with differentiated substrate binding. Examples of this phenomenon include PulA (PDB ID: 3 WDH) from Anoxybacillus sp. LM18–11 has CBM68 and CBM48 structural domains, the limiting dextrase (HvLD) from Hordeum vulgare (PDB ID: 2Y4S) possesses CBM21 and CBM48 structural domains at the N-terminus, and BaPul13A (PDB ID: 2 WAN) from B. acidopullulyticus has additional N1, N2, CBM41, and CBM48 structural domains, while Escherichia coli str. K-12 (PDB ID: 2 WSK) has only one CBM48. Apart from the characteristic catalytic (β/α) 8-barrel of GH13, the typical pullulanase structure despite the overall high structural similarity between the GH13_12-14 multistructural domains of PULI, the sequence identity and similarity of pullulanases is low, even for the catalytic structural domain. Thus, GH13_12-14 enzymes differ in terms of substrate specificity and preference. In addition to the catalytic (β/α)8-barrel characteristic of GH13, typical pullanase structures have one or more N-terminal structural domains, including CBMs, and a C-terminal structural domain common to most GH13 enzymes. These auxiliary structural domains are associated with specific differences among the three PULI GH13 subfamilies. Unfortunately, one or more N-terminal structural domains are missing in some structures of GH13_12 and GH13_14. Recently, the first structure of a branched substrate in complex with PULI was identified, namely, barley limiting dextrinase (PDB entry 4 J3 W, Table 3.1). The structures of four GH13_11 enzymes have been identified (Table 3.1). The crystal structures of Klebsiella pneumoniae pullulanase and its complexes with glucose (G1), maltose (G2), isomaltose (isoG2), maltotriose (G3), or maltotetrasaccharide (G4) have been refined at a resolution of approximately 1.7–1.9 Å by using the synchrotron source of Spring-8. The crystals used for X-ray diffraction experiments were prepared by the suspension-drop method and the vapor diffusion method. The refined model of the enzyme shows possession of 920–1052 amino acid residues, 942–1212 water molecules, 4 or 5 calcium ions, bound sugar molecules, and five structural domains (N1, N2, N3, A, and C). The N1 structural domain is only clearly visible in the structure of the complex with maltotriose or maltotetrasaccharide. The N1 and N2 structural domains are characteristic of pullulanase, while the N3, A, and C-structural domains have a weak

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similarity to the isoamylase of Pseudomonas aeruginosa. The N1 structural domain was found to be a novel carbohydrate-binding structural domain with a calcium site (CBM41). The GH13_11 crystal structure shows the same overall multimodal organization: an N-terminal carbohydrate-binding module family 48 (CBM48), the catalytic structural domain, and a C-terminal β-sandwich structural domain (Fig. 3.3). Most GH13 enzymes contain this C-terminal β-sandwich structural domain. Its function in relation to debranching is unknown, but the C-terminal structures of other GH13 enzymes, such as GH13 branching enzymes, have been shown to be involved in substrate binding and/or shield the hydrophobic residues of the catalytic structural domain from contact with the solvent. In the GH13_11 ISA from C. reinhardtii, the C-terminal structural domain is involved in dimerization. Comparison of the composition of N-terminal CBM domains of various SDBEs. The careful structure data and literature references of concerned SDBEs may be found in Table 3.2. The color code of the CBM domains is in keeping with the domain organization summary. Visual images of the structures were achieved by PyMol computer code. This figure was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9]. The binding pockets of GH13 SDBEs are highly similar across types. Several important binding and catalytic residues are located on the loops around the pocket (Fig. 3.4). The crystal structure of a barley limiting dextrase E510A mutant complexed with 62-α-D-maltotriose (G3G13, a straight-chain amylose fragment, PDB ID 4 J3 W) and 63-α-D-glucosyl maltotriose (GG23-G23, a pullulan fragment, PDB ID 4J3X) shows the active center of SDBE [22]. A summary of the structure is shown in surface kind, and an exaggerated read of the ring containing the preserved binding residues within the pocket is shown as a cartoon drawing in several colors. Active residues and ligands are indicated by lines. The ligands G3G13 and GG23-G23 are shown in cyan and inexperienced, respectively. The image of the structures was achieved by PyMol computer code. This figure was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9] Structure-based multiple sequence comparison of SDBEs with known structural information revealed that five conserved sequence regions (CSRs) are distributed across these loops, and they may play a key role in ensuring the formation of the correct catalytic conformation during substrate recognition (Fig. 3.5) [22, 54]. Among them, the CSR regions (I–IV) show almost absolute conservation in both pullulanases and isoprenozymes and are widely detected in the large family of GH13 starch-acting hydrolases such as amylases and cyclodextrin glycosyltransferases [55, 56]. In addition, the catalytic triad Asp-Glu-Asp is located in CSR II-III-IV. In contrast, motif I is an additional consensus sequence found in GH13 SDBEs and shows a high discriminatory feature with the unique conserved sequences of YNWGYDP in pullulanases and NYWGY in isoamylases [3]. According to the conserved analysis, sequences from the GH57 family usually contain an N domain of GH57N_APU and a glucanase-like C-terminal domain, which is very different from pullulanases [5, 19, 57]. However, the precise 3D structure of the type II pullulanases from GH57 remains unknown.

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W. Xia et al.

Table 3.2 Protein engineering of starch debranching enzymes Enzyme PulPY

Source Pyrococcus yayanosii CH1

Strategy Combined truncations of N- and C-terminal domains

PulSL3

Alkalibacterium sp. SL3

N-terminus truncation

PulB

Bacillus naganoensis

N-terminus truncation

Tailoring of the active sites lining the catalytic pocket Rational design based on homology modeling and sequence analysis Active hydrogen bond network (AHBN) Evolutionary coupling saturation mutagenesis

BaPul

Bacillus acidopullulyticus

Truncation of N-terminal domains

Hydrogen bond-based engineering KvPulA

Klebsiella variicola SHN-1

Single amino acid substitution

BlIam1

Bacillus lentus

Semi-rational design on essential residues

Variant and effect Δ28N + Δ791C had an increased optimum temperature of 100 °C and a sevenfold specific activity relative to the wild type rPulSL3△N had better thermostability, higher substrate affinity, and higher catalytic efficiency than the wild type PULΔN5 showed a 2.3fold increase in kcat/Km values compared with the wide type D787C exhibited a 1.7fold increase in kcat/Km value and higher stability D328H/N387D/A414P showed an increase of 5 ° C in Tm and 12.9-fold t1/2 at 65 °C N680D/T477N showed improved thermostability The best variant K631V/ Q597S exhibited 5.6 folds increases in kcat/Km value The activities of M1 (ΔCBM41) and M5 (ΔCBM41ΔX25) were 2.9- and 2.4-fold that of the wild type, respectively L627R showed 117% activity and increased acid tolerance at pH 4.0 F581L, F581Q, F581R, F581T, F581V, and F581Y exhibited 3 °C increase in optimum temperature. F581L and F581V exhibited 1.6 and 1.7-fold kcat/Km values Thermal stability of R505P and acidic stability of R505E were enhanced. The kcat/Km

Reference [39]

[40]

[41]

[42]

[43]

[44] [45]

[46]

[47]

[48]

[49]

(continued)

3

Sequence, Structure, and Engineering of Microbial Starch Debranching Enzymes

53

Table 3.2 (continued) Enzyme

Pul

Source

Bacillus deramificans

Strategy

Rational design based on stable region prediction

N-terminal domain truncation

BtPul

Bacillus thermoleovorans

Combined usage of three computational design predictors (FoldX, I-Mutant 3.0, and dDFIRE)

Variant and effect values of G608V have been promoted by 49% D437H/D503Y showed 4.3 times half-life and 2.0-fold catalytic efficiency D437H/D503Y-T1 exhibited a 1.7-fold halflife of 203 h at 60 °C than D437H/D503Y G692M showed 3.8 °C increased Tm and 2.1fold half-life at 70 °C

Reference

[50]

[51]

[52]

This table was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9].

Fig. 3.4 Catalytic pocket of starch debranching enzymes (SDBEs)

The C-terminus of pullulanases has a β-sandwich structure organized as 5β/3β with a calcium-binding site (calcium binding site 5). The C-terminus of type II pullulanase possesses AamyC, fibronectin type III (Fn III), CBM20, and in some cases an S-layer homologous structural domain. The AamyC structural domain consists of a β-sheet and is found at the C-terminus of enzymes of the α-amylase family. This structural domain has been found to be involved in other substrate binding sites of the catalytic structural domain and in separating the α-glucan chains of starch, thereby destroying the starch granules. It is also involved in obtaining the correct orientation of the substrate at the active site of the enzyme. The fibronectin type III (FnIII) structural domain has a seven-stranded β-sandwich structure similar to the immunoglobulin structural domain. This structural domain is thought to play a role in substrate binding [22, 54].

54

W. Xia et al.

Fig. 3.5 Conserved sequence motifs of starch debranching enzymes (SDBEs). The preserved sequence motifs square measure known by a structure-guided multiple supermolecule sequence alignment. The chemical action triad Asp-Glu-Asp square measure is marked with an associate degree asterisk. Sequences were aligned by ClustalW, and similarity was rendered by ESPript 3.0 (https://espript.ibcp.fr/ESPript/ESPript/) [53]. This figure was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9]

The C-structural domain of neoaurobilin has the least sequence similarity to other enzymes of the α-amylase family. Conserved residues involved in substrate binding in the GH13 family are also evident in this subfamily. However, the C-structural domain has been found to stabilize the catalytic structural domain by protecting hydrophobic residues of the structural domain from aqueous solvents. In addition to the substrate binding sites found in the N-structural domain, the NA site (the interface between the structural domains N and A) and the A-structural domain, substrate binding sites were also found in the C-structural domain of T. vulgaris α-amylase I. Although the C and N structures of T. vulgaris α-amylase II exist separately from the rest of the protein, the C-structural domain has been shown to interact more with the catalytic structural domain than the N-structural domain. The C-structural domain of isoamylase folds into a right-handed parallel β-helix to form the catalytic structural domain, which consists of ten intact coils of three twisted parallel β-sheets and three incomplete coils surrounding the N and C termini of structural domain C.

3.4

Protein Engineering

In practical applications, high catalytic efficiency and thermal stability are desirable properties of SDBEs in starch liquefaction/glycosylation processes. However, due to biological adaptations and evolutionary limitations, natural enzymes still have some unavoidable defects, such as structural instability and low catalytic efficiency, despite the abundant resources of enzymes with different properties in microbial or genetic databases [58, 59]. Therefore, the use of rational or semirational design combined with stochastic mutagenesis strategies to improve the properties of SDBEs through targeted modifications is necessary to meet the application requirements and

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has become a hot research topic in this field [4, 54]. Some representative reports on protein engineering of SDBEs in recent years are shown in Table 3.2. Firstly, peptide truncation is a widely used modification method for SDBEs because of the variability and functional unknowability of multiple binding domains [60–62]. A variant of the combined truncated N- and C-terminal structures of the highly heat-stable type II pullulanase PulPY from Pyrococcus yayanosii CH1, Δ28 N + Δ791C, showed a further improved optimum temperature and nearly sevenfold specific activity compared to wild-type PulPY [63]. The N-terminal truncated rPulSL3ΔN from alkaline bacterial SL3 showed higher substrate affinity, improved thermal stability, and higher catalytic efficiency, while the C-terminal truncated rPulSL3ΔC lost both α-amylase and pullulanase activities [64]. Using an optimization strategy based on disorder prediction, truncated variants of B. naganoensis pullulanase, PULΔN5, PULΔN45, PULΔN78, PULΔN106, and PULΔC9, were designed and showed better activity and catalytic efficiency [65]. Chen et al. (2016) found that B. acidopullulyticus pullulanase (BaPul) variants of truncated M1 (ΔCBM41) and M5 (ΔCBM41ΔX25) exhibited both higher levels of secretory expression and higher activity by 2.9- and 2.4-folds, respectively, suggesting that deletion of the N-terminal structural domain may be more suitable for enhanced catalysis through industrial applications [66]. These cases suggest that reducing the number of structural domains or the size of the protein by truncation is an effective approach to improve thermal adaptation and other enzymatic properties of the abovementioned SDBEs. However, deletion of the N-terminal CBM68 structural domain of Anoxybacillus sp. WB42 pullulanase did not affect enzyme catalysis but significantly reduced thermal stability and substrate selectivity [60]. With the development of structural biology, structure-based designs are increasingly used for targeted modifications of enzymes, as they are very helpful in the localization of mutation sites. Phe581 was identified to be associated with the thermal stability and enzymatic activity of the pullulanase of Klebsiella mutants SHN-1, as several alternative mutations at this single site usually show higher optimal temperature and kcat/Km values [67]. Mutants R505P, R505E, and G608V improved the thermal stability, acid stability, and kcat/Km values of isoprenoidase from Bacillus lentilis JNU3 by single point mutations of residues near the important active site [39]. Four thermally resistant variants of pullulanase from B. deramificans (D503F, D503Y, D437H, and D437H/D503Y) were made by employing a stable region prediction strategy supported by similarity modeling. The half-lives of mutants D503F, D437H, and D503Y at 55 °C were one.6, 1.6, and 2.3 times greater than those of the wild sort, respectively. Moreover, there was an additive impact between these mutations because the double mutant D437H/D503Y showed a four.3-fold higher half-life and a twofold higher chemical action potency relative to the wild sort [40]. Moreover, the truncated variant D437H/D503Y-T1, which lacked the CBM41 structural domain at the N-terminal end, exhibited a 7-fold higher half-life of 203 h at 60 °C compared to the variant D437H/D503Y [41]. Saturation of residues at intervals in the active pocket of B. naganoensis pullulanase resulted in variant D787C with a one.7-fold increase in kcat/km values and better stability at 60 °C

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W. Xia et al.

and pH 4.0 [42]. The superimposed variants D328H/N387D/A414P of pullulanase PulB from B. naganoensis showed an improvement of 5 °C in melting temperature (Tm) and a prolonged t1/2 at 65 °C, which was 12.9-fold compared to the wild type [43]. Substitution of surface residues and introduction of disulfide bonds in the C domains with success improved the thermostability of the pullanase of Anoxybacillus sp. WB42 [44]. The superimposed variant K419R/T245C/A326C/ W651C/V707C of PulAC exhibited higher chemical process performance throughout starch saccharification with higher reaction temperature, higher catalytic efficiency values, and stronger thermal resistance at 65 °C. In addition, computer-aided rational design has attracted increasing interest within the last decade. In silico analysis of the natural philosophy of limiting dextrinase from H. vulgare by molecular dynamics (MD) simulations discovered that durable interactions such as H-bonds, salt bridges, and hydrophobic contact area units were weakened or lost at high temperatures, foreseen that seven salt bridge area units were important for the thermal resistance of the protein molecule, and provided new insight into the authentication of heat-sensitive regions in pullulanases [45]. A computationally assisted strategy combining the design tools FoldX, I-Mutant 3.0, and dDFIRE was developed for designing the thermal stability of pullulanase from Bacillus thermoleovorans [46]. Six dominant variants were identified from 17 designed mutants, and the best variant, G692M, exhibited better thermal stability with a 2.1-fold prolongation of T50 at 70 °C compared to the wild type. In an engineering strategy based on hydrogen bonding to alter the optimal pH and acid tolerance of BaPul by changing the pKa of the catalytic residues, the positive variant L627R showed better acid adaptation and effectively accelerated the starch glycation process [47]. The activity, thermal stability, and pH sensitivity of B. naganoensis pullulanase were modified by comutation of N680D/T477N through the “active hydrogen bonding network” (AHBN) approach, which focuses on the hydrogen bonding interactions between residues located in or around the active center. Mutations in residues at the periphery of the AHBN may exert beneficial effects, while mutations within the AHBN may inhibit activity [48]. Recently, a coevolutionaryguided target site identification method, called “evolutionary coupled saturation mutagenesis” (ECSM), was developed to enhance the activity of B. naganoensis pullulanase [49]. The evolutionary coupling (EC) tool offered correct data regarding the pairs of residues that are in contact within the three-dimensional structure. They calculated the biological process coupling (EC) scores within intra- and inter-domain mistreatment using the EVcouplings server (http://evfold.org) and the EVcomplex server (https://evcomplex.hms.harvard.edu), respectively. Seven residue pairs with scores higher than 1.0, particularly D614/H539, E530/T520, D541/D473, E777/T730, K631/Q597, V328/I565, and Y392/Y571, were chosen for mutation. Seven double mutants showed a pair of 6- to 5.6-fold increases in kcat/km compared to the wild type among all 1056 double variants, with K631V/Q597S being the best. However, the quadruple variants obtained from the mixture of those double mutants did not cause an additional increase in activity [49]. Undoubtedly, these studies provide a

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good guide for identifying possible hotspots and selecting mutation targets to modify other SDBEs. Future SDBE research should include three valuable goals. First, with the help of emerging fields such as bioinformatics and computational biology, we will continue to explore and design enzymes that are suitable for a variety of newly developed enzyme combinations and processing conditions. The second is to identify and eliminate bottlenecks in the preparation of high-yield enzymes by optimizing host cells and developing new fermentation strategies. Third, study the content of different SDBEs in different sources. Hydrolysis ability and application performance of 1,6 glucoside-linked polysaccharides. Further research on these issues is expected to promote the industrial application of SDBEs. Acknowledgments This chapter was modified from the paper published by our group in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [9]. The related contents are reused with permission.

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Chapter 4

Production and the Applications in Preparation of Branched Sugar Products of Starch Debranching Enzymes Wei Xia, Sheng Chen, and Jing Wu

Abstract Starch debranching enzymes (SDBEs) are bidirectional catalytic enzymes that can catalyze pullulan, starch, and other branches of polysaccharide substrates. α-Hydrolysis of 1,6-glycosidic bonds can also be catalyzed under certain conditions. α-Reverse synthesis of 1,6-glycosidic bonds. Therefore, it can be used in many fields, such as saccharification and degradation of starch raw materials, debranching, and reverse synthesis of branched sugar products. Because the SDBE strain obtained by direct screening has a low fermentation yield in nature, it is difficult to separate pure protein, and the production cost is high, which cannot meet the needs of industrial production. Therefore, heterologous expression of SDBEs through genetic engineering has become a research hotspot in this field. The most commonly used SDBE gene expression systems are Escherichia coli, Bacillus, and Pichia pastoris. Keywords Starch-debranching enzymes · Pullulanase · Isoamylase · Enzyme production

4.1

Introduction

Debranching enzymes are widely used in the production of ethanol fuel, cyclodextrin, resistant starch, maltotriose syrup, and other products, which are in great demand in industrial applications. However, pullulanase, the main producer of debranching enzymes, is mainly Novozymes of Denmark, with high cost. It is also produced in China by Kingsley and Longcote, with a small proportion. The current research reports that pullulanase is highly expressed in Escherichia coli, Bacillus subtilis, and Bacillus subtilis. However, in principle, neither Escherichia coli nor W. Xia · S. Chen · J. Wu (✉) State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, 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. 2023 J. Wu, W. Xia (eds.), Industrial Starch Debranching Enzymes, https://doi.org/10.1007/978-981-19-7026-9_4

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Bacillus subtilis are considered safe microorganisms. Bacillus subtilis has food safety problems and a long fermentation time caused by methanol. Pullulanase (EC 3.2.1.41) is a widely used starch debranching enzyme. The expression of pullulanase in the two wild-type bacteria was very low, and the enzyme activities were 0.171 and 3.86 U
mL-1 in Bacillus flavothermus KWF-1 [1] and Anoxybacillus SK3–4 [2]. With the development of DNA recombination technology, pullulanase has been expressed in many microbial hosts, including Escherichia coli [3], Bacillus subtilis [4–6], and Bacillus licheniformis [7], and its highest extracellular enzyme activities are now 1567.9, 24.5, 18.0, 1164.8, and 2101.3 U
mL-1. However, due to the larger molecular weight and more complex structure, many pullulanase sources have some folding problems when expressed in host bacteria. When pullulanase from Bacillus delamifica is expressed in Escherichia coli, a large number of inclusion bodies are formed in the cytoplasm gap, and the efficiency of extracellular secretion is very low. Therefore, the safe and efficient expression of debranching enzymes is of great significance for their industrial application. In addition, it can specifically cleave the α-glycosidic bond. In addition to the 1,6-glycosidic bond, pullulanase also has a strong reverse synthesis ability. This property of pullulanase can be used to synthesize new starch deep processing branched sugar products, such as branched maltooligosaccharides and branched cyclodextrins, and has broad application prospects.

4.2

Production of SDBEs

In view of the importance of SDBEs in the starch process business, an outsized range of studies have been conducted to explore the consequences of physical and biological process parameters on the assembly of untamed microorganisms manufacturing debranching enzymes. Although good stress has been placed on cistron biological research and expression and continuous fermentation to obtain a high titer, optimizing culture conditions continues to be efficient thanks to improving yield. The effects of various chemical parameters, together with carbon and chemical element sources, surfactants, metal ions, temperature, pH scale, and stirring, on protein production were studied. The foremost necessary physical parameters moving biological growth and protein production are temperature, pH, stirring, and pressure. Generally, pullulanases are produced by microorganisms during a comparatively wide temperature range, and therefore, the pH scale worth of the expansion medium has a profound impact on the assembly, stability, and activity of living thing enzymes. With the growth of industrial demand, heterologous recombinant expression has become increasingly crucial for the industrial application of SDBEs because the low enzyme production capacity of wild microorganisms usually cannot meet the needs of practical processing [6, 8–15]. Studies have long been utilized to increase the output of SDBEs with sensible industrial application potential. Within the past decades, Escherichia coli or B. bacillus strains and yeasts have been widely used

4

Production and the Applications in Preparation of Branched Sugar. . .

63

as hosts for secreting and expressing recombinant pullulanase and isoamylase. However, aside from a number of studies, most of the reportable bodily fluid expression levels are still not up to a hundred U/mL, which is sometimes the case (Table 4.1) [16–21]. To enhance the matrix secretion level of enzymes, it is vital to investigate the mechanism that affects the animate thing secretion level of SDBEs.

4.2.1

Heterologous Expression of SDBEs in E. coli Strains

With the advent of the proteome era, research on protein structure and function has been further deepened in recent years, and the application scope of recombinant protein technology has become increasingly extensive. The Escherichia coli expression system is the most widely used protein expression system at present. It has a clear genetic background, is easy to cultivate and control, has a simple conversion operation, has a high expression level, has a low cost, and has a short cycle. At the same time, it is a commonly used and economical protein expression system, which was the earliest developed gene expression technology. To date, the E. coli expression system is still considered an important and widely applicable tool in the field of protein expression. However, the productivity of the economic pET plasmid system and Escherichia coli BL21 (DE3) expression host in the production of pullulanase from Bacillus cereus Nws-bc5 and thermophilic bacteria was unsatisfactory [16, 34, 41]. The results showed that the composition of promoters, operons, and alternative components of the expression cellular inclusion might have an effect on the recombinant and iatrogenic expression of eubacteria acidophilus BaPul13A in E. coli. Mistreatment of the vector pET-28a (+), which carries components composed of animal product operator, lacI cistron, and T7 promoter, the very best soluble macromolecule yield was raised to 1156 U/mL [23] by strictly dominating the essential expression strategy in a 5-L fermentation tank. Additionally, a series of optimization strategies have been developed to enhance the secretion level. In addition, 412 U/mL Thermotoga lettingae TMO pullulanase was produced in Escherichia coli BL21 Coden and (DE3) host by optimizing the soundness conditions of a dissolved chemical element (DO) [40]. Through N-terminal truncation [29], addition of Triton X-100 wetting agent [28], supplementation of natural diffusion answer [30], and glycine-fed batch culture [3], the living thing activity of pullulanase from attenuated bifidobacteria was raised to 502, 814.2, 879.3, and 2523.9 U/mL, respectively. Within the case of overexpression of isoamylase from Bifidobacterium lugens in Escherichia coli MDS42 by optimizing induction conditions and fermentation methods, the best rumored expression level of SDBEs was 22983.0 U/mL activity and 18.8 mg/mL macromolecule [42].

B. subtilis WS9

B. choshinensis B. choshinensis E. coli BL21(DE3) E. coli BL21(DE3) E. coli BL21(DE3) E. coli BL21(DE3) B. subtilis ATCC6051.

B. deramificans

B. naganoensis

B. subtilis WB800 B. subtilis WB800 E. coli BL21(DE3)

B. subtilis WB600 B. subtilis WB600 B. subtilis WB600

E. coli BL21(DE3)/ pLysS B. subtilis 5951.8 543.0 1005.8 814.2 502.0 2523.9 963.9 625.5 26.5 24.5 2.8

Protease deletion and optimal feeding Addition of MgCl2 to the medium Response surface methodology Adding surfactants TritonX-100 N-terminal truncation Glycine feeding Supplement with natural osmolytes Protease deletion, promoter optimization Enhancer regulation Screening promoter and host Optimizing promoter and signal peptide

60.9 102.8 580.0

90.7

Optimizing dual promoter

pHY300PLK/ PHpaII-PamyQ pHY300PLK/ PHpaII-PamyQ pNCMO2 pNCMO2 pET-20b(+) pET-20b(+) pET-20b(+) pET-20b(+) pBE-MCS/PamyLPspovG pMA0911/PsacB pMA0911/P43 pBE-Papr/ SPsacB-pulB PHpaII-pul PHpaII-pul pET-22b(+) Optimizing expression conditions Optimized feeding strategy Autoinduction by lac operator

4.8

–c

pET-42

pET-20b(+)

E. coli BL21(DE3)

504.0

Yield (U/mL)

1156.3

pWB980

B. subtilis WB600

Strategy Using a high copy number and highly stable shuttle plasmid Tight control of basal expression

Vector

Host

B. cereus Nws-bc5

Gene source Pullulanase Anoxybacillus sp. LM18–11 B. acidopullulyticus

Table 4.1 Recombinant expression of starch debranching enzymesa

3.2 2.4 41.4

5.9 6.3 –

5.4 10.8 46.2 2.9 1.2 8.3 1.48

[32] [33] [34]

[17] [18] [19]

[26] [27] [28] [29] [3] [30] [31]

[25]

[24]

– 65.6

[16]

[23]

[22]

Reference

3.28/ OD600 –

5.5

Fold changeb

64 W. Xia et al.

E. coli MDS42 P. pastoris GS115 E. coli JM109 Saccharomyces cerevisiae

B. subtilis BS001 E. coli BL21(DE3) E. coli BL21-Coden Plus (DE3)-RIL E. coli BL21(DE3)

P. pastoris SMD1168 P. pastoris KM71H P. pastoris KM71H P. pastoris X-33

pSX2 pGAPZαA pHIF2 and pMIF3 pGK8

pET-28a(+)

pPIC9k pPICZαA pPICZαA pPICZαA, pGAPαA pCBS35 pET-28a(+) p ET15b 13.8 22983.0 318.0 90.0 86.0

Optimization of induction Continuous high-density fermentation – –

350.8 8.5 15.0 33.5 28.6 1555.0 10.0 412.0

Strong promoters combination – Optimizing dissolved oxygen (DO)-stat protocol –

– – – –

b

[42] [43] [20] [21]

[41]

– 54.0 3.5 – –

[38] [39] [40]

[35] [36] [36] [37]

21.9 – 2.3

– – – –

This table was reused with permission from the paper published in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [44] The fold modification means that the magnitude relation of the catalyst expression level thereto of the native strain or in restraint condition c The dashed image represents there’s no relevant information

a

Bacillus subtilis BK07 B. subtilis PY22 Geobacillus thermoleovorans NP33 PulA Klebsiella variicola Thermotoga lettingae TMO Thermus. thermophilus Isoamylase Thermobifida fusca B. lentus JNU3 F. odoratum KU P. amyloderamosa

4 Production and the Applications in Preparation of Branched Sugar. . . 65

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4.2.2

W. Xia et al.

Heterologous Expression of SDBEs in Bacillus Strains

For the past few years, Bacilli strains such as Bacillus subtilis and B. choshinensishas have become candidate expression hosts for food industry proteins. Bacillus subtilis, as an endophytic spore Gram-positive bacterium, is an ideal host for the expression and secretion of foreign proteins in prokaryotic expression systems and an important model strain in the field of microbial research, with high application value because of its nonpathogenic nature, strong ability to secrete proteins, good fermentation foundation, and production technology. With the rapid development of biotechnology applications, the demand for an optimal expression system is growing. Compared with the Escherichia coli expression system, the B. subtilis expression system has the advantages of high protein production, good biological activity, low endotoxin content, and good egg white solubility. Compared with the eukaryotic expression system, its expression cycle is short, the cost is low, and the expression amount is high. Moreover, B. subtilis is generally considered a biosafety strain and an ideal expression host for protein production in important pharmaceutical and food industries, such as α-amylase, protease, xylanase, lipase, and β-food enzyme preparations such as glucanase. Bacilli strains are also widely used as expression hosts for the production of SDBEs. It was reported that a novel fusiform plasmid with high copy number and high stability was used to increase the production of Anoxybacillus sp. LM18–11 pullulanase on the host Bacillus subtilis WB600, resulting in a high yield of 504 U/ mL [22]. Moreover, new host strains constructed by removing proteases showed better performance. By binding strong promoters extracted from the host cell genome, a recombinant Bacillus subtilis BS001 strain containing the combined promoter (PsodA+fusA+amyE) produced 1555 U/mL PulA in a 50-liter fermentor, 21.9 times the flask level [38]. By adding MgCl2 and response surface optimization, the expression activity of B. deramificans pullulanase in B. choshinensishas was increased to 543 U/mL and 1005.8 U/mL, respectively [26, 27]. In addition, the safety of B. subtilis (GRAS), which is generally recognized as safe, makes it more suitable for the efficient expression of starch dedendriase. However, similar to the commercial Escherichia coli system, the commercial host strains Bacillus subtilis WB600 or WB800 have different expression strategies for Bacillus longvalley pullulanase, and their production is still below 60 U/mL [17–19, 32, 33]. The optimized Bacillus subtilis ATCC6051 was expressed with the pammyl-pspovg plasmid [31]. Eight extracellular proteases were removed, and the production of Bacillus Nagasaki pullulanase was increased to 625.5 U/mL. Similarly, using the dual-promoter PHpaII-PamyQ, B. deramificans pullulanase was overexpressed in the modified B. subtilis WS9 with nine protease deletions, and the production reached the highest of all reported pullulanases to our knowledge of 5951.8 U/mL [24, 25].

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4.2.3

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Heterologous Expression of SDBEs in Yeast Strains

An efficient expression system in the field of biomedicine is crucial for the production of recombinant proteins. With the rapid development of gene engineering technology and protein heterologous expression, an increasing number of expression systems have been established and widely used. As a food-grade eukaryotic microorganism, Saccharomyces cerevisiae has the characteristics of rapid reproduction, simple culture, convenient genetic operation, etc. It is one of the ideal expression systems for producing recombinant proteins. Pichia pastoris has become one of the most successful foreign egg white expression systems due to its clear genetic background, simple operation, rapid growth, inexpensive culture, suitability for high-density fermentation, and the advantages of posttranslational modification and processing systems of eukaryotic cells. It is widely used for the expression of foreign proteins. In the past 20 years, yeast expression systems have experienced rapid development and received extensive attention, which has produced great economic and social values. On the other hand, the yeast expression system is affected by many factors, including the characteristics of the foreign gene itself, the characteristics of the strain, the expression environment, and the fermentation process, and its expression of different foreign proteins also show great differences. Recently, yeast strains have also been used as good hosts for highly efficient production of heterologous SDBEs. Earlier studies found that integrating Pseudomonas amyloderamosa isoamylase into the recombinant S. cerevisiae strain created only 86 U/mL of low activity [21]. In contrast, isoamylase from Bacilli JNU3 was produced by P. pastoris. Through continuous high cell density fermentation, the output of animate thing isoamylase reaches a high value of 318 U/mL [43]. Additionally, under the action of the promoters PGAP and PAOX1, the open-chain enzyme Gt apu of Bacilli thermophilus. The animate expression of C in Pichia pastoris was 33.5 U/mL and 28.6 U/mL, respectively. However, different studies have shown that inducible promoters also contribute to achieving high yields. As a representative, the comparatively high animate actinomycete pullulanase production of 350.8 U/mL was successfully achieved in Pichia pastoris SMD1168 solely at the shake flask level through alcohol induction [35]. In summary, some valuable advances have been made in this field in recent years, mainly focusing on optimizing the expression elements and secretion pathways of host cells, optimizing the high-density fermentation strategy, and removing N-terminal peptide chains or domains [45, 46]. The drawback was that the assembly of enzymes with excellent industrial properties, such as BaPulA from eubacteria acidophilus, remains meagre. Humor expression, particularly the safe and economical production of food-grade expression systems, remains a drag that warrants attention within the application of SDBEs, which requires additional analysis and development.

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Conclusions

In recent decades, SDBEs have attracted extensive research interest as important biocatalysts for starch processing and downstream product preparation. SDBEs, represented by pullulanase, are one of the most important debranching enzymes widely used in the starch processing industry. They are mainly used in combination with other starch decomposing enzymes and are widely used in starch saccharification as debranching enzymes to improve sugar production. In addition, SDBEs are also used in other industrial processes, such as baking enzymes in food, saccharifying starch into glucose and glucose syrup, maltose and maltotriose syrup, the brewing industry, and the detergent industry. In recent years, previous studies on the identification and characterization of enzymes have enriched resources and can be used for various applications with different needs. To date, various SDBEs have significant similarities and differences in catalytic performance. Among them, thermostable enzymes are receiving considerable attention because they are active under industrial conditions. From an industrial point of view, the important application features of SDBEs such as pullulanase and isoamylase include high thermal stability, functions in a wide pH range, and activities independent of calcium ions. The bond type specificity of catalytic hydrolysis between α-1,6 and α-1,4 bonds is also particularly important. For the existing research, another important consideration for the practical application of SDBEs is that the yield of heterogenous production is usually kept at a relatively low level, except for a few prominent cases, especially for thermostable enzymes with great potential from extreme microorganisms. Because enzyme production in natural bacterial and archaeal hosts is generally low, the expression of SDBEs in heterogeneous hosts has attracted the interest of a large number of researchers. These commonly used hosts mainly include Escherichia coli, Bacillus subtilis, and Pichia pastoris. These studies on fermentation production show great application value and should be considered the most important link to industrialization. Acknowledgments This chapter was modified from the paper published by our group in Biotechnol Adv. (Xia W, Zhang K, Su L, Wu J. 2021) [44]. The related contents are reused with permission.

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Chapter 5

Recombinant Expression of Starch Debranching Enzymes in Escherichia coli Kang Zhang, Zhanzhi Liu, and Jing Wu

Abstract Starch debranching enzymes have been recombinantly expressed in Escherichia coli, which has the advantages of a clear genetic background, simple molecular operation, easy cultivation, short fermentation cycle, and low fermentation cost. While starch debranching enzymes, especially pullulanase, are prone to form inclusion bodies in the cytoplasm of recombinant E. coli strains, many strategies have been used to improve their soluble expression. Thus, in this study, kinds of factors affecting the recombinant expression of starch debranching enzymes in E. coli, such as supplements, enhancers, and induction regulation were investigated. Pullulanase mutants were constructed, and the effects of the mutation on the thermal stability and catalytic efficiency of the enzyme were investigated. The N-terminal domain of pullulanase was truncated to different degrees to investigate the effect of domain truncation on the secretion efficiency, enzymatic properties, and application performance of pullulanase. Isoamylase was recombinantly expressed in different E. coli host strains. Keywords Escherichia coli · Host strains · Starch debranching enzymes · Fermentation cultivation condition · Mutants · Domain truncation

5.1

Introduction

With the wide application of pullulanase in starch processing, the demand for pullulanase is gradually increasing. How to realize the efficient production of pullulanase has become a hot spot of research and development? In recent years, scholars at home and abroad have mainly carried out screening, mutation, and K. Zhang · Z. Liu · J. Wu (✉) State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, 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. 2023 J. Wu, W. Xia (eds.), Industrial Starch Debranching Enzymes, https://doi.org/10.1007/978-981-19-7026-9_5

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fermentation condition optimization of pullulanase-producing wild strains. Although the enzyme production level can be improved by means of mutation and fermentation optimization, the improvement range is still very limited due to the strict regulatory mechanism of wild strains. As one of the most effective methods of protein overexpression, genetic engineering has been widely used in the industrial production of various enzyme preparations. To improve the enzyme production level, domestic scholars have tried to improve the activity unit of pullulanase by means of genetic engineering combined with fermentation optimization [1]. The Escherichia coli expression system has the advantages of a clear genetic background, simple molecular operation, easy cultivation, short fermentation cycle, and low fermentation cost [2]. E. coli is the most widely used and deeply studied expression system at present because of its rapid synthesis of heterologous protein and high yield, and the expression of recombinant protein can exceed 30% of its total protein. At present, many commercial proteins are produced on a large scale with E. coli as the host [3, 4]. However, proteins often form inclusion bodies when recombinantly expressed in E. coli, especially for pullulanase. It has been reported that pullulanases from various sources easily form inclusion bodies when expressed in E. coli [5–7]. The possible reason is that pullulanase has a large molecular weight, complex structure, and a strong tendency to form aggregates. At present, many methods have been reported to promote soluble expression of recombinant protein in E. coli, mainly including recombinant protein modification, host strain modification, expression plasmid element optimization, and cultivation condition optimization. First, studies have shown that the molecular weight and structural complexity of proteins have a great influence on their soluble expression and secretion efficiency. The larger the molecular weight and the more complex the structure of proteins, the easier they are to form inclusion bodies, and the lower the extracellular secretion efficiency [8]. In 2001, an enzyme biosystems company reported in a patent that removing 106 amino acids from the N-terminus of B. naganoensis pullulanase could increase the exocrine level of the recombinant enzyme by 1.6-fold [9]. Researchers believe that this may be caused by the decrease in the molecular weight of the truncated protein. Second, studies have shown that coexpression of chaperones can help or promote the correct folding of proteins and effectively reduce the probability of inclusion body formation due to misfolding of recombinant proteins [10, 11]. Using some modified E. coli stains as hosts, such as glutathione reductase mutants or thioredoxin reductase mutants, can help the recombinant protein form disulfide bonds in the cytoplasm, which shows an obvious effect in expressing some proteins with disulfide bonds. Moreover, a weak promoter, low culture temperature, and induction intensity were used to express the target protein, which reduced the speed of protein synthesis and promoted the correct folding of the target protein. Adding glycine, surfactants, or some metal ions to the medium may enhance the permeability of the cell membrane and promote protein secretion. Li et al. increased the extracellular expression of α-CGT by more than 11 times by adding 150 mM glycine and 20 mM Ca2+ [12]. Secretion of recombinant protein into the periplasmic space or fermentation broth can also promote soluble protein expression. This is because the oxidation of the periplasmic space is helpful for the correct

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folding of the precursor protein, while the proteolytic enzymes in the periplasmic space are much less abundant than those in the cytoplasm, which can avoid the degradation of the protein. Compared with the periplasmic space, the space in fermentation broth is wider, which can avoid the accumulation of proteins in cells and form inclusion bodies [13].

5.2

Effect of Fermentation Conditions on Soluble Expression of Recombinant Pullulanase

In a previous study, we constructed genetically engineered bacteria producing pullulanase and carried out molecular modification of natural pullulanase. After transformation, the pullulanase mutant can be well mixed with glucoamylase. To improve the fermentation unit of pullulanase, it is necessary to optimize the fermentation conditions. In this study, the shaking flask fermentation conditions of the recombinant strain were optimized, and then the high-density 3-L fermentation control technology of the recombinant strain was optimized.

5.2.1

Recombinant B. deramificans Pullulanase Easily Formed Active Protein Aggregates

The results of shake flask fermentation showed that the activity of pullulanase was measured after 36 h of induction with 0.4 μmol/L IPTG at 37 °C. The extracellular enzyme activity was only 3.2 U/mL, and the intracellular enzyme activity was 9.9 U/ mL (Fig. 5.1a). SDS–PAGE images showed that there were a large number of pullulanase proteins in the insoluble part of the cells, which may be inactive “inclusion bodies” (Fig. 5.1b); however, no “inclusion bodies” were found in the cells without the expression vector. In addition, electron microscopy images showed that the induced recombinant cells contained a large number of “inclusion bodies,” occupying a large space of cells (Fig. 5.2a). During the processing of the fermentation sample of the recombinant strain, it was found that pullulanase activity still existed in the precipitate after the cell was broken, and the activity of the precipitate part (converted into enzyme activity per mL fermentation broth) reached 28.3 U/mL, which was 2.1 times the soluble enzyme activity (the sum of extracellular and intracellular supernatant) of the recombinant enzyme. This phenomenon is different from the recombinant expression of a variety of enzymes (isoamylase, α-amylase, α-CGTase, cutinase, and amidase) studied by our laboratory. Although there are different degrees of insoluble proteins in cell fragments after expression of the above other enzymes, they are not active. This phenomenon of insoluble proteins with activity may be a special phenomenon of pullulanase.

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Fig. 5.1 Pullulanase activity distribution (a) and SDS–PAGE analysis (b) of E. coli BL21(DE3) (pET24-ompA/pulA). Reused with permission from Bioresource Technol. Duan X, Chen J, Wu J. 2013. Lane 1, periplasmic protein of E. coli containing pET24-ompA/pulA; Lane 2, insoluble protein of E. coli containing pET24-ompA/pulA; Lane 3, periplasmic protein of E. coli containing pET24-ompA without the pullulanase-encoding gene; Lane 4, insoluble protein of E. coli containing pET24-ompA without the pullulanase-encoding gene; Lane M, protein molecular weight markers

Fig. 5.2 Transmission electronic microscopy observation of intracellular localization of the recombinant pullulanase in E. coli BL21(DE3)(pET24-ompA/pulA without (a) and with glycinebetaine (b). Scale bars 0.2 μm

The amino acid sequence of B. deramificans pullulanase was analyzed by AGGRESCAN software [14], and it was found that it had obvious characteristics of forming active protein aggregates (Table 5.1). There are 21 hot spots in the

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Table 5.1 Prediction of the aggregation tendency of different pullulanases overexpressed in bacteria Sources Bacillus sp. m5-13 Bacillus cereus E33L Pullulanibacillus naganoensis B. deramificans Bacillus acidopullulyticus Klebsiella pneumoniae Thermotoga maritima

Length (AA) 1331 780 926 928 921 1030 773

Na4vSSa -16.50 -15.10 -11.9 -11.1 -9.8 -9.2 -2.2

Aggregation tendency ++ ++ ++ ++ + + Aggregate

Na4vSS characteristic values corresponding to different morphological proteins: inclusion body, 2.51; natural globular protein, -4.26; soluble protein, -5.18; protein aggregate, -12.96; unfolded protein, -28.73

a

structure, 11 of which are located on the protein surface: 162–173, 234–244, 334–353, 6–13, 21–26, 268–274, 372–381, 810–820, 865–871, 878–881, and 911–919. They are mainly distributed in the X25, X45, CBM48, and C domains, especially the CBM48 domain. In addition, the amino acid sequences of several pullulanases in the database were analyzed. Most of the pullulanases had the tendency to form active protein aggregates to varying degrees (Table 5.1), which may be an important reason for the low expression of pullulanase in E. coli. The above software analysis showed that B. deramificans pullulanase had the characteristics of naturally forming protein aggregates, and the experimental results further confirmed that the pullulanase was active in the cell precipitation part, so it was speculated that the recombinant enzyme in the precipitation part was the active protein aggregate. Studies have shown that when a large number of heterologous proteins are overexpressed and accumulate in cells, great metabolic pressure will be placed on host cells. If the molecular chaperones responsible for protein folding in cells have difficulty coping with the pressure caused by protein overexpression, overexpressed proteins will not fold correctly and eventually aggregate to form insoluble protein aggregates. Protein aggregates can be divided into partially misfolded, active protein aggregates and misfolded, inactive inclusion bodies. Ramshini et al. found that the larger the molecular weight and the more complex the structure of the protein, the easier it is to form protein aggregates [8]. In addition, Ivanova et al. found that the amino acid sequence of a protein determines whether it is easy to form protein aggregates. The phenomenon that B. deramificans pullulanase in this study easily forms protein aggregates may be related to many factors, such as the natural properties of the amino acid sequence of the protein, the complexity of the protein structure, and the high molecular weight [15]. As the formation of protein aggregates will seriously reduce the yield of soluble protein, improve the complexity of subsequent protein separation and purification processes, and increase the production cost, improving the solubility of recombinant protein and reducing the formation of protein aggregates has become one of the severe problems faced by recombinant protein expression.

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Effect of Induction Temperature on Fermentation of the Recombinant Strain

Induction temperature is one of the important factors affecting the soluble expression of recombinant protein [16–18]. In general, induced expression of recombinant E. coli at low temperature can reduce the formation of inclusion bodies/protein aggregates and increase the proportion of soluble recombinant proteins. To investigate the effect of temperature on the fermentation of recombinant pullulanase, the recombinant strain was induced and cultured at 25, 30, and 37 °C, and the results were analyzed. As shown in Fig. 5.3, with increasing culture temperature, the activity of soluble pullulanase (extracellular and intracellular supernatants) decreased gradually. When the temperature was 37 °C, the cell biomass was the highest, and the dry cell weight reached 5.6 g/L, which was 1.5 times and 1.3 times the dry cell weight at 25 and 30 °C, respectively; however, the pullulanase activity in the extracellular and intracellular supernatants of the recombinant strain was only 2.1 and 10.2 U/mL, respectively. When the induction temperature was 30 ° C, although the dry cell weight decreased to 4.3 g/L, the extracellular enzyme activity and intracellular enzyme activity increased to 9.0 and 14.3 U/mL, respectively. When the induction temperature was 25 °C, the activity of soluble pullulanase reached 36.5 U/mL, which was 3.0 times that at 37 °C. Garcia-Fruitós et al. found that low temperature could significantly improve the quality of recombinant protein in the process of recombinant expression of two fusion proteins VP1-GFP and 6

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0 25

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Fig. 5.3 Effects of culture temperature on cell growth and soluble pullulanase activity. DCW (black), extracellular pullulanase activity (white), periplasmic pullulanase activity (light gray). Reused with permission from Bioresource Technol. Duan X, Chen J, Wu J. 2013

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VP1-LAC; however, when cultured at high temperature, the folding efficiency of recombinant protein decreased, and it was easier to form protein aggregates [19]. To increase the correct folding rate of recombinant pullulanase and increase the soluble expression level of pullulanase, the induction temperature should be controlled according to the experimental requirements in the following experiments.

5.2.3

Effect of Inducer Concentration on Fermentation of the Recombinant Strain

The expression vector pET24-ompA was used in this study. The T7 promoter of pET24-ompA was the most widely used promoter in the pET expression system. The T7 expression system is under strict regulation by the LacI repressor protein. When not induced, the T7 RNA polymerase gene is not transcribed, and the T7 promoter is silent; when an inducer is added for induction, the repressor protein binds with the inducer and changes conformation, which separates from the manipulation gene and activates the transcription and expression of T7 RNA polymerase. Then, target gene expression under the control of the T7 promoter is activated. The commonly used inducers are IPTG and lactose. IPTG is strong but expensive and is generally used in small-scale experiments; lactose induction intensity is low, and the price is inexpensive, so it can be used in large-scale fermentation experiments. The concentration of inducer has an important influence on the expression level of recombinant protein, especially when the target protein easily forms protein aggregates. When the concentration of inducer is too high, the target protein usually exists in an insoluble state. Therefore, it is necessary to investigate and optimize the concentration of inducer. The recombinant strain was first cultured at 37 °C for 2 h and then cooled to 25 °C when the OD600 reached approximately 1.0. Then, IPTG at final concentrations of 0, 0.025, 0.05, 0.1, 0.2, and 0.4 mmol/L was added to the medium for 36 h. Then, the dry cell weight and the pullulanase activities in the extracellular and intracellular supernatants were determined. As shown in Fig. 5.4, with increasing IPTG concentration, the dry cell weight gradually decreased. When the IPTG concentration was greater than 0.1 mmol/L, the decline rate of cell biomass accelerated. When the concentration of IPTG was 0.4 mmol/L, the dry cell weight was only 2.5 g/L, which was 62.5% of the dry cell weight without IPTG. These results showed that a high concentration of IPTG could significantly inhibit the growth of the cells because IPTG was toxic to the recombinant cells. When the concentration of IPTG was between 0 and 0.05 mmol/L, the activity of pullulanase in the extracellular and intracellular supernatants increased gradually with increasing concentration of inducer. The maximum activities of extracellular and intracellular enzymes were 13.0 and 23.0 U/mL, respectively. When the concentration of IPTG was higher than 0.05 mmol/L, the activity of pullulanase decreased with increasing IPTG concentration. When the IPTG concentration

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reached 0.4 mmol/L, the extracellular and intracellular enzyme activities were only 1.0 and 5.2 U/mL, respectively, which was similar to the results reported by Jhamb et al. [20]. Jhamb et al. found that the concentration of IPTG had an important effect on the solubility of the recombinant protein. When the concentration of IPTG was reduced to 0.01 mmol/L, the soluble ratio of xylanase increased significantly, and the proportion of insoluble protein decreased. The reason for the above phenomenon may be that when induced by a high concentration of IPTG, a large number of target proteins were produced rapidly in the cells of recombinant strains, which formed a crowding effect in the narrow cells. If these proteins could not be transported to the periplasmic space in time and folded correctly with the help of molecular chaperones, they would accumulate each other and form aggregation cores. Finally, with the increase in aggregated proteins, insoluble protein aggregates were formed. Meanwhile, a low concentration of inducer can reduce the protein expression speed, and the protein concentration inside the cell is maintained at a low level. The cell itself can basically meet the requirements of protein secretion and folding. The synthesized protein is gradually transported to the periplasmic space and folded to form soluble active protein.

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Effect of Osmotic Pressure Regulator on Fermentation of Recombinant Strain

The results in Sects. 5.2.2 and 5.2.3 show that the total activity of soluble pullulanase is only 36.2 U/mL even if the temperature and induction intensity are reduced, and a large number of protein aggregates will still be formed in the cells; the addition of surfactants in shake flasks can unfold and dissolve some aggregates, and some unfolded proteins can refold to form active forms [14]. However, surfactants easily form foams. If they are added and ventilated in the fermenter, the amount of foam will be even larger, which will be difficult to control. The formation of foam may cause liquid fluid and bacterial contamination in the fermentation fermenter. Moreover, most surfactants (such as Triton X-100) are harmful to human health and cannot be directly used in the production of food enzymes. De Marco and Ignatova found that some small molecules of osmotic pressure regulators naturally existing in cells cannot only significantly improve the thermal stability of some proteins but also promote the correct folding of proteins [21, 22]. These small molecular osmotic pressure regulators are known as “chemical molecular chaperones” because of their unique functions of improving the thermal stability of natural proteins and promoting the refolding of unfolded peptides. In this study, three osmotic pressure regulators (proline, potassium glutamate, and betaine) were selected to investigate their effects on the soluble expression of pullulanase.

5.2.4.1

Effects of Osmotic Pressure Regulators on the Fermentation of the Recombinant Strain

The recombinant strain was cultured at 37 °C until the OD600 reached approximately 1.0 and cooled to 25 °C. Then, osmotic pressure regulators (20 mmol/L proline, 20 mmol/L proline+0.5 mol/L NaCl, 20 mmol/L potassium glutamate, 20 mmol/L potassium glutamate+0.5 mol/L NaCl, 20 mmol/L betaine, and 20 mmol/L betaine +0.5 mol/L NaCl) and 0.05 mmol/L IPTG were added to the culture medium for 36 h. Then, the dry cell weight and the activity of pullulanase in extracellular and intracellular supernatants were measured. As shown in Fig. 5.5, pullulanase activity in extracellular and intracellular supernatants of four groups of samples with added proline, proline +0.5 mol/L NaCl, potassium glutamate, and potassium glutamate +0.5 mol/L NaCl in the culture medium decreased to varying degrees. The total activity of soluble pullulanase (the sum of extracellular and intracellular enzymes) of the sample supplemented with betaine alone was 41.6 U/mL, which was 1.3 times that of the control group. Moreover, betaine also promoted the growth of cells. It can be seen from the transmission electron microscope picture (Fig. 5.2b) that there is no obvious protein aggregate in the cells induced by IPTG, which proves that betaine can effectively promote the correct folding of recombinant pullulanase and improve the soluble expression of pullulanase. When betaine and NaCl were added together, the total soluble recombinant enzyme activity decreased slightly, and the

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dry cell weight also decreased significantly. This phenomenon may be because TB medium already contains many inorganic salts (2.31 g/L KH2PO4 and 16.43 g/L K2HPO4), which can generate enough osmotic pressure to promote the absorption of betaine. The addition of 0.5 mol/L NaCl resulted in a high osmotic pressure in the medium, which was not conducive to the growth of the recombinant bacteria and eventually led to a decrease in the expression of recombinant pullulanase. Diamant et al. also found that E. coli can reduce protein aggregation caused by high culture temperature by absorbing betaine added to the culture medium. Researchers believe that chemical molecular chaperones can improve the viscosity of cells, affect the dynamic interaction between molecular chaperones and folding substrates, promote the correct folding of proteins and reduce protein aggregates [23]. In this study, the effect of three osmotic pressure regulators on the solubilization of pullulanase was significantly different, which may be due to the difference in the size of the small molecule branched chain and the different types of protein aggregation, resulting in the different interactions between the protein surface and different small molecular substances.

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Effects of Betaine Concentration and Addition Time on Fermentation of the Recombinant Strain

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The effects of betaine concentration on the fermentation of the recombinant strain were investigated by adding 0, 10, 20, 40, and 80 mmol/L. As shown in Fig. 5.6a, the activities of pullulanase in the extracellular and intracellular supernatants of the control group (without betaine) were 10.6 and 21.2 U/mL, respectively. With increasing betaine concentration, the activity of soluble pullulanase first increased and then decreased. When the betaine concentration was 20 mmol/L, the total activity of soluble pullulanase was 45.3 U/mL. With increasing betaine concentration, the activity of soluble pullulanase began to decrease. The soluble pullulanase activity was only 22.1 U/mL when the betaine concentration was 80 mmol/L, which was 69.5% of the control group. To investigate the effect of betaine addition time on the fermentation of the recombinant strain, 20 mmol/L betaine was used as an osmotic pressure regulator to promote the extracellular secretion of pullulanase. The results showed that the total activity of soluble pullulanase was the highest when betaine was added at 0 h, which was approximately 48.9 U/mL, which was 163% of the control group (without betaine). The increase in pullulanase activity decreased with increasing addition time; when betaine was added at 6 h and 12 h, the soluble pullulanase activity was only 14.3% and 10.1% higher than that of the control group (Fig. 5.6b). In recent years, a large number of studies have confirmed that small molecular osmotic pressure regulators (such as proline, potassium glutamate, and betaine) can not only improve the thermal stability of proteins in vivo but also promote refolding of misfolded proteins in vitro [21, 23, 24]. Researchers believe that the existence of an osmotic pressure regulator can improve the viscosity of the protein dissolving

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system and effectively promote the folding intermediate into the natural folding pathway in the early stage of protein folding to reduce the occurrence of misfolding events. When the temperature of the protein expression system decreased significantly, increased intracellular viscosity also appeared.

5.2.5

Optimization of High-Density Fermentation Conditions of the Recombinant Strain

The results of the shake flask experiment (see Sects. 5.2.1–5.2.4) showed that the recombinant strain easily formed active protein aggregates during fermentation. The soluble expression level of pullulanase could be improved by using low fermentation temperature, low induction intensity, and adding surfactant as well as betaine. To achieve high-density fermentation of the recombinant pullulanase engineering strain, based on the optimization of fermentation conditions in shake flasks, the two-stage fermentation process previously established in the laboratory was adjusted appropriately, and a high-efficiency soluble expression fermentation process of recombinant pullulanase was established. In the first stage, the initial carbon source was exhausted, and dissolved oxygen began to rebound (the end of batch fermentation). The specific growth rate μset = 0.2 h-1 was controlled by the exponential flow addition method. This exponential feeding method can make the concentration of glycerol in the culture medium low, thus avoiding the accumulation of acetic acid caused by excessive glycerol and reducing the influence on the later induction stage of enzyme production. The second stage was the induction phase. When the cell concentration OD600 = 50, the glycerol feeding mode was changed from exponential feeding to constant speed feeding, and then the flow acceleration was gradually reduced according to the growth status of the bacteria; at the same time, lactose was added to induce the expression of recombinant protein (Fig. 5.7).

5.2.5.1

Effect of Fermentation Temperature on High-Density Fermentation of the Recombinant Strain

As shown in Sect. 5.2.2 of the shaking flask experiment, the total soluble enzyme activity of recombinant pullulanase induced at 37 °C was very low, which was only 52.8% and 33.7% of that at 30 and 25 °C, respectively, which proved that induction temperature was one of the key factors for soluble expression of recombinant pullulanase. In this study, a two-step temperature control culture method was used in the high-density production of recombinant pullulanase in a 3-L fermenter: cell growth stage and induction culture stage. To study the effect of temperature control on the production of recombinant pullulanase, the effects of three different temperature control methods on pullulanase production by the recombinant strain were investigated. The three different temperature control methods are as follows: mode A, first growing bacteria at 37 °C and then inducing culture at 30 °C;

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mode B, using 30 °C for cell growth and induction; mode C, first growing bacteria at 30 °C and then cooling to 25 °C for induction (subsequently, A, B, or C were used to represent the three temperature control methods). It can be seen from Fig. 5.8a that the growth rates of different temperature control methods were different. Temperature control mode A had the fastest growth rate, and

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the maximum dry cell weight was 80.3 g/L at 33 h; the maximum values of cell growth were achieved at 35 and 38 h in temperature control modes B and C, respectively, with dry cell weights of 78.1 and 77.5 g/L, respectively. The effect of different temperature control methods on the growth rate of bacteria was obvious, but the influence on the maximum bacterial volume was weak. However, the enzyme production decreased with increasing temperature (Fig. 5.8b): under the condition of temperature control mode A, the enzyme activities of intracellular and extracellular supernatants were 28.5 and 3.1 U/mL, respectively; under the condition of temperature control mode B, the enzyme activities of intracellular and extracellular supernatants were 107.2 and 6.3 U/mL, respectively; under the condition of temperature control mode C, the enzyme activities of intracellular and extracellular supernatants were 315.4 and 8.5 U/mL, respectively. The fermentation temperature is very important for the production of pullulanase by high-density fermentation of the recombinant strain, and a lower temperature control mode will be adopted in the following experiments.

5.2.5.2

Effect of Betaine on High-Density Fermentation of the Recombinant Strain

It can be seen from the results of high-density fermentation in Sect. 5.2.5.1 that the activity of pullulanase in the intracellular supernatant can reach 315.4 U/mL under temperature control mode C, which is 11.1 times the intracellular fermentation level under temperature control mode A. In addition, the early shake flask fermentation experiment also found that the addition of betaine to the culture medium on the basis of reducing temperature could further improve the enzyme production level. After adding betaine, the total activity was approximately 48.9 U/mL, which was 1.6 times that of the control group (see Sect. 5.2.4). However, whether the optimized conditions are effective in a 3-L fermenter still needs to be verified. In the high-density fermentation process, the temperature control and the addition of betaine were superimposed to investigate whether the addition of betaine in the fermenter can effectively improve the enzyme production level. When OD600 = 15 (early logarithmic growth), 20 mmol/L betaine was added to the medium at one time; when OD600 = 50 (medium logarithmic growth), exponential feeding was stopped, and 0.8 g/L/h lactose was added to induce the expression of pullulanase. The fermentation temperature was controlled at 30 °C in the growth stage and 30 and 25 °C in the induction phase. It can be seen from Fig. 5.9a that after adding betaine, the maximum dry cell weights of cells induced at 30 and 25 °C were 69.1 and 60.5 g/L, respectively, which were 88.5% and 78.1% of the dry cell weight of cells without betaine under the corresponding temperature control mode. Although the biomass of cells with betaine was lower than that without betaine, the enzyme production level was significantly improved. As shown in Fig. 5.9b, the maximum intracellular enzyme activity at 30 and 25 °C induction was 606.7.1 and 946.5 U/mL, respectively, which were 5.7 and 3.0 times higher than those without betaine at the corresponding temperature.

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The activity of extracellular enzymes was 11.0 and 17.4 U/mL after adding betaine, which was similar to the activity of the extracellular enzyme without betaine (4-9b). The above results demonstrate that adding betaine to the medium is the key to improving the soluble expression of pullulanase. Although the soluble expression of recombinant pullulanase in E. coli can be effectively improved by lowering the temperature and adding betaine, pullulanase mainly exists in the periplasmic space. When E. coli produces foreign proteins with signal peptides, it mainly secretes recombinant proteins into the periplasmic space through the cell membrane mediated by signal peptides; the process of protein “secretion” from the periplasmic space to the culture medium is nonspecific leakage, and the strength of this leakage determines the proportion of recombinant enzymes in the fermentation medium and periplasmic space. Previous studies in our laboratory found that glycine (0.75%) could effectively promote the extracellular leakage efficiency of recombinant proteins such as α-CGTase, α-amylase and sucrose isomerase in E. coli and finally secrete most of the recombinant enzymes into the culture medium. However, glycine had little effect on the extracellular leakage of recombinant pullulanase. Generally, the leakage of recombinant proteins in E. coli is affected by many factors, including culture composition, fermentation conditions, protein molecular weight, and protein surface charge distribution. The reason for the extremely low extracellular secretion efficiency of recombinant pullulanase in E. coli needs further study.

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Comparison of Fermentation Parameters of Recombinant Strain in Shake Flask and 3-L Fermenter

As unit cell productivity is an important parameter in the process of enzyme fermentation, it is necessary to calculate and compare the unit cell productivity in shake flasks and fermentation fermenters. As shown in Table 5.2, in the high-density fermentation process (without betaine), the unit cell productivity at 30 and 25 °C was 1.5 × 103 and 4.6 × 103 U/gcell, respectively. The unit cell productivity at 30 °C was only approximately 32.6% of that at 25 °C. The reason may be that the cell growth rate is low and the cell metabolism is slow at a low temperature (25 °C). At this time, the protein synthesis rate is also low, and the probability of misfolding of recombinant protein is reduced. Finally, the total amount of soluble pullulanase is increased, and the unit cell productivity is also high. At the same time, the unit cell productivity of the recombinant strain at 25 °C in a shake flask was 8.2 × 103 U/gcell, which was 1.8 times higher than that of high-density fermentation at the same temperature. The reason may be that the growth of the cell is limited and the metabolism of the cell is slow in the shake flask, while in the high-density fermentation, the conditions of nutrient transfer and dissolved oxygen in the culture medium are very good, the cell metabolism is vigorous and the growth is fast, and the speed of protein synthesis in the cell is higher than that in the shake flask. Due to the vigorous synthesis of intracellular protein and the shortage of natural molecular chaperones in the fermentation fermenter, the recombinant protein cannot be folded effectively, and the quality of the recombinant protein is reduced, making it prone to form aggregates. Therefore, the specific productivity of soluble protein is lower than that of the shake flask. When betaine was added to the fermenter, the specific productivity per cell in the 3-L fermenter reached 8.0 × 103 and 16.0 × 103 U/gcell at 30 and 25 °C, respectively, which were 5.3 and 3.5 times higher than that of the corresponding temperature without betaine. These results further indicated that betaine could improve the soluble expression of pullulanase.

Table 5.2 Comparison of fermentation parameters for the recombinant strain in shake flasks and 3-L fermenters. Reused with permission from Bioresource Technol. Duan X, Chen J, Wu J. 2013

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Soluble activity (U/mL) 23.0 ± 0.6 36.1 ± 0.8 49.3 ± 0.7 31.6 ± 1.3 115.8 ± 9.10 618.3 ± 40.6 321.8 ± 21.5 963.9 ± 69.4

Productivity (U/mL/h) 0.9 ± 0.1 1.3 ± 0.1 1.8 ± 0.1 1.0 ± 0.1 3.0 ± 0.2 16.3 ± 1.2 8.5 ± 0.6 25.4 ± 1.5

Productivity per cell (×103 U/ gcell) 6.1 ± 0.3 8.2 ± 0.3 10.2 ± 0.5 0.4 ± 0.02 1.5 ± 0.1 8.0 ± 0.6 4.6 ± 0.3 16.0 ± 1.1

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Effect of N-Terminal Domain Excision on the Thermostability and Secretory Efficiency of Pullulanase

Studies have shown that the molecular weight and structural complexity of proteins have a great influence on their soluble expression and secretion efficiency. Proteins with higher molecular weights and more complex structures more easily form inclusion bodies, and their extracellular secretion efficiency is lower [8]. In 2001, Enzyme Bio-Systems reported in a patent that the extracellular secretion level of recombinant B. naganoensis pullulanase could be increased by 1.6 times by removing 106 amino acids from the N-terminus [9]. The researchers believe that this may be due to the decrease in the molecular weight of the truncated protein. B. deramificans pullulanase is composed of 928 amino acids with a molecular weight of 101 kDa. The structural model analysis of B. deramificans pullulanase showed that it had six domains, CBM41, X25, X45, CBM48, A, and C. The CBM41 and CBM48 domains belong to the carbohydrate-binding domain 41 family and the 48 family, respectively, which have binding functions with starch substrates, while the functions of the X25 and X45 domains are unknown [25]. CBM48, A, and C domains are closely linked to form a whole CBM41, X25, and X45 domains are connected by a flexible linker, which has a strong swing. In Sect. 5.2, the soluble expression level of pullulanase could be significantly improved by optimizing the fermentation process and adding betaine. The enzyme activity of the intracellular supernatant reached 946.5 U/mL after 36 h of fermentation in a 3-L fermenter, while the extracellular enzyme activity was only 17.4 U/mL. Therefore, the recombinant pullulanase was mainly located in the periplasmic space, and the extracellular secretion efficiency was low. This phenomenon may be caused by the high molecular weight and complex structure of the recombinant pullulanase. In this study, based on the structural analysis of B. deramificans pullulanase, the N-terminal domain of B. deramificans was excised to different degrees, and the effects of domain excision on the secretion efficiency and enzymatic properties of pullulanase were investigated.

5.3.1

Construction, Recombinant Expression, and Purification of the N-Terminal Truncated Mutant of Pullulanase

5.3.1.1

Construction and Recombinant Expression of the N-Terminal Truncated Mutant

The structure analysis showed that B. deramificans pullulanase had six domains, among which the CBM48 domain, catalytic domain, and C domain were closely linked to form a whole with relatively complete structure and function. The structure

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of these three domains was similar to that of isoamylase, which may have a complete catalytic function as a starch debranching enzyme. In addition to the CBM41 domain, the other two X25 and X45 domains are not clearly classified. It is speculated that these three domains may be related to the binding of macromolecular polysaccharide substrates. Because pullulanase mainly deals with the branching bonds of dextrins with small molecular weights in practical applications, the binding capacity of macromolecular substrates is not high. If the N-terminal domain is removed, the debranching function of pullulanase on small molecular substrates may still be retained. In addition, the B. deramificans pullulanase protein has a high molecular weight and complex structure, easily forms active protein aggregates and has low secretion efficiency. The removal of the N-terminal domain can reduce the molecular weight and structural complexity of the protein, which may promote the secretion efficiency of pullulanase. Therefore, we constructed a series of N-terminal domain-deleted pullulanase mutants. Considering that there is a 14 amino acid linker between the CBM41 domain and X45 domain, CBM41 domain excision can be divided into two types: one is Puld1′, which removes only the CBM41 domain, and the other is Puld1, which removes the CBM41 domain and linker (as shown in Fig. 5.10). Using the plasmid pET24-ompA/pulA containing the wild-type pullulanase gene as a template, puld1′, puld1, puld2, and puld3 with different degrees of truncation in the N-terminal domain were amplified by PCR to construct four expression plasmids with truncated mutants: pET24-ompA/d1′, pET24-ompA/d1, pET24-ompA/d2, and pET24-ompA/d3. The plasmids were verified and transformed into E. coli BL21 (DE3) to obtain the recombinant strains. The recombinant strains were inoculated into shake flasks for culture and induced expression. After centrifugation, the fermentation supernatants were used for protein electrophoresis and enzyme activity determination. The results of SDS–PAGE electrophoresis (Fig. 5.11) showed that wild-type pullulanase and mutants Puld1′, Puld1, and Puld2 were soluble in E. coli, except for Puld3. Moreover, the protein bands of mutants Puld1′, Puld1, and Puld2 in the supernatant of fermentation broth were thicker than those of wild-type pullulanase. Protein electrophoresis showed that the amount of extracellular protein secreted by the truncated pullulanase-producing

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Fig. 5.11 SDS–PAGE analysis of extracellular fractions for wild-type enzyme and the truncated variants. Lane M protein markers; lane 1, wild-type enzyme; lane 2, Puld1′; lane 3, Puld1; lane 4, Puld2; lane 5, Puld3

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strain was significantly higher than that of the strain expressing wild-type pullulanase. The activity of recombinant wild-type pullulanase and the truncated mutant in the extracellular supernatant, intracellular supernatant, and intracellular precipitation showed that although the total enzyme activity of the mutant and wild-type enzyme did not change significantly, the ratio of extracellular enzyme activity to total enzyme activity of three mutants (Puld1′, Puld1, and Puld2) increased significantly. The extracellular enzyme activity of natural enzymes only accounted for 15.4% of the total enzyme activity. The proportion of extracellular enzymes of mutant Puld1′, Puld1, and Puld2 increased significantly, reaching 48.5%, 52.9%, and 58.7%, respectively, 3.1, 3.4, and 3.8 times that of the wild-type enzyme (Table 5.3). The ratio of enzyme activity to total enzyme activity in the intracellular supernatant and intracellular precipitation of the truncated mutant decreased, and the proportion of intracellular supernatant decreased most obviously. The extracellular secretion efficiency of Puld1′ and Puld1 was similar, and the extracellular secretion efficiency of Puld2 with a smaller molecular weight was higher than that of Puld1′ and Puld1. The above results showed that the secretion efficiency of N-terminal truncated pullulanase was significantly improved, which indicated that the extracellular secretion efficiency of the protein was indeed related to the molecular weight. The

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increase in protein secretion efficiency may be due to the decrease in the molecular weight of pullulanase after truncation, which is more conducive to leakage from the periplasmic space to the extracellular medium. In addition, no pullulanase activity was detected in the supernatant, intracellular supernatant, and intracellular precipitate of Puld3. This indicates that the existence of the X45 domain may be important for promoting the correct folding of pullulanase and maintaining its conformation. Its removal may lead to the failure of pullulanase to fold into a space stereoscopic structure with activity. The results of protein electrophoresis showed that Puld3 mainly existed in the form of a large number of inactive inclusion bodies in the cell precipitate, so the following experiments did not involve Puld3.

5.3.1.2

Isolation and Purification of the N-Terminal Truncated Mutant

To further study the enzymatic properties of recombinant pullulanase and its mutants, recombinant pullulanase was isolated and purified. The supernatant of the recombinant protein was treated by heat treatment, 50% (w/v) ammonium sulfate precipitation, dialysis, and DEAE-Sepharose anion exchange chromatography. It can be seen from Fig. 5.12 that the purified recombinant pullulanases purd1’, puld1, and puld2 are electrophoretically pure. As shown in Table 5.4, the specific activity of the recombinant pullulanase mutant puld1’ was increased from 84.7 to 357.0 U/mg, with a purification multiple of 4.2 times; the specific activity of the mutant puld1 was increased from 89.2 to 372.9 U/mg with a purification multiple of 4.1 times; the specific activity of the mutant puld1 was increased from 62.50 to 250.1 U/mg, with a purification multiple of 4.0. The specific activities of the mutants puld1′, puld1, and puld2 were 88.1%, 92.1%, and 61.8% of the natural enzyme, respectively. The specific activities of the three mutants (pullulan polysaccharide as substrate)

Fig. 5.12 SDS–PAGE analysis of purified wildtype enzyme and the truncated variants. Lane M protein markers, lane 1, Puld1′; lane 2, Puld1; lane 3, Puld2; lane 4, wild-type enzyme

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Table 5.4 Purification scheme of truncated pullulanase variants Purification steps Crude enzyme Heat treatment Ammonium sulfate precipitation Anion exchange chromatography

Specific activity (U/mg) Puld1′ Puld1 Puld2 84.7 89.2 62.5 177.4 180.3 134.7 239.1 241.9 169.7 357.0 372.9 250.1

Purification fold Puld1′ Puld1 1.0 1.0 2.1 2.0 2.8 2.7 4.2 4.1

Puld2 1.0 2.2 2.7 4.0

decreased, among which the specific activities of puld1’ and puld1 were similar, approximately 90% of the natural enzyme; the specific activity of puld2 decreased the most, approximately 40%. This phenomenon may be related to the function of the N-terminal domain. The CBM41 domain and X25 domain may have binding ability with pullulan polysaccharide. Their removal will affect the catalytic activity of pullulanase. Among them, puld2 has the most removed domain, and the enzyme activity is most affected.

5.3.2

Construction, Expression, and Purification of Superimposed Mutants

5.3.2.1

Construction of Superimposed Mutants

Although the mutants puld1’, puld1, and puld2 obtained in the previous section have higher extracellular secretion efficiency, their specific activities are lower than those of natural enzymes, which may be due to the reduction in substrate binding capacity due to the excision of the N-terminal domain. The mutant D437H/D503Y obtained in a previous study has better substrate binding ability and catalytic efficiency than natural enzymes. Therefore, we considered combining point mutation with N-terminal truncated mutation to construct a new mutant. Since there is no obvious difference between puld1′ and puld1, the subsequent experiments only performed superposition mutation on D437H/D503Y and puld1 and puld2. The overlapping mutants D437H/D503Y/D1 and D437H/D503Y/D2 were recombined and expressed in E. coli. The SDS–PAGE results of the fermentation supernatant of the recombinant strain are shown in Fig. 5.13. The protein size was consistent with the theoretical value, and the protein bands of D437H/D503Y/d1 and D437H/D503Y/d2 were significantly thicker than those of the control D437H/ D503Y/D2. In addition, the activity of the recombinant enzyme showed that the proportion of extracellular supernatant of mutant D437H/D503Y/D1 and D437H/D503Y/D2 was 57.3% and 60.8%, respectively, which was 5.3 times and 5.6 times that of control D437H/D503Y (Table 5.5). The results showed that the soluble enzyme activity (extracellular and intracellular supernatants) of the two mutants accounted for approximately 80% of the total enzyme activity, and only approximately 20% of

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Fig. 5.13 SDS–PAGE analysis of extracellular fractions for D437H/D503Y and the truncated variants. Lane M, protein markers; lane 1, D437H/D503Y; lane 2, D437H/D503Y/D1; lane 3, D437H/D503Y/D2

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Table 5.5 Location of pullulanase activity of D437H/D503Y and its truncated variants Pullulanase D437H/D503Y D437H/D503Y/ D1 D437H/D503Y/ D2

Extracellular ratio (%) 10.9 57.3

Intracellular ratio (%) 31.5 21.4

Intracellular precipitation ratio (%) 57.6 21.3

60.8

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18.7

the enzyme activity existed in the form of active protein aggregates. The soluble enzyme and active protein aggregates of D437H/D503Y accounted for 42.4% and 57.6% of the total enzyme activity, respectively. It can be concluded that the superposed mutants have better extracellular secretion and expression abilities than the control and may have better production potential.

5.3.2.2

Isolation and Purification of Superimposed Mutants

It can be seen from Fig. 5.14 that the purified pullulanase superimposed mutants D437H/D503Y/D1 and D437H/D503Y/D2 reached electrophoretic purity after purification. As shown in Table 5.6, the specific activity of the recombinant pullulanase mutant D437H/D503Y/D1 increased from 81.5 to 415.7 U/mg, with a purification multiple of 5.1 times; the specific activity of the mutant D437H/D503Y/ D2 increased from 129.4 to 563.5 U/mg, with a purification multiple of 4.4 times. The specific activities of D437H/D503Y/D1 and D437H/D503Y/D2 were 1.04 and 1.41 times higher than those of D437H/D503Y and 1.03 and 1.39 times that of the natural enzyme, respectively.

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Fig. 5.14 SDS–PAGE analysis of purified D437H/ D503Y and the truncated variants. Lane M, protein markers; lane 1, D437H/ D503Y; lane 2, D437H/ D503Y/d2; lane 3, D437H/ D503Y/d1. Reused with permission from Appl Environ Microbiol. Duan X, Wu J. 2015

Table 5.6 Summary of purification steps of the truncated variants of D437H/D503Y

Purification steps Crude enzyme Heat treatment Ammonium sulfate precipitation Anion exchange chromatography

Specific activity (U/mg) D437H/ D437H/ D503Y/d1 D503Y/d2 81.5 129.4 228.2 332.5 293.4 431.7

Purification fold D437H/ D503Y/d1 1 2.8 3.6

D437H/ D503Y/d2 1.0 2.6 3.3

415.7

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4.4

563.5

5.3.3

Analysis of Enzymatic Properties of Mutants

5.3.3.1

Determination of Optimum pH for Mutants

The activities of N-terminal truncated and superimposed mutants of pullulanase were measured in an acetic acid buffer system (pH 3.5–5.0) and a phosphate buffer system (pH 5.0–6.0) at 60 °C. The enzyme activity measured at the optimum pH was set as 100%, and the activity at other pH values was calculated as the percentage of enzyme activity at the optimum pH. As shown in Fig. 5.15, the optimal pH of all N-terminal truncated and superimposed mutants is 4.5, which is the same as that of natural enzyme and D437H/D503Y, but the pH range of different mutants varies. The relative activity of truncated pullulanase was only approximately 50% above the optimum pH, which was significantly lower than that of the natural enzyme by approximately 80%. Under the optimum pH, the relative activities of puld1 and puld2 were 60% and 31%, respectively. It can be seen that the pH range of puld2 is the narrowest, and the pH range of puld1 is shifted to the direction of low pH. The

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Relative enzyme activity (%)

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pH Fig. 5.15 Optimal pH of truncated pullulanase variants. Reused with permission from Appl Environ Microbiol. Duan X, Chen J, Wu J. 2013

increased pH range of the superimposed mutants may be due to the wider pH range of D437H/D503Y.

5.3.3.2

Optimum Temperature and Temperature Stability

As shown in Fig. 5.16a, the optimum temperature of puld1 and puld2 is 55 °C, which is the same as that of natural enzymes. The activities of puld1 and puld2 were 97.0% and 89.3%, respectively, at 60 °C, which were higher than those of natural enzymes at 60 °C. The optimum temperature of D437H/D503Y/D1 and D437H/D503Y/D2 was 60 °C, which was 5 °C higher than that of the natural enzyme and the same as that of the double mutant D437H/D503Y. The activity of puld1 and D437H/D503Y/ D1 was approximately 85% at 70 °C, which was the same as that of the double mutant D437H/D503Y, while the activity of puld2 and D437H/D503Y/D2 was only 39% and 51% at 70 °C, respectively. The thermal stability of the truncated pullulanase mutant was measured at pH 4.5 and 60 °C. As shown in Fig. 5.16b, the half-lives of puld1 and puld2 were 57 h and 34 h, respectively, which were 2.6 and 1.5 times those of the natural enzyme, respectively. The half-lives of the superimposed mutants D437H/D503Y/D1 and D437H/D503Y/D2 were 203 h and 160 h, respectively, which were 9.0 and 7.1 times that of the natural enzyme and 1.7 and 1.3 times that of the double mutant D437H/D503Y (120 h), respectively. It can be seen that the stability of N-terminal

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Fig. 5.16 Optimal temperature (a) and thermostability (b) of truncated pullulanase variants. Reused with permission from Appl Environ Microbiol. Duan X, Chen J, Wu J. 2013

truncated mutants of pullulanase has been improved to varying degrees; when the truncated mutation and the point mutation are superimposed, the two mutations overlap each other, and the thermal stability of the superimposed mutant is further improved. According to the three-dimensional crystal structure of B. acidopullulyticus pullulanase and B. deramificans pullulanase simulation structure analysis, we can see that the N-terminal domain of pullulanase is flexible, especially the terminal CBM41 domain, which may have a negative impact on the overall thermal stability of pullulanase. Many previous studies have shown that protein thermal stability can be effectively improved by deleting or replacing regions with poor thermal stability. In this study, the N-terminal domain was truncated to improve the thermal stability of pullulanase, which confirmed the effectiveness of this method. Whether domain excision affects the catalytic properties of pullulanase still needs to be verified by subsequent experiments.

5.3.3.3

Determination of the Kinetic Parameters of Mutants

The kinetic parameters of the natural enzyme, double mutant D437H/D503Y and N-terminal truncated enzyme (puld1, puld2, D437H/D503Y/D1, and D437H/ D503Y/D2) are shown in Table 5.7. Compared with the natural enzyme, the N-terminal truncated mutation reduced the binding ability of the mutants puld1 and puld2 to pullulan substrate. The km values of pullulanase were 1.6 times and 4.4 times higher than those of natural enzymes, respectively. Among them, the N-terminal domain of pullulanase with the most truncated N-terminus was also the worst, which indicated that the N-terminal domain of pullulanase was involved in the binding of polysaccharide macromolecular substrates and contributed to the substrate binding ability. The Vmax values of the mutants puld1 and puld2 were 1.1 and

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Table 5.7 Kinetic parameters of truncated pullulanase variants. Reused with permission from Appl Environ Microbiol. Duan X, Chen J, Wu J. 2013 Enzyme Native enzyme Puld1 Puld2 D437H/D503Y Puld1 + D437H/D503Y Puld2 + D437H/D503Y

Km (mg/mL) 0.70 ± 0.04 1.12 ± 0.06 3.11 ± 0.16 0.38 ± 0.02 0.56 ± 0.04 1.05 ± 0.06

Vmax (U/mg) 469.7 ± 23.4 517.1 ± 25.8 550.5 ± 27.1 436.9 ± 21.6 415.5 ± 20.7 576.8 ± 9.10

kcat (s-1) 1900.4 ± 95.0 2081.8 ± 104.0 1609.7 ± 80.5 2031.0 ± 97.5 2685.9 ± 124.2 2851.5 ± 87.3

kcat/Km (mL/mg
s1 ) 2712.9 ± 135.6 1867.1 ± 93.3 517.4 ± 25.8 5403.0 ± 270.1 4810.8 ± 240.5 2710.5 ± 91.0

1.2 times higher than those of the native enzyme, respectively. This may be because the truncation of the N-terminal domain reduces the steric hindrance of pullulanase, resulting in an increase in the maximum reaction speed. In addition, the truncation of the N-terminal domain obviously has some adverse effects, especially on the catalytic efficiency. It can be seen from Table 5.7 that the kcat/Km values of puld1 and puld2 are only 68.8% and 19.1% of those of natural enzymes. This is mainly due to the decreased binding ability of N-terminal truncated mutants to substrates or the decrease in both substrate binding capacity and kcat coefficient. Although the stability of the truncated mutants was improved, the kinetic parameters of the enzyme decreased. Therefore, reducing the negative effects of N-terminal truncation on enzyme kinetic parameters is particularly important. In a previous study, site-directed mutagenesis of pullulanase was carried out. The substrate binding ability of the double mutant D437H/D503Y was significantly improved, its Km value was only 54.3% of that of the natural enzyme, and the kcat value of the double mutation was also higher than that of the natural enzyme. In this study, the N-terminal truncation and double mutation were superimposed, and the kinetic parameters of the superimposed mutants were investigated. The results showed that the substrate binding abilities of D437H/D503Y/D1 and D437H/ D503Y/D2 were significantly improved compared with those of the corresponding nonsuperimposed mutants puld1 and puld2, and their Km values were 50% and 34% of those of puld1 and puld2, respectively. The kcat values of the superimposed mutants D437H/D503Y/D1 and D437H/D503Y/D2 were 1.4 and 1.5 times higher than those of natural enzymes, respectively. Their catalytic efficiency (kcat/Km) was 1.8 and 1.0 times higher than that of natural enzymes, respectively. These results indicated that the substrate binding ability and catalytic ability of the superimposed mutants were significantly improved, which compensated for the adverse effects caused by simple N-terminal truncation. The superimposed mutants not only have better stability but also greatly improve the catalytic kinetic parameters, which may have better application value.

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Substrate Specificity Analysis of Mutants

Using pullulan polysaccharide, dextrin (DE 10), soluble starch, amylopectin, and glycogen at a 1% (w/v) concentration as substrates, the relative enzyme activities of the natural enzyme, site-specific mutant, and truncated mutant were measured at pH 4.5 and 60 °C. It can be seen from Table 5.8 that pullulan polysaccharide is the best substrate for all pullulanases, followed by dextrin with a DE value of 10. All enzymes have no glycogen hydrolysis activity. Compared with the natural enzyme, the pullulanase double mutant had a slight decrease in the relative activity of soluble starch, but the relative activity of dextrin and amylopectin was significantly increased, especially for amylopectin. It may be that the double mutant pullulan enhances the affinity of branched starch substrates with more complex branches, which is more conducive to the hydrolysis of large branched substrates. However, the double mutant pullulan has no hydrolytic activity on glycogen, probably because glycogen branches of glycogen are denser, pullulan has difficulty reaching the branching point, and all cannot attack the branching bond. The relative activities of truncated pullulanase puld1 and puld2 to dextrin, soluble starch, and amylopectin were all decreased. Among them, the hydrolytic activity of the above substrates was decreased most by puld2, which had the most removed domains. As shown in Table 5.8, the relative activity of puld2 to dextrin, soluble starch, and amylopectin is 60%, 32%, and 0% of that of a natural enzyme, respectively, indicating that the hydrolysis ability of puld2 to substrate with higher molecular weight and more dense branches is poorer, which is determined by the function of CBM41 and X25 domains. The addition of double mutation and truncated mutation can partly compensate for the decrease in the relative activity of dextrin, soluble starch, and amylopectin caused by N-terminal truncation. Table 5.8 shows that D437H/D503Y/D1 and D437H/D503Y/D2 have different degrees of improvement in the relative activities of dextrin, soluble starch, and amylopectin. The superposition of D1 and D437H/ D503Y increased the relative activity of amylopectin from 9.6% to 37.3%. The double mutation can compensate for the substrate specificity change caused by the truncation of the CBM41 domain. After superposition of Puld2 and D437H/D503Y, although the relative activity of amylopectin was increased, the extent of improvement was limited. The enhancement of the substrate binding capacity of

Table 5.8 Substrate specificity of truncated pullulanase variants Substrates (1% (W/V)) Pullulan Dextrin (DE 10) Soluble starch Amylopectin a

Relative activity (%) Natural Puld1 enzyme 100 100 64.2 59.4 62.5 37.9 16.4 9.6

ND, no enzyme activity was detected

Puld2 100 38.5 20.2 0NDa

D437H/ D503Y/d1 100 61.4 48.1 37.3

D437H/ D503Y/d2 100 41.4 32.7 0.3

D437H/ D503Y 100 82.8 58.3 64.1

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S. aeruginosa can only partially compensate for the decrease in substrate binding capacity caused by domain truncation.

5.3.4

Application of Mutants in Starch Saccharification

Thirty percent (dry basis) corn starch was used as the substrate, which was liquefied by α-amylase for 1.5 h, cooled to 60 °C and adjusted to pH 4.5. Then, 100 U/g starch (dry basis) glucoamylase and 0.5 U/g starch (dry basis) pullulanase (puld1, puld2, D437H/D503Y/D1, and D437H/D503Y/D2) were added, and the reaction lasted for 62 h. As shown in Fig. 5.17, within 25 h of the initial reaction, the DX values of all reaction groups increased rapidly to 82–90%. Among them, the DX value with puld1 increased fastest, followed by D437H/D503Y/D1, then D437H/D503Y/D2, and puld2. The DX values of all samples reached a maximum after 50 h of reaction. The maximum DX values of puld1, puld2, D437H/D503Y/D1, and D437H/D503Y/ D2 were 93%, 92.5%, 95%, and 94%, respectively. Combined with the saccharification data in the previous section and the control experiment (the results are not shown), the maximum DX value of puld1 and puld2 is equivalent to that of the natural enzyme (92.9%); the maximum DX value of D437H/D503Y/D1 (95.0%) is 0.5% lower than that of D437H/D503Y (approximately 95.5%); the maximum DX value of D437H/D503Y/D2 is 94.1%, which is better than that of the natural enzyme but is 1.4% lower than that of double mutation DX.

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Time (h) Fig. 5.17 Glucose production by a combination of glucoamylase and truncated pullulanase variants. Reused with permission from Appl Environ Microbiol. Duan X, Chen J, Wu J. 2013

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Surfactant Promotes the Soluble Secretory Expression of Pullulanase in E. coli

In previous research by our laboratory, pullulanase from B. deramificans was expressed in E. coli, and a pullulanase mutant with a high application value was obtained by site-directed mutagenesis and domain excision. However, further studies showed that the enzyme easily formed a large number of active protein aggregates in E. coli, and the efficiency of extracellular secretion was poor. According to the literature reports, surfactants can promote the partial unfolding of inclusion bodies or protein aggregates, and partially unfolded proteins can be refolded to form soluble active proteins [19, 26, 27]. In addition, surfactants also dissolve cell membrane lipids and increase cell permeability, which are commonly used to promote extracellular protein secretion of E. coli. In this study, we investigated the effect of surfactants on the activity aggregate of pullulanase and its promoting effect on the soluble secretory expression of pullulanase in E. coli.

5.4.1

Depolymerization of Pullulanase Active Aggregates by Surfactants

Previous studies in the laboratory found that the expression of pullulanase from B. deramificans in E. coli would form a large number of inclusion bodies, and these inclusion bodies were active aggregates with pullulanase activity. The recombinant E. coli was cultured, the supernatant and the bacteria were separated by centrifugation, and then the cells were broken by ultrasonication. The enzyme activity of the samples was determined, and the results are shown in Fig. 5.18. The results showed that the precipitate (possibly inclusion body) collected after cell wall breakage was suspended in 20 mmol/L pH 4.5 acetate buffer, and the enzyme activity was measured. The activity was 80.5 U/mL, which was 1.3 times the soluble activity of the recombinant enzyme (fermentation liquid and wall-breaking supernatant). After centrifugation, the enzyme activity of the supernatant was only 2.7 U/mL. This result suggested that the inclusion body that is not soluble in water formed by pullulanase in E. coli is not a real inclusion body but a group of active proteins with catalytic activity. According to the data reported [19, 26], some surfactants (such as Tween-80 and Triton X-100) can promote partial folding of inclusion bodies or protein aggregates, and these partially folded proteins can be refolded to become soluble active proteins. In this study, the abovementioned breaking cells were suspended in 0.1% (w/v) Tween-80 or Triton X-100 buffer and then centrifuged to measure enzyme activity after incubation at room temperature for 10 min. The activity of supernatant enzyme containing Tween-80 was 75.6 U/mL and that of Triton X-100 was 73.8 U/mL, 93.9%, and 91.7% of pullulanase activity of the active aggregates, respectively. In addition, after treatment with Tween-80 or Triton X-100, the solid was significantly

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Fig. 5.18 Activity of pullulanase in different components of the E. coli recombinant strain. a, Extracellular pullulanase activity; b, Intracellular pullulanase activity; c, The activity of the pullulanase was measured with 20 mM acetic acid buffer solution at pH 4.5 for wall breaking precipitation; d, After the wall breaking precipitate was suspended in 20 mM acetic acid buffer at pH 4.5, the supernatant was centrifuged to measure the pullulanase activity; e, After the wall breaking precipitate was suspended in acetic acid buffer containing 0.1% (w/v) Tween-80, the supernatant was centrifuged to measure the pullulanase activity; f, After the wall breaking precipitate was suspended in acetic acid buffer containing 0.1% (w/v) Triton X-100, the supernatant was centrifuged to measure the pullulanase activity. Reused with permission from Bioresource Technol. Duan X, Zou C, Wu J. 2015

reduced after centrifugation. This indicates that Tween-80 and Triton X-100 depolymerize and refold the active aggregates of pullulanase into soluble protein.

5.4.2

The Influence of Surfactant Species on the Growth and Enzyme Production of E. coli

From a previous study, we can see that some surfactants depolymerize the active aggregates of pullulanase into soluble proteins. However, the process requires breaking cells to obtain the activity aggregates of pullulanase, and then soluble pullulanase can be obtained by surfactant treatment. The process is complicated and difficult to use in industrial amplification. According to the data reported [19, 26], surfactants cannot only promote inclusion bodies or protein aggregation to become soluble active proteins but also dissolve lipids and increase the permeability of the cell membrane and are often used to promote the extracellular expression of proteins. Therefore, adding surfactant to the culture of recombinant E. coli may not only

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Fig. 5.19 Effects of surfactant type on DCW (a) and pullulanase production (b). DCW (black), extracellular pullulanase activity (white), intracellular pullulanase activity (light gray). Reused with permission from Bioresource Technol. Duan X, Zou C, Wu J. 2015

reduce the formation of inclusion bodies but also promote the secretion of pullulanase. To select the appropriate surfactant, 2 g/L Tween-80, Triton X-100, Span-80, S-185, Sarkoyl, SDS, and NP40 were added to the recombinant E. coli culture medium, and the results are shown in Fig. 5.19. The total enzyme activity was 1.54 and 1.53 times higher than that of the control after adding Sarkoyl or SDS in the medium, and the extracellular secretion efficiency of pullulanase was also increased to 98.4% and 96.8%, respectively, which were 1.94 and 1.90 times that of the control. However, the dry cell weight of E. coli decreased sharply to 1.8 and 2.0 g/ L, which was only 25.8% and 29.5% of the control. In addition, the viscosity of the bacteria was higher in the medium with the two surfactants added. This indicates that E. coli may undergo cell cleavage under the action of these two surfactants, which makes the mass of the matter in the cytoplasm leak out of the cell. The results showed that the cleavage of E. coli was still obvious when the concentration of sarkoyl and SDS was reduced to 1 g/L in the medium, but the extracellular secretion efficiency of pullulanase decreased to 83.8% and 79.5%, respectively. Adding Tween-80, Triton X-100, Span-80, S-185, or NP40 to the medium not only significantly improved the total activity of soluble protein and extracellular secretion efficiency of pullulanase but also did not significantly increase cell cleavage. Among them, Triton X-100 increased the total enzyme activity and extracellular secretion efficiency of pullulanase to the most significant degree, which was 115.0 U/mL and 85.9%, respectively, which were 45% and 69.1% higher than that of the control group. Therefore, Triton X-100 is the most favorable surfactant for recombinant E. coli to express pullulanase. Based on the above research, surfactants have two functions in the process of expressing pullulanase in E. coli: on the one hand, they can pass through the outer membrane of E. coli and effectively depolymerize the pullulanase aggregates, thus increasing the expression of pullulanase soluble protein; on the other hand, they can

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also improve the permeability of the cell membrane, significantly improving the extracellular secretion efficiency of pullulanase.

5.4.3

Effect of Triton X-100 Concentration on Growth and Enzyme Production of E. coli

According to the previous Sect. 5.4.2, Triton X-100 can significantly improve the extracellular secretion efficiency of pullulanase in E. coli. The effect of Triton X-100 concentration on the growth and enzyme production of recombinant E. coli was investigated in this section because the promotion effect of Triton X-100 on the extracellular expression of pullulanase might be small at a low concentration, and cell growth might be seriously inhibited if the concentration of Triton X-100 is too high. The recombinant E. coli was cultured in medium supplemented with 1, 2, 5, 10, or 20 g/L Triton X-100. The growth and enzyme production of the recombinant E. coli are shown in Fig. 5.20. The results showed that with increasing Triton X-100 concentration, the growth of the recombinant strain became increasingly worse. Especially when the concentration of Triton X-100 was more than 5 g/L, the cell concentration significantly decreased. In the range of 0–5 g/L Triton X-100, the total activity and extracellular secretion efficiency of pullulanase increased significantly with increasing Triton X-100 concentration. When the concentration of Triton X-100 was 5 g/L, the total activity and extracellular secretion efficiency of pullulanase were 125.1 U/mL and 96.8%, respectively, which were 57.6% and 90.6% higher than those of the control (without Triton X-100). When the Triton X-100 concentration exceeded 5 g/L, the total activity and extracellular secretion

Fig. 5.20 Effects of Triton X-100 concentration on DCW (a) and pullulanase production (b). DCW (black), extracellular pullulanase activity (white), intracellular pullulanase activity (light gray). Reused with permission from Bioresource Technol. Duan X, Zou C, Wu J. 2015

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efficiency of pullulanase changed little. In summary, the optimal concentration of Triton X-100 was 5 g/L.

5.4.4

Effect of Triton X-100 Addition Time on the Growth and Enzyme Production of E. coli

According to previous studies, the addition of surfactants (such as Triton X-100) to the culture medium can inhibit the growth of E. coli. According to the data reported [19], delayed addition of surfactant can reduce its inhibition on the growth of bacteria to a certain extent. Therefore, the effects of adding Triton X-100 and not adding Triton X-100 (control) at 24, 36, and 48 h on the growth and enzyme production of E. coli were further studied. The results are shown in Fig. 5.21. According to Fig. 5.21a, adding Triton X-100 at 24 h could inhibit the growth of bacteria, and the dry cell weight of bacteria was 78.7% of the control. However, the addition of Triton X-100 at 36 or 48 h had little effect on the growth of bacteria. When Triton X-100 was added at 24, 36 or 48 h, the final extracellular pullulanase activity was 181.9, 192.0 and 216.3 U/mL, respectively, which were 3.7, 4.0 and 4.6 times higher than that of the control. Therefore, the addition of Triton X-100 at the late stage of fermentation not only did not affect cell growth but also significantly increased the extracellular activity of pullulanase. This result is similar to that reported by Ding Runrong et al. on promoting extracellular expression of α-CGTase in E. coli by delaying the addition of surfactants [19]. Considering the promotion effect of Triton X-100 on extracellular secretion of recombinant protein and the E. coli fermentation cycle, Triton X-100 was added at 48 h of fermentation.

Fig. 5.21 Effect of Triton X-100 addition time on growth (a) and enzyme production (b). ■, control; ●, 24 h; ▲, 36 h; ◆, 48 h. Reused with permission from Bioresource Technol. Duan X, Zou C, Wu J. 2015

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Effect of Triton X-100 on High-Density Fermentation of E. coli in a 3-L Fermenter

In the preliminary study of our laboratory, a high-efficiency expression technology of recombinant pullulanase was established based on the high-density fermentation process of an E. coli 3-L fermenter. In this process, the pH value is approximately 7.0 by adding ammonia water, and the DO is approximately 30% by controlling the stirring speed and ventilation rate and adding a certain proportion of pure oxygen. When the initial carbon source (glycerol) was completely consumed (marked by the sudden increase in DO), glycerol was fed exponentially. The specific growth rate of E. coli was controlled to 0.2 h-1, and the temperature was controlled at 30 °C. When the dry cell weight was DCW = 30 g/L, the temperature was reduced to 25 °C, and lactose was fed at a certain speed to induce the expression of recombinant protein; at this stage, glycerol was fed according to the growth of the cell by first decreasing the speed and then a constant speed. This process can control the concentration of glycerol at a very low level and avoid the accumulation of byproducts such as acetic acid. In addition, the soluble expression of pullulanase in E. coli was effectively promoted by low-temperature induction and betaine addition. The total enzyme activity reached 956.5 U/mL through high-density fermentation in a 3-L fermenter, but only a small amount of pullulanase was secreted into the fermentation broth [28]. Based on previous studies, we expect to improve the soluble expression and extracellular secretion efficiency of pullulanase by adding Triton X-100 in the later stage of fermentation. However, because Triton X-100 is a nonionic surfactant, a large number of bubbles will be produced under aeration and high-speed stirring. Therefore, we chose to stop ventilation in the late fermentation stage and reduce the rotational speed (100 rpm) before adding Triton X-100. According to Fig. 5.22, the dry cell weight of cells, the extracellular activity and secretion efficiency of pullulanase were 84.1 g/L, 17.6 U/mL, and 1.9%, respectively, at 40 h of fermentation. In the absence of Triton X-100, the intracellular and extracellular pullulanase activity almost did not increase after 40 h. Therefore, 5 g/L Triton X-100 was added at 40 h. After treatment with Triton X-100 for 6 h, the dry cell weight and intracellular enzyme activity decreased to 80.5 g/L and 128.1 U/mL, respectively, while the extracellular pullulanase activity increased to 812.4 U/mL, and the extracellular secretion efficiency of pullulanase was 86.4%, which was 46.2 and 45.5 times higher than that without Triton X-100. Although Triton X-100 significantly increased the extracellular activity and secretion efficiency of pullulanase, it did not promote the total amount of soluble pullulanase, which was inconsistent with the results of shake flask fermentation. According to the literature [29], this may be due to the differences in the properties of inclusion bodies (or aggregates) formed under different fermentation conditions. The production intensity of pullulanase was 23.8 U/mL/h in a 3-L fermenter, which was 14.4 times that in a shake flask. Due to the high molecular weight and complex structure of pullulanase, some inactive proteins may be formed in the process of rapid synthesis due to the lack of time for folding, and these proteins may aggregate to

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Fig. 5.22 Expression of pullulanase in E. coli as a function of Triton X-100. ■, DCW; ●, intracellular pull-down activity; ▲, extracellular pull-down activity. Reused with permission from Bioresource Technol. Duan X, Zou C, Wu J. 2015

form some inactive inclusion bodies. They have no biological activity even if they are treated with surfactants. This may be the main reason why inclusion bodies formed on the 3-L fermenter are different from those in shake flasks. In addition, Triton X-100 may not be suitable for large-scale fermentation due to its severe foaming during aeration agitation.

5.5

Optimization of the Induction Method Combined with the Glycine Feeding Strategy to Promote the Extracellular Expression of Pullulanase in E. coli

Reducing the formation of inclusion bodies is an important way to increase the production of recombinant protein. Many researchers have reported strategies to reduce the formation of inclusion bodies in E. coli [30], mainly including reducing the speed of recombinant protein synthesis by low-temperature culture, reducing the amount of inducer or using low-intensity promoter, assisting the recombinant protein to fold correctly by coexpressing molecular chaperone, adding osmotic pressure regulator to reduce the folding speed and avoid misfolding, and secreting the recombinant protein outside the cell to avoid the aggregation of recombinant protein in the narrow space in the cell. In the production of enzyme preparation, if the production bacteria can secrete the target protein directly into the fermentation broth, it can not only effectively improve

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soluble expression but also simplify the extraction process and reduce the cost. According to the literature report [31], the strategies to improve the extracellular secretion of E. coli mainly include changing the transport pathway of recombinant protein, fusing the transporter onto the target protein to assist the transport, coexpressing proteins that can destroy the cell membrane, and adding some reagents to improve the permeability of the cell membrane. Adding glycine to the medium can effectively improve the cell membrane permeability, which is a common method to enhance the extracellular secretion of E. coli. In our previous Sect. 5.2, the soluble expression of pullulanase was effectively improved by reducing the induction temperature and adding an osmotic pressure regulator. The total enzyme activity was increased to 956.5 U/mL by high-density fermentation in a 3-L fermenter, but only a small amount of pullulanase was secreted into the fermentation broth. In the previous Sect. 5.4, although the soluble secretory expression level of pullulanase was effectively improved by adding surfactants, it may not be suitable for large-scale production due to its serious foaming in aeration agitation. To solve the above problems, we will optimize the induction method and add glycine to improve the pullulanase production level.

5.5.1

Effect of Inducer Type and Concentration on Growth and Enzyme Production of E. coli

In this study, the T7lac promoter was used as the promoter of pullulanase expression in E. coli. The promoter usually uses IPTG or lactose as an inducer to initiate the expression of recombinant protein. IPTG is a highly effective inducer that is very stable and difficult for bacteria to metabolize and decompose. Lactose is a disaccharide that can be decomposed and utilized by E. coli. It can be used as an inducer and a carbon source. In this section, the effects of different concentrations of IPTG and lactose on the growth and enzyme production of E. coli were compared. The results are shown in Figs. 5.23 and 5.24. Compared with the control without any inducer, the expression of pullulanase was significantly increased by IPTG and lactose. It can be seen from Fig. 5.23 that the addition of IPTG has a certain inhibitory effect on the growth of bacteria, and the higher the concentration, the more obvious the inhibition. The optimal concentration of IPTG was 20 μmol/L, and the total pullulanase activity was 2.2 times higher than that of the control (without inducer). It can be seen from Fig. 5.24 that lactose not only slightly promotes the growth of bacteria but also significantly promotes the expression of pullulanase. Under the optimal concentration of lactose (5 g/L), the activity of pullulanase increased 3.9 times compared with the control. In addition, IPTG has potential toxicity to the human body and is very stable, which limits its

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Fig. 5.23 Effect of IPTG concentration on growth (a) and enzyme production (b). DCW (black), extracellular pullulanase activity (white), intracellular pullulanase activity (light gray). Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

Fig. 5.24 Effects of lactose concentration on DCW (a) and pullulanase production (b) in E. coli. DCW (black), extracellular pullulanase activity (white), intracellular pullulanase activity (light gray). Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

large-scale use in industry (especially the food industry). Lactose is not only nontoxic but also cheaper than IPTG. Therefore, lactose is more suitable for the expression of pullulanase than IPTG. It should be noted that the activity of pullulanase decreased significantly when the induction intensity was too high, which may be because the protein synthesis speed was too fast to cause pullulanase to fold and form inclusion bodies. Therefore, the induction intensity should be strictly controlled in future studies.

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Effect of Glycine Concentration on Growth and Enzyme Production of E. coli

Because of the special double membrane structure, E. coli usually locates the secreted foreign proteins in the periplasmic space before releasing them into the fermentation broth through outer membrane leakage. However, this leakage is often very weak. For example, under the induction of 5 g/L lactose, the secretion efficiency (extracellular activity/total activity) of pullulanase was only 26.8%. According to the data [32, 33], glycine can enhance the permeability of the cell membrane and effectively improve the secretion efficiency of recombinant protein. The mechanism of this enhancement is believed to be that glycine can destroy the peptidoglycan layer of the cell wall, resulting in a loose cell wall and partial cell membrane damage [33]. In this section, the recombinant strain was cultured in glycine medium containing 0, 2.5, 5.0, 7.5, 10, and 12.5 g/L. The growth and enzyme production of the recombinant strain are shown in Fig. 5.25. On the one hand, increasing the concentration of glycine can effectively improve the secretion efficiency of pullulanase; on the other hand, glycine concentrations above 5 g/L can significantly inhibit the growth of the recombinant strain. Especially when the concentration was over 10 g/L, the concentration and enzyme production level of the recombinant strain decreased sharply. The optimal concentration of glycine was 10 g/L. Under these conditions, the total activity and extracellular secretion efficiency of pullulanase were 278.9 U/mL and 42.1%, respectively, which were 40.5% and 56.9% higher than those without glycine.

Fig. 5.25 Effects of glycine concentration on DCW (a) and pullulanase production (b) in E. coli. DCW (black), extracellular pullulanase activity (white), intracellular pullulanase activity (light gray). Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

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Effect of Lactose Flow Acceleration on Pullulanase Production by High-Density Fermentation of E. coli in a 3-L Fermenter

The shaking flask results showed that the induction intensity had a great influence on the growth and enzyme production of the recombinant strain, and an induction intensity that was too high was not conducive to the production of pullulanase. Therefore, we induced the recombinant strain by feeding lactose into a 3-L fermenter and compared the effect of lactose flow acceleration of 0.2, 0.4, and 0.8 g/L/h on pullulanase production. The fermentation results are shown in Fig. 5.26. When the lactose flow acceleration is 0.2 or 0.4 g/L/h, the bacterial volume can reach 105.2 and 103.2 g/L, respectively, while when the lactose flow acceleration is 0.8 g/L/h, the cell mass decreases to 92.5 g/L. This indicates that the lower induction intensity has little

Fig. 5.26 Effects of lactose feeding rate on DCW (a), extracellular pullulanase activity (b), and total pullulanase activity (c). ■, 0.2 g/L/h; ●, 0.4 g/L/h; ▲, 0.8 g/L/h. Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

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effect on the growth of the recombinant strain, while the higher induction intensity may lead to a decrease in bacterial volume due to excessive metabolic pressure. It can be seen from Fig. 5.26 that the acceleration of lactose flow has a great influence on the enzyme production of the recombinant strain, and too high or too low of an induction intensity is not conducive to enzyme production. The optimal lactose flow rate was 0.4 g/L/h. Under these conditions, the extracellular activity and total activity of pullulanase were 88.7 and 2731.1 U/mL, respectively. Although 10 g/L glycine was added to the fermentation medium, only 3.2% of pullulanase was secreted into the fermentation broth.

5.5.4

Effect of Induction Time on Pullulanase Production by High-Density Fermentation of E. coli in a 3-L Fermenter

The effect of induction time on the expression of recombinant protein in E. coli is mainly reflected in two aspects: on the one hand, if the induction time is too early and the cell metabolic load is too large, which is not conducive to growth; on the other hand, if the induction time is too late, the permeability of cell membrane phospholipids changes, which is unfavorable for the transport of recombinant protein. To determine the appropriate induction time, this experiment will start when the cell concentration is 6, 15, or 30 g/L. The fermentation results are shown in Fig. 5.27. Although the dry cell weight and total pullulanase activity were the highest when the recombinant strain was induced at a high bacterial concentration, the extracellular secretion efficiency of pullulanase was only 3.2%. However, when the recombinant strain was induced at the early growth stage, although the extracellular secretion efficiency of pullulanase was the highest (73.7%), the growth of the bacteria was seriously affected, and the dry cell weight was only 33.3 g/L. The results showed that the total enzyme activity also decreased to 941.6 U/mL; when the recombinant strain was induced at a bacterial concentration of 15 g/L, the growth and total activity of pullulanase were not significantly inhibited, which were 101.8 g/L and 2636.3 U/mL, respectively, and the extracellular pullulanase activity reached 731.3 U/mL, which was the highest under these three conditions. Therefore, the optimal induction time was 15 g/L bacterial concentration. The results showed that the induction time had a great influence on the extracellular secretion efficiency of pullulanase. According to the data reported [34], when cell growth enters the stable phase or the growth rate becomes slow, the content of phosphatidylethanolamine (PE) and phosphatidylglycerol (PG), which can promote protein transportation, will decrease sharply, and the cell membrane itself will become more rigid. Therefore, the induction of high concentrations of bacteria is very unfavorable to the extracellular secretion of pullulanase. The residual glycine concentration in the fermentation broth was further detected, and the results are

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Fig. 5.27 Effects of induction point on DCW (a), extracellular pullulanase activity (b), total pullulanase activity (c), and glycine concentration (d). ■, 6 g/L; ●, 15 g/L; ▲, 30 g/L. Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

shown in Fig. 5.27d. Under the condition of early growth induction, the consumption rate of glycine was significantly slower than that of high concentration induction, and it was not consumed until approximately 21 h of fermentation. At this time, the growth of the recombinant strain entered a stable phase. However, if induction occurred when the concentration of bacteria was 15 or 30 g/L, the growth of bacteria was in the early logarithmic phase when glycine was consumed. In the process of cell growth, glycine can replace L- and D-alanine in the peptidoglycan layer, which loosens the cell wall structure and enhances cell permeability. Therefore, when induction began at a bacterial concentration of 6 g/L, the medium containing enough glycine during the growth of the recombinant strain might be another reason for the high secretion efficiency of pullulanase.

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Effect of Glycine Feeding Strategy on Pullulanase Production by High-Density Fermentation of E. coli in a 3-L Fermenter

6

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Glycine feedingrate (g/L/h)

Glycine concentration (g/L)

Previous studies have shown that the production of pullulanase may be promoted by providing sufficient glycine during the growth of the recombinant strain, while glycine at high concentrations (>5 g/L) will seriously inhibit the growth of the recombinant strain. Therefore, by feeding a certain amount of glycine to maintain the residual glycine concentration of 1–5 g/L in the fermentation broth, it is possible to achieve the goal of not affecting the growth of the bacteria but also promoting the extracellular secretion of the recombinant protein. The acceleration of glycine flow can be determined by sampling the glycine concentration in the fermentation broth. The glycine concentration and glycine flow acceleration in the fermentation broth are shown in Fig. 5.28. Under the glycine feeding strategy, the dry cell weight, pullulanase activity and extracellular activity were 74.0 g/L, 2523.5 U/mL, and 1567.9 U/mL, respectively. Compared with those without glycine, although the dry cell weight and total pullulanase activity decreased by 27.5% and 4.3%, respectively, the extracellular pullulanase activity increased by 114.4% (Fig. 5.29). As shown in Fig. 5.30, the content of recombinant pullulanase (79 kDa) in the fermentation broth increased significantly with the progress of fermentation, which indicated that the glycine feeding strategy could effectively improve the extracellular secretion level of pullulanase in E. coli.

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Fig. 5.28 Glycine concentration and glycine feeding rate. Glycine concentration; glycine feeding rate, solid line; glycerol feeding rate, dotted line. Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

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Fig. 5.29 Effects of glycine feeding on DCW and enzyme production of E. coli. ■, DCW; ●, extracellular enzyme activity; ▲, total enzyme activity. Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

Fig. 5.30 SDS–PAGE analysis of fermentation supernatant (diluted 3 times). 1, 23 h; 2, 30 h; 3, 35 h; 4, 39 h; 5, 42 h; 6, 47 h

The comparison of the parameters of pullulanase production by recombinant E. coli in a 3-L fermenter is shown in Table 5.9. The total activity of pullulanase was the highest in all conditions at a rate of 0.4 g/L/h at a high bacterial concentration (30 g/L). However, due to the low extracellular secretion efficiency, the extracellular enzyme production per unit cell was only 0.86 × 103 U/g DCW. When induction

Lactose speed (g/L/h) 0.8 0.2 0.4 0.4 0.4 0.4

Induction time (DCW g/L) 30 30 30 6 15 15

Glycine feeding No No No No No Yes

Total activity (U/mL) 1172.8 ± 71.5 2142.3 ± 92.2 2731.1 ± 150.2 941.6 ± 36.7 2636.3 ± 115.9 2523.5 ± 90.8

Extracellular activity (U/mL) 66.3±2.7 30.1 ± 1.2 88.7 ± 2.9 693.8 ± 38.1 728.0 ± 44.4 1567.9 ± 61.1

DCW (g/L) 92.5 ± 3.9 104.0 ± 5.3 102.9 ± 4.5 33.3 ± 1.8 101.2 ± 3.9 70.5 ± 3.3

Extracellular productivity per cell (×103 U/g DCW) 0.72 ± 0.06 0.29 ± 0.03 0.86 ± 0.07 20.83 ± 2.40 7.19 ± 0.74 22.24 ± 2.00

Table 5.9 Comparison of parameters of pullulanase production by recombinant Escherichia coli in a 3-L fermenter. Reused with permission from Bioresource Technol. Zou C, Duan X, Wu J. 2014

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began under the condition of a low bacterial concentration (6 g/L), the extracellular enzyme production per unit cell reached 20.83 × 103 U/g DCW, which was 1.9 times that induced by 15 g/L DCW. However, early induction resulted in the lowest bacterial concentration and total enzyme activity. When induction began at 15 g/L DCW, the intensity of extracellular enzyme production was increased to 22.24 × 103 U/g DCW, which was the highest among all conditions. Therefore, early induction and glycine feeding strategies are important factors in improving extracellular enzyme production per unit cell.

5.5.6

“Mixed Conformation” Model of Pullulanase Expression in E. coli

Because of the double membrane structure of E. coli, the recombinant protein is usually located in the periplasmic space instead of directly releasing it into the culture medium. Then, a limited number of recombinant proteins are released to the outside of the cell through nonspecific leakage, which leads to the accumulation of a large number of recombinant proteins in the periplasmic space. According to 3.2.1, pullulanase from B. deramificans has a strong self-aggregation trend, so it will aggregate in a narrow periplasmic space to form active aggregates. In addition, due to the high molecular weight and complex structure of pullulanase, in the process of rapid synthesis, some inactive proteins may be formed due to a lack of time for folding. These proteins may aggregate to form some inactive inclusion bodies, which are not bioactive even when treated with surfactants. Therefore, the rapid expression of pullulanase in E. coli may form a variety of mixed conformations, such as normal pullulanase, misfolded “pullulanase,” pullulanase active aggregates and inactive inclusion bodies (as shown in Fig. 5.31). To reduce the formation of inclusion bodies, low-temperature induction was used to reduce the synthesis rate of recombinant protein, chemical chaperone was added to promote the correct folding of recombinant protein, and surfactant was added to promote the depolymerization of active aggregates, which significantly improved the soluble expression level of pullulanase. In this study, the extracellular secretion level of pullulanase in E. coli was significantly improved by optimizing the concentration of inducer, induction time and feeding glycine. By releasing the recombinant protein into the extracellular space, it can not only avoid the formation of inclusion bodies in the narrow periplasmic space but also simplify the extraction process and reduce the cost.

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Fig. 5.31 The mixed-state conformation model of pullulanase expressed in E. coli

5.6

Recombinant Expression of Thermobifida fusca Isoamylase in E. coli BL21(DE3)

Isoamylase (EC 3.2.1.68) is a kind of starch debranching enzyme. It can hydrolyze α-1,6-glycosidic bonds in glycogen, amylopectin, and β-limit dextrin. It can partially hydrolyze α-limit dextrin but cannot hydrolyze pullulan polysaccharide. Isoamylase has important application value in the starch processing industry. It can produce glucose, maltose, cyclodextrin, resistant starch, and so on by mixing with amylases such as glucoamylase, β-amylase, and CGTase. Due to the important role of isoamylase, a series of studies have been carried out for many years, and a variety of isoamylase-producing microbial strains have been found, including Pseudomonas amyloderamosa, Bacillus amyloliquefaciens [35], E. coli [36], Flavobacterium odoratum [37, 38], and Sulfolobus solfataricus [39]. With the development of genetic engineering technology, an increasing number of isoamylases have been recombined and expressed in E. coli or yeast [40]. However, there are still some problems, such as less developed isoamylases and low yields.

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T. fusca is a moderately thermophilic actinomycete isolated from composting. It can produce a variety of hydrolases (including glycoside hydrolase and cutinase) and can degrade almost all components except lignin and pectin in the plant cell wall. T. fusca is one of the main natural degradation microorganisms of the plant cell wall in nature [41]. At present, many glycoside hydrolases have been isolated and identified from T. fusca, including cellulase, xylanase, α-glucosidase, and α-amylase [42–44]. However, there is no report on whether T. fusca can produce starch debranching enzymes. In recent years, the publication of an increasing number of microbial genome sequences has provided a more convenient and effective way to exploit enzyme-coding gene resources from microorganisms. In 2007, the complete genome sequence analysis of T. fusca YX was completed. Database annotation showed that there were approximately 45 glycoside hydrolase coding genes in the T. fusca genome, and 5 of them may have starch debranching enzyme activity [41]. In this part, we compared the amino acid sequences of five potential starch debranching enzymes by blasting analysis in the NCBI database and analyzing the characteristic motifs of starch debranching enzymes and found that only Tfu_1891 may be the coding gene of isoamylase. Using the T. fusca genome as a template, the isoamylase-encoding gene was PCR amplified and recombinantly expressed in E. coli BL21 (DE3). After fermentation optimization, the isoamylase activity reached 6876.9 U/mL.

5.6.1

Cloning and Sequence Analysis of the T. fusca Isoamylase Gene

In 2007, Lykidis et al. on the basis of sequencing and analyzing the whole genome of T. fusca, considered that there were five potential pullulanase coding genes in the T. fusca genome, which were Tfu_0582, Tfu_0585, Tfu_0833, Tfu_0985, and Tfu_1891. First, the protein sequences of the five coding genes were submitted to NCBI for blast analysis. The results showed that they all had the characteristic sequence of the carbohydrate hydrolase family and three conserved key catalytic residues. As shown in Table 5.10, Tfu_0582, Tfu_0585, Tfu_0833, Tfu_0985, and Tfu_1891 correspond to branching enzymes, α-amylase, α-glucosidase, α-amylase, and debranching enzymes, respectively. In addition to the characteristic sequence of the carbohydrate hydrolase family and three conserved key catalytic residues, starch debranching enzymes also have a pullulanase characteristic motif (NWGYDP) or isoamylase characteristic motif (NYWGY) [45]. The characteristic motifs of these five proteins were analyzed, and only Tfu_1891 was found to have an isoamylase characteristic motif: NYWGY. Therefore, only Tfu_1891, found from amino acid sequence alignment,

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Table 5.10 Comparative analysis of the amino acid sequences of the five potential starch debranching enzymes in the T. fusca genome Potential starch debranching enzymes Tfu_0582 Tfu_0585 Tfu_0833 Tfu_0985 Tfu_1891

Amino acid length (AA) 749 655 544 605 707

Characteristic motifsa No No No No Yes

Results of database comparison and analysis Branching enzyme α-Amylase α-Glucosidase α-Amylase Debranching enzyme

a

Characteristic motifs of starch debranching enzymes include pullulanase characteristic motif (NWGYDP) and isoamylase characteristic motif (NYWGY) Table 5.11 A pairwise comparison between the amino acid sequences of the isoamylase. Reused with permission from Appl Microbiol Biotechnol. Duan X, Chen S, Chen J, Wu J. 2013 Sourcesa a b c d e f g

NCBI login number P10342.3 AAB63356.1 BAA82695.1 NP_343483.1 YP_289947.1 YP_003680738.1 ACY48768.1

Amino acid similarity (%) a b c d 100 60 62 29 100 71 31 100 31 100

e 31 37 35 49 100

f 32 36 34 48 77 100

g 29 31 32 52 54 52 100

a

a, P. amyloderamosa; b, F. odoratum; c, M. odoratus; d, S. solfataricus; e, T. fusca; f, N. dassonvillei; g, R. marinus

has the typical characteristics of starch debranching enzymes and is the object of further study. Using T. fusca genomic DNA as a template, the Tfu_1891 gene was PCR amplified, and gel electrophoresis showed that the size of the PCR product was slightly larger than 2000 bp, which was consistent with the gene size published in the database. Sequencing results showed that the target gene sequence was consistent with the published T. fusca YX Tfu_1891 sequence (NCBI accession number: CP000088.1:22160312218154). The length of Tfu_1891 was 2124 bp, and the length of the corresponding protein amino acid sequence was 707 AA. The amino acid sequence analysis showed that Tfu_1891 was a good candidate with the highest homology with isoamylase from N. dassonvillei, and the homology was approximately 77%. The homology with isoamylase from R. marinus, S. solfataricus, F. odoratum, M. odoratus, and P. amyloderamosa was 54%, 49%, 37%, 35%, and 31%, respectively (Table 5.11).

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Construction of Recombinant Plasmid and Strain

The cloning vector pMD18T/Tfu_1891 containing the target gene was subjected to double restriction enzyme digestion, gel recovery, and purification to obtain the target gene, which was connected with the expression vector pT7-7 recovered by the same restriction enzyme digestion and transformed into E. coli JM109. The recombinant plasmid was digested by NdeI and HindIII and showed two bands of approximately 2.5 kb and 2.1 kb, respectively, which were consistent with the expected size, demonstrating that the Tfu_1891 gene was successfully inserted into the expression vector (Fig. 5.32). The recombinant plasmid was transformed into strain E. coli BL21 (DE3), yielding recombinant strain E. coli BL21 (DE3) (pT7–7/ Tfu_1891).

5.6.3

Shake Flask Fermentation of the Recombinant Strain and Purification

The recombinant strain was cultured at 30 °C and induced by IPTG. The enzyme activity was measured at intervals. After 36 h of fermentation, the enzyme activity was the highest, reaching 879.2 U/mL. SDS–PAGE was used to detect the protein of the fermentation supernatant and the whole cell of the recombinant strain. There were bands in the fermentation broth and cells that were consistent with the theoretical molecular weight of the target recombinant protein, and the band size was approximately 79 kDa (Fig. 5.33). After the supernatant was purified by ammonium sulfate precipitation and DEAESepharose anion exchange chromatography, the specific activity of recombinant isoamylase was increased from 136.3 to 1223.5 U/mg, the purification fold was Fig. 5.32 Agarose gel electrophoresis of pT7-Tfu_1891 digested by NdeI and HindIII. Lane 1, plasmid digested with NdeI and HindIII; Lane M1, DL2000 DNA Marker; Lane M2, λ-EcoT14 I digest DNA Marker

1

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M2

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19329 7743 6223

pT7-7 Tfu_1891

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Fig. 5.33 SDS–PAGE analysis of the supernatant of the fermentation broth and purified enzyme. Lane 1, whole cell pellet; Lane 2, ammonium sulfate fraction; Lane 3, recombinant enzyme purified by chromatography; Lane M, molecular weight markers. Reused with permission from Appl Microbiol Biotechnol. Duan X, Chen S, Chen J, Wu J. 2013

Table 5.12 Summary of purification steps of the recombinant isoamylase. Reused with permission from Appl Microbiol Biotechnol. Duan X, Chen S, Chen J, Wu J. 2013

Purification steps Crude enzyme Ammonium sulfate precipitation Anion exchange chromatography

Total protein (mg) 72.6 4.9

Total activity (U) 36814.3 16062.1

Enzyme recovery (%) 100.0 43.6

Specific activity (U/mg) 136.3 885.1

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8.2 times, and the recovery rate was 9.0% (Table 5.12). It can be seen from Fig. 5.33 that the purified recombinant enzyme reached electrophoretic purity.

5.6.4

High-Density Fermentation of the Recombinant Strain in a 3-L Fermenter

Due to the high dosage of isoamylase in the application, a previous study found that recombinant isoamylase could reach 870.5 U/mL in shake flasks, and the enzyme production level was not high enough. To better meet the needs of subsequent application experiments, the fermentation process of the recombinant strain was preliminarily investigated in a 3-L fermenter.

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Overexpression of recombinant enzymes usually causes metabolic pressure on the host strain and then affects cell growth, protein expression and plasmid stability. To relieve the pressure of induced protein expression on the bacteria and improve the enzyme production level, 0.8 g/L/h lactose was added when the cell density of OD600 reached 50 (the fermentation period was approximately 15 h). It can be seen from Fig. 5.34 that when the cultivation time is 36 h, the bacterial concentration reaches the maximum, the biomass is 68.5 g/L at this time, and the cell concentration begins to decrease after 2 h of continuous cultivation. It can be seen from the enzyme activity curve that isoamylase activity began to appear after 3 h of induction (fermentation cycle 18 h), and then the enzyme activity increased rapidly with the extension of induction time. After 21 h of induction, the enzyme activity reached a maximum of 6876.9 U/mL. After that, the enzyme activity began to decrease slowly with decreasing cell concentration.

5.7

Recombinant Expression of T. Fusca Isoamylase in E. coli MDS42

E. coli MDS42 is a genome-reducing strain that does not have mobile elements and was constructed based on E. coli K12 MG1655 by deleting more than 700 nonessential genes (approximately 15% of the template genome), such as mobile DNA fragments (insertion sequence and recessive prophage) and toxic genes. The reduction in the genome reduces the redundant part of the E. coli gene and regulation loop, reduces unnecessary protein synthesis in the process of strain growth and

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metabolism and the nutrient concentration required in growth and metabolism, and improves the metabolic efficiency of the strain [46]. Recently, the E. coli MDS42 strain has been used to produce drug proteins and L-threonine on a small scale. In this part, the isoamylase gene (Tfu_1891) from T. fusca was cloned and expressed in E. coli MDS42, and the fermentation conditions were optimized in shake flasks and fermenters.

5.7.1

Construction of the Recombinant Strain E. coli MDS42/Tfu_1891-pSX2

The extracted plasmid Tfu_1891/pET24a(+) was used as a template, and the Tfu_1891 gene was PCR amplified and contained NdeI and KpnI sites at the ends. Agarose gel electrophoresis showed that the size of the PCR product was slightly larger than 2000 bp, which was similar to that of the known isoamylase gene fragment. Then, the PCR product was purified, A-added, and inserted into the pMD18T simple vector, yielding plasmid Tfu_1891/pMD18T. Then, the 1891 gene fragment was obtained by digesting the plasmid Tfu_1891/pMD18T with the restriction endonucleases NdeI and KpnI and was ligated with the expression vector pSX2, which was digested by the same restriction endonucleases. The ligation product was transformed into E. coli MDS42, yielding the recombinant strain E. coli MDS42/Tfu_1891-pSX2.

5.7.2

Expression of Recombinant Strain

The recombinant strain E. coli MDS42/Tfu_1891-pSX2 and E. coli MDS42 without the plasmid were inoculated into the corresponding LB liquid medium, cultured for 8 h, and then transferred to the corresponding TB fermentation medium. After incubation for 2 h at 37 °C and 200 rpm, 0.02 mmol/L IPTG was added for induction for 34 h. The isoamylase activity in the wall breaking supernatant of the recombinant strain E. coli MDS42/Tfu_1891-pSX2 was 1565.6 U/mL, but no isoamylase activity was detected in the fermentation supernatant and wall breaking supernatant of the control strain E. coli MDS42. SDS–PAGE analysis showed that there were bands with the same theoretical molecular weight (79 kDa) as the target protein in the recombinant strain E. coli MDS42/Tfu_1891-pSX2 (Fig. 5.35) [47], demonstrating that isoamylase was expressed in the recombinant strain E. coli MDS42/Tfu_1891pSX2.

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Fig. 5.35 SDS–PAGE analysis of protein expression of blank and recombinant strains. Lane M, molecular weight markers; Lane 1, wall breaking supernatant of the control strain; Lane 2, wall breaking supernatant of recombinant bacteria; Lane 3, wall breaking precipitate of recombinant bacteria. Reused with permission from Appl Biochem Biotechnol. Ran H, Wu J, Wu D, Duan X. 2016

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5.7.3

Optimization of Fermentation Conditions for Recombinant Strain E. coli MDS42/Tfu_1891-pSX2 by Shake Flask Cultivations

5.7.3.1

Effect of Induction Temperature on the Growth and Enzyme Production of the Recombinant Strain

The induction temperature is one of the important factors affecting the growth of recombinant bacteria and the expression of recombinant proteins [4]. To investigate the effects of different induction temperatures on the growth and enzyme production of the recombinant strain, 0.2 mmol/L IPTG was added at 25, 30, and 37 °C. The results are shown in Fig. 5.36. With increasing induction temperature, the biomass of bacteria decreased gradually. The highest cell concentrations were 8.2 g/L at 25 °C, 7.4 g/L at 30 °C, and 6.9 g/L at 37 °C. However, the enzyme production was contrary to the growth of the bacteria. At 37 °C, the isoamylase activity was the highest, which was 1565.6 U/mL, which was 2.5 times and 3.7 times higher than that at 30 °C and 25 °C, respectively. At the same time, as shown in Fig. 5.37, SDS– PAGE analysis showed that more inclusion bodies were formed at 37 °C than at 30 and 25 °C. Generally, when induced by high temperature, the synthesis rate of recombinant protein is too fast, resulting in a reduction in the folding efficiency of recombinant protein and the easier formation of insoluble inclusion bodies in the cell [28]. Comprehensive analysis showed that the reason for the poor growth of the bacteria under the high-temperature induction condition may be the overexpression of recombinant enzyme, which caused pressure on the growth and metabolism of the recombinant strain, but the effect was not significant. Under high-temperature induction conditions, the isoamylase activity was significantly higher than that at

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Temperature (°C) Fig. 5.36 Comparison of DCW and intracellular isoamylase activity in E. coli MDS42 induced with 0.02 mmol/L IPTG at 25, 30 and 37 °C. DCW (black), isoamylase activity (white) Fig. 5.37 SDS–PAGE analyses of intracellular soluble and insoluble fractions at different induction temperatures. M, molecular weight markers; 1, 2, 3, intracellular soluble fraction induced with 0.02 mM IPTG at 37, 30 and 25 °C, respectively; 4, 5, 6, intracellular insoluble fraction induced with 0.02 mM IPTG at 37, 30, and 25 °C

other temperatures. Therefore, 37 °C was selected as the temperature for the induction of enzyme production in shake flask fermentation.

5.7.3.2

Effect of Inducer Type and Concentration on the Growth and Enzyme Production of the Recombinant Strain

The promoter of the expression vector pSX2 used in this study is T5/LacO, which is a strict and strong promoter [48]. IPTG or lactose are generally used as inducers to

Recombinant Expression of Starch Debranching Enzymes in Escherichia coli

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Fig. 5.38 Comparison of DCW and intracellular isoamylase activity in E. coli MDS42 after 34 h of culture in TB medium at different lactose (a) and IPTG (b) concentrations. (a) Comparison of DCW and isoamylase activity at different lactose concentrations ranging from 1.25 to 20 g/L. (b) Comparison of DCW and isoamylase activity at different IPTG concentrations ranging from 0 to 0.4 mmol/L. DCW (black), isoamylase activity (white)

induce protein expression. IPTG is a kind of sulfur-containing lactose analog, which will not be decomposed in the induction process, and the induction intensity is high; however, when lactose is used as an inducer, it will be degraded by β-galactosidase, and the induction intensity is low. Therefore, to explore the effects of IPTG and lactose on the growth and enzyme production of the recombinant strain, lactose at final concentrations of 1.25, 2.5, 5.0, 10, 15, and 20 g/L and IPTG at 0, 0.02, 0.05, 0.1, 0.15, 0.2, 0.3, and 0.4 mmol/L were added to the culture medium at 37 °C. The cell dry weight and isoamylase activity in the cell wall breaking supernatant were measured. As shown in Fig. 5.38, both lactose and IPTG could induce the expression of isoamylase in E. coli MDS42, but no isoamylase activity was detected in shake flasks without inducer. At the same time, as shown in Fig. 5.38a, with the increase in lactose-induced concentration, cell dry weight and isoamylase activity showed a trend of first increasing and then decreasing. When the concentration of lactose reached 5 g/L, the cell dry weight reached the maximum value of 8.3 g/L, while the activity of isoamylase reached the maximum value when the final concentration of lactose increased to 10 g/L, which was 951.5 U/mL. The isoamylase activity of lactose was lower than that of IPTG. As shown in Fig. 5.38b, isoamylase activity increased with increasing IPTG concentration when the final concentration of induction was between 0 and 0.05 mmol/L IPTG. The isoamylase activity was 1928.7 U/mL when the IPTG concentration was 0.05 mmol/L, which was twice as high as that of lactose. It should be noted that when the IPTG concentration was higher than 0.05 mmol/L, the biomass (between 6.9 and 6.3 g/L) and isoamylase activity did not change much. The above results showed that a high concentration of IPTG did not inhibit the growth of cells, which may be due to the deletion of many mobile gene fragments of E. coli MDS42 [49], which greatly improved the stability of the genome and plasmid [50], thus reducing the toxicity of IPTG to recombinant cells.

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The above results showed that in the E. coli MDS42 protein expression system, the expression level of the target protein was significantly higher when IPTG was used as an inducer than when lactose was used as an inducer, which was different from previous research results in E. coli K12 LJ110 and E. coli BL21(DE3) protein expression systems [51–53]. To further investigate the induction effect of lactose, we used the optimized technology of isoamylase production by recombinant E. coli BL21(DE3) in a 3-L fermenter (isoamylase activity was 6876.9 U/mL), and the expression of isoamylase in recombinant E. coli MDS42 was induced by adding lactose into a 3-L fermenter. The results showed that the expression level of isoamylase was very low in the E. coli MDS42 protein expression system. The reason is that when lactose is used as an inducer, the expression of β-galactosidase in cells has a significant effect on the expression of inducible protein. Because β-galactosidase can hydrolyze lactose into glucose, galactose, and isomeric lactose, the repressor protein can be inactivated when it combines with isomeric lactose in the cell, and mRNA synthesis, gene transcription, and target protein synthesis can be promoted. The expression of β-galactosidase in E. coli BL21(DE3)/Tfu_1891pET24a(+) and E. coli MDS42/Tfu_1891-pSX2 was compared when induced by lactose. The expression of β-galactosidase was determined by adding lactose at a final concentration of 10 g/L to the fermentation medium of the two recombinant strains. After induction at 37 °C for 34 h, the activity of β-galactosidase in cells was determined. Finally, the β-galactosidase activity in the E. coli BL21 (DE3) expression system was 0.6 U/mg DCW, while that in the E. coli MDS42 expression system was only 0.08 U/mg DCW. Therefore, in the E. coli MDS42 protein expression system, the low activity of β-galactosidase may be the reason for the poor effect of lactose induction.

5.7.4

Optimization of Fermentation Conditions for Recombinant Strain E. coli MDS42/Tfu_1891-pSX2 by 3-L Fermenter Cultivation

After optimization of fermentation conditions, the isoamylase activity was 1928.7 U/ mL. Compared with 879.2 U/mL in E. coli BL21(DE3), the yield of isoamylase in E. coli MDS42 was improved. To further improve the yield of isoamylase and lay the foundation for industrial production, the fermentation process of the recombinant strain in a 3-L fermenter was optimized. First, for intracellular enzymes, better growth conditions are a prerequisite for the efficient expression of intracellular enzymes [51]. The glycerol feeding rate will affect the metabolic flow, thus affecting the biomass, enzyme expression, and the formation of byproducts such as acetic acid [54]. Therefore, it is very important to select the appropriate glycerol feeding rate for the fermentation of recombinant bacteria. In E. coli MDS42, approximately 15% of the gene fragments were deleted, which improved the metabolic efficiency of the strain and reduced the energy

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demand in the fermentation process [55]. Therefore, we modified the glycerol feeding strategy described by Fang et al. [16]. As shown in Fig. 5.39a, when the glycerol in the medium was consumed, the exponential feeding medium was started to control the specific growth rate of 0.165 h-1. When the growth of bacteria was close to the late logarithmic stage (approximately 18 h of fermentation), exponential feeding was stopped, the feeding rate was reduced to 41.5 mL/h at the current flow rate for 4 h, and a constant flow rate was added to the end of the fermentation process. In addition, the expression of heterologous proteins usually leads to a metabolic burden in cells, thus reducing cell concentration, protein expression and plasmid stability [53]. Therefore, to obtain a higher level of heterologous protein expression, the induction temperature, induction time, and inducer concentration can be optimized to adjust the metabolic burden of cells.

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Effect of Induction Temperature on the Growth and Enzyme Production of the Recombinant Strain

In shake flask fermentation, the isoamylase activity under high temperature was obviously higher, which provided a certain reference for the scale-up experiment of isoamylase. Therefore, we first investigated the effects of temperature on the growth and enzyme production of the recombinant strain in a 3-L fermenter. When the cell concentration OD600 reached approximately 40, isoamylase expression was induced at 25, 30, and 37 °C by adding 0.05 mmol/L IPTG. As shown in Fig. 5.39a, when the induction temperature is 30 °C, the growth of the strain is the best. After 36 h of fermentation, the cell concentration was 96.2 g/L, while at 37 and 25 °C, the cell concentrations were 85.2 and 66.3 g/L, respectively. Therefore, temperatures that are too high or too low have adverse effects on the growth of the strain. In addition, we also detected glycerol and acetic acid in the fermentation process of the recombinant strain and found that there was no glycerol accumulation in the whole fermentation process. As shown in Fig. 5.39b, acetic acid production increases with increasing temperature, but its content is below 1.0 g/L. In theory, it will not affect the growth of bacteria. The isoamylase activity at 37 °C was significantly higher than that at 30 and 25 °C. The isoamylase activity and yield reached maximum values of 13,813.3 U/mL and 12.0 mg/mL, respectively, at 37 °C for 20 h. However, the maximum isoamylase activity and yield were 7856 U/mL and 6.8 mg/mL and 3137.0 U/mL and 2.7 mg/mL at 30 and 25 °C, respectively. In summary, if the temperature is too low, the growth of the bacteria is limited, and the cell metabolism is slow, which is not conducive to the growth of the bacteria and the production of enzymes. Meanwhile, if the temperature is too high, the cell metabolism speed is too fast, and the intracellular protein synthesis speed is too fast, it will have a certain negative impact on the growth of the strain, but the content of intracellular soluble protein is higher than that in the low-temperature environment. Therefore, the optimum temperature for enzyme production was 37 °C.

5.7.4.2

Effect of Induction Time on the Growth and Enzyme Production of the Recombinant Strain

To determine the optimal induction time, when the recombinant strain OD600 reached 3, 40, and 80, 0.05 mmol/L IPTG was added to induce enzyme production. As shown in Fig. 5.40a, when the OD600 was induced to 3, cell growth was seriously inhibited, growth stopped after 30 h of fermentation, and the cell biomass was only 30.3 g/L. However, when the OD600 was 40 and 80, the cell growth was normal, and there was no significant difference. After 39 h of fermentation, the cell concentrations were 87.6 g/L and 85.2 g/L, respectively. During the fermentation process of the recombinant strain, the fermentation fermenter was monitored in real time. In the early induction, after 13.5 h of fermentation, the feeding pump was turned off, and the dissolved oxygen rebounded after 1.5 h, which indicated that

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Fig. 5.40 Comparison of the time profiles for biomass (a), recombinant isoamylase expression (b), glycerol (c) and acetic acid concentration (d) induced at OD600 of 3, 40 and 80 in a 3-L fermenter. Reused with permission from Appl Biochem Biotechnol. Ran H, Wu J, Wu D, Duan X. 2016

glycerol accumulated in the fermentation process. When the concentration of bacteria was detected by timely sampling, abnormal growth of bacteria was found. After 21.5 h of fermentation, feeding was stopped. In the middle and late induction feeding stages, dissolved oxygen rebounded immediately after the feeding pump was turned off, indicating no glycerol accumulation. The results showed that when the OD600 was 3, glycerol accumulated seriously at 21 h, which was 8.5 g/L, while acetic acid production increased to 6.8 g/L and then began to decline. When the OD600 was 40 and 80, acetic acid production and glycerol accumulation in the fermentation fermenter were much lower than those in early induction. It should be noted that when the OD600 was 40, there was no glycerol accumulation in the fermentation fermenter, and the acetic acid content was less than 1.0 g/L. Shiloacha et al. reported that acetic acid is the uncoupling agent for phosphorylation and electron transfer in the TCA cycle [49]. When the concentration of acetic acid in the fermentation

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medium is greater than 2 g/L, the growth of bacteria will be slowed or blocked, and the expression of recombinant protein will be inhibited. Therefore, when the OD600 was 3, IPTG induced the expression of the target protein too early, which caused pressure on cell growth and metabolism, resulting in glycerol accumulation, producing a large amount of acetic acid, and finally inhibiting growth. This is a common phenomenon in the fermentation of E. coli MDS42 and E. coli BL21 [16, 52, 56]. The enzyme production at different induction stages is shown in Fig. 5.40d. Isoamylase activity first increased and then decreased with increasing induction time. When the OD600 was 3, the growth of the recombinant strain was inhibited, resulting in the lowest isoamylase activity of 3644.0 U/mL and an isoamylase yield of 3.0 mg/mL. When the induced OD600 was 40, the isoamylase activity and yield were 13813.3 U/mL and 12.0 mg/mL, respectively, which were 1.1 times the highest protein expression level (12183.4 U/mL, 10.6 mg/mL) when the induced OD600 was 80, which was 2 times that of recombinant isoamylase in E. coli BL21(DE3). According to the growth and enzyme production results, the optimal induction time was 40 when the OD600 was reached.

5.7.4.3

Effect of Inducer Concentration on the Growth and Enzyme Production of the Recombinant Strain

Too low of an IPTG concentration will lead to insufficient induction intensity, and too high of an IPTG concentration may be harmful to cell growth [57]. Therefore, it is very important to select the appropriate IPTG concentration for the expression of heterologous proteins. In the previous shake flask fermentation, 0.05 mmol/L IPTG was enough to induce the expression of recombinant enzyme, and higher IPTG concentrations (greater than 0.4 mmol/L) had no effect on cell growth. However, the cell concentration in the 3-L fermenter was much higher than that in the shake flask, so more IPTG may be needed to induce the expression of the recombinant enzyme. To investigate the effect of IPTG concentration on cell growth and enzyme production in a 3-L fermenter, 0.05, 0.15, and 0.25 mmol/L were added to logarithmic medium to induce enzyme production. As shown in Fig. 5.41a, there was no significant difference in the growth trend of bacteria under three different IPTG induced concentrations, and the cell concentrations were 85.2, 80.1, and 82.4 g/L at 36 h, respectively. In addition, the contents of acetic acid and glycerol in the fermentation fermenter were also detected, as shown in Fig. 5.41b. There was no accumulation of glycerol in the fermentation fermenter medium induced by different concentrations of IPTG, and the acetic acid content was lower than 1.0 g/L. As shown in Fig. 5.41c, the trend of enzyme production was consistent under different IPTG concentrations. With increasing induction time, the isoamylase activity also increased. After 36 h, the enzyme activity reached a maximum and then began to decline. The isoamylase activity increased with increasing IPTG concentration, but the increasing range decreased slowly. When the dosage of IPTG was 0.25 mmol/L, the maximum enzyme activity of isoamylase was 22983.0 U/mL, and the yield was 20.0 mg/mL, which was 1.1 and 1.7 times higher

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Fig. 5.41 Comparison of the time profiles for biomass (a), recombinant isoamylase expression (b) and acetic acid concentration (c) induced with different IPTG concentrations in a 3-L fermenter. Reused with permission from Appl Biochem Biotechnol. Ran H, Wu J, Wu D, Duan X. 2016

than that of 0.15 and 0.05 mmol/L, respectively. Since the isoamylase activity did not change much when the IPTG dosage was 0.15 mmol/L and 0.25 mmol/L, it was unnecessary to increase the amount of IPTG (more than 0.25 mmol/L) to increase the expression of isoamylase.

5.7.5

Comparison of Fermentation Parameters of Recombinant Isoamylase Production in Shake Flasks and 3-L Fermenters

As unit cell productivity is an important parameter in the fermentation process of recombinant enzyme production [28], we calculated and summarized the unit cell productivity in shake flasks and fermentation fermenters. As shown in Table 5.13, when 0.02 mmol/L IPTG was added to the flask at 37 °C to induce enzyme

3 80 40 40 40 40

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E. coli BL21(DE3) [47]

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DCW (g/L) 8.2 7.4 6.9 6.5 – 3644.0 12,183.4 13,813.3 20,082.7 22,983.0 6876.9

Activity (U/mL) 425.6 626.1 1565.6 1928.7 879.2 151.8 312.4 383.7 557.8 638.4 191.0

Productivity (U/mL/h) 11.8 17.4 43.5 53.6 24.4

127.0 143.0 162.1 250.7 278.9 100.4

Productivity per cell (×103U/gcell) 51.9 84.6 226.9 296.7 –

Table 5.13 Comparison of parameters for intracellular isoamylase production in shake flasks and 3-L fermenters. Reused with permission from Appl Biochem Biotechnol. Ran H, Wu J, Wu D, Duan X. 2016

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production, the unit cell productivity was 226.9 × 103 U/g cell, which was 2.7 and 4.4 times higher than that at 30 and 25 °C, respectively. The reason may be that the protein synthesis rate and cell metabolism are faster at higher induction temperatures (37 °C), which ultimately leads to high cell productivity. In the process of highdensity fermentation in a fermenter, compared with shaking flask fermentation, the nutrient and nutrient conditions of the medium are very sufficient, dissolved oxygen and pH can be controlled more accurately, the growth and metabolism of bacteria are fast, and the speed of protein production in cells is higher than that in shake flasks. When protein expression was induced at different times, the unit cell productivity was 162.1 × 103 U/gcell at medium induction, which was 1.3 and 1.1 times higher than that of early induction and late induction, respectively. When the concentration of IPTG was 0.25 mmol/L, the protein synthesis rate and cell productivity were 638.4 U/mL/h and 278.9 × 103 U/gcell, respectively, which were 3.3 and 2.8 times that in E. coli BL21 (DE3), respectively. The results showed that IPTG could induce the protein expression of E. coli MDS42 in a wide range. When 0.05 mmol/L IPTG was added in the late induction period and 0.05 mmol/L and 0.25 mmol/L IPTG were added in the medium-term stage, the protein expression levels were much higher. Meanwhile, the synthesis rate and unit cell productivity of the recombinant isoamylase strain were higher than those in E. coli BL21 (DE3). Acknowledgments This chapter was modified from the paper published by our group in Bioresour. Technol. (Duan X, Chen J, Wu J. et al., 2013) [5], Appl. Environ. Microbiol. (Duan X, Chen J, Wu J. 2013) [58], Bioresour. Technol. (Zou C, Duan X, Wu J. 2014) [53], Appl. Environ. Microbiol. (Duan X, Wu J. 2015) [59] Bioresour. Technol. (Duan X, Zou C, Wu J. 2015) [7], and Appl. Biochem. Biotechnol. (Ran H, Wu J, Wu D, Duan X. 2016) [30]. The related contents are reused with permission.

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Chapter 6

Production of Starch Debranching Enzymes in Bacillus Strains Kang Zhang, Zhengfei Yan, and Jing Wu

Abstract Bacillus is a kind of Gram-positive bacterium that has a single membrane, and Bacillus subtilis, Bacillus licheniformis, Bacillus amyloliquefaciens, Bacillus megatherium, and Brevibacillus choshinensis have been constructed as expression systems for the production of various proteins. In this study, the expression of starch debranching enzymes in Bacillus was investigated, mainly focusing on the expression of pullulanase in B. choshinensis and B. subtilis. First, the expression of pullulanase in B. choshinensis was improved through shake-flask cultivation and 3-L fermenter optimization. Because magnesium ions can promote the expression of “high activity” pullulanase in B. choshinensis, the mechanism of this promoting effect was studied from the aspects of morphological changes, transcription, and expression of related genes in B. choshinensis. Second, to improve the expression of pullulanase in B. subtilis, a CRISPR/Cas9 gene editing system was established in B. subtilis, and a host strain with improved fermentation performance was obtained through gene disruption. Then, high-efficiency expression of pullulanase in B. subtilis was achieved by optimizing the host strain protease concentration, increasing the negative charge of the cell wall, optimizing the expression vector, and optimizing fermentation conditions. Keywords Brevibacillus choshinensis · Bacillus subtilis · Fermentation cultivation optimization · Magnesium ion · Host strain modification · Expression vector

K. Zhang · Z. Yan · J. Wu (✉) State Key Laboratory of Food Science and Resources, Jiangnan University, Wuxi, China School of Biotechnology and Key Laboratory of Industrial Biotechnology Ministry of Education, 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. 2023 J. Wu, W. Xia (eds.), Industrial Starch Debranching Enzymes, https://doi.org/10.1007/978-981-19-7026-9_6

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Introduction

Bacillus is a kind of aerobic Gram-positive bacteria that can form spores. The cell wall does not contain endotoxin. It is mostly found in soil, water, air, and decaying organic matter [1]. Since the genus Bacillus was proposed in 1872, it has a history of more than 100 years. Many species of Bacillus, including Bacillus subtilis, Bacillus amyloliquefaciens, Bacillus licheniformis, Bacillus megaterium, and Brevibacillus choshinensis can be used as expression hosts for the production of various industrial enzymes. Until 1989, 50% of industrial enzymes were produced by Bacillus. B. choshinensis is a Gram-positive bacterium with short rod-shaped cells [2]. It was originally screened from the soil by Udaka et al. Because of its strong ability to secrete proteins out of the cell, it has now been developed into a recombinant protein expression system. When expresses recombinant proteins, B. choshinensis has the following advantages: can synthesize a large number of recombinant proteins with biological activity; directly secrete protein into fermentation broth, which is conducive to downstream extraction; almost no protease is produced, which prevents the recombinant protein from being degraded; easy to cultivate and does not produce spore; simple molecular operation, genetic stability; nonpathogenic and safe bacteria, can be used in the food industry. The above advantages promote the wide application of B. choshinensis in industrial enzyme expression. B. subtilis, as a model organism in the genus Bacillus, is the first strain in the genus Bacillus to be used as an expression host strain. B. subtilis is a food-safe strain and is listed as generally recognized as safe (GRAS) by the US Food and Drug Administration. It has long been used in the preparation of fermented foods. At present, researchers have used B. subtilis, B. licheniformis, and B. amyloliquefaciens as hosts to express pullulanase. Wang et al. studied the effects of different promoters and signal peptides on the expression of pullulanase in B. subtilis using the promoter Papr and signal peptide SPacB to increase the extracellular activity of pullulanase to 2.82 U/mL [3]. Zhang et al. improved the expression of pullulanase in B. subtilis by optimizing the feeding strategy and fermentation conditions, which yielded the highest activity of 102.75 U/mL [4]. Liu et al. improved the expression of pullulanase in B. subtilis through host strain modification that knocked out eight extracellular protease genes, signal factor F (spoIIAC) and surfactin (srfAC), and promoter optimization, which yielded the highest activity of 625.5 U/mL [5]. Meng et al. improved the expression of pullulanase in B. subtilis by mining strong promoters from transcriptome data and promoter combinations, which yielded a triple promoter PsodA + fusA + amyE that showed the highest activity of 1555 U/mL in a 50-L fermenter cultivation [6]. Abdel-Naby et al. used B. licheniformis as a host to express pullulanase and carried out batch fermentation and continuous fermentation with immobilized cells, and the yields of pullulanase were 14.8 and 16.8 U/mL, respectively [7]. Sun et al. knocked out the sporeproducing gene of B. amyloliquefaciens and used the modified strain to express pullulanase. By optimizing the fermentation conditions, the output of pullulanase reached 6.9 ASPU/mL. In short, although the expression level of pullulanase

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expressed by Bacillus as a host is relatively low, it can secrete most of the pullulanase outside the cell. In addition, many pullulanases with excellent properties are derived from Bacillus. Selecting a suitable Bacillus system for expression may achieve high-efficiency secretion and expression of pullulanase. In this study, strategies to improve the expression of pullulanase in B. choshinensis and B. subtilis were investigated, mainly focusing on fermentation optimization, host strain modification, promoter, and signal peptide optimization.

6.2

Recombinant Expression of Pullulanase in B. choshinensis

B. choshinensis is a kind of Bacillus that shows excellent secretory capability. It has been engineered to produce a very low level of extracellular protease and cannot form spores in the late stage of fermentation. B. choshinensis was also confirmed to be nonpathogenic. These characteristics make B. choshinensis a favorable host strain to express pullulanase. Since 2015, Zou et al. expressed Bacillus deramificans pullulanase in B. choshinensis. The fermentation media and conditions were optimized on the basis of shake flasks and 3-L fermenter cultivations. The mechanism by which Mg2+ promotes the expression of high-specific activity and thermostable pullulanase was investigated.

6.2.1

Construction of the Recombinant Strain B. choshinensis (pNCMO2/pulA-d2)

First, the pulA-d2 gene fragment was obtained by PCR amplification with plasmid pET-24a-ompA/pulA-d2 as a template. Then, the fragment was ligated with the pMD-18 T vector and transferred to E. coli JM109 by chemical transformation. The pulA-d2 gene fragment was obtained by double restriction enzyme digestion and ligated with the vector fragment pNCMO2, which is an E. coli-B. choshinensis shuttle vector. The ligation product was first transferred to E. coli JM109 by chemical transformation and then transformed into the expression host B. choshinensis by electrotransformation, yielding the recombinant strain B. choshinensis (pNCMO2/pulA-d2). The recombinant bacteria were transferred to a shake flask containing TM medium and cultivated for 48 h. The pullulanase activity in the supernatant of the fermentation broth was 50.1 U/mL. After the bacteria were collected, the cells were broken, and the pullulanase activity of the supernatant of the cell disruption solution was 0.9 U/mL. After the broken pellet was suspended, no pullulanase activity was detected. SDS–PAGE analysis was performed on the supernatant of the fermentation broth, the supernatant of the cell disruption solution and the precipitate, and the results are shown in Fig. 6.1. There

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Fig. 6.1 SDS–PAGE analysis of fermentation broth. Lane M, molecular weight markers; Lane 1, supernatant of fermentation broth; Lane 2, supernatant of cells after disruption; Lane 3, precipitation of cells after disruption. The arrow indicates pullulanase. The 14-kDa protein band in lanes 2 and 3 corresponds to the cells lyzed by adding lysozyme. Reused with permission from J Ind Microbiol Biotechnol. Zou C, Duan X, Wu J. 2015

are very few intracellular soluble proteins and very few inclusion bodies. Most of the pullulanase is secreted into the fermentation broth. This shows that B. choshinensis is a host capable of soluble secretion and expression of pullulanase.

6.2.2

Optimization of Shake-Flask Cultivations

6.2.2.1

Original Fermentation Medium Optimization

The commonly used media, including 2SY, TM, TM-derived medium, LB, TB, TSB, and M9, were individually used as the original medium for shake-flask cultivations. After shake-flask cultivations, the pullulanase activity of TM was 49.4 U/mL, which was much higher than that of the others. Therefore, TM was used as an original medium in further optimization (Fig. 6.2).

6.2.2.2

Carbon Source Optimization

The carbon source of 10 g/L glucose in TM medium was replaced with glycerol, sucrose, maltose, lactose, and soluble starch at the same concentration. After shakeflask cultivations, the pullulanase activity of glucose showed the highest activity and DCW. Then, the concentration of glucose was optimized at 10, 20, 30, and 40 g/L. After shake-flask cultivations, 20 g/L glucose showed the highest activity and DCW,

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Fig. 6.2 Effects of the initial medium on cell growth and pullulanase production. DCW (black), pullulanase activity (white). Reused with permission from J Ind Microbiol Biotechnol. Zou C, Duan X, Wu J. 2015

Fig. 6.3 Effects of carbon source (a) and glucose concentration (b) on cell growth and pullulanase production. DCW (black), pullulanase activity (white). Reused with permission from J Ind Microbiol Biotechnol. Zou C, Duan X, Wu J. 2015

which were 54.3 U/mL and 3.1 g/L, respectively. Therefore, 20 g/L glucose was used as the carbon source in further optimization (Fig. 6.3).

6.2.2.3

Nitrogen Source Optimization

The nitrogen source in the medium was replaced with (NH4)2SO4, NH4Cl, yeast extract, beef extract, peptone, cottonseed meal, polypeptone, meat extract, and

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Fig. 6.4 Effects of nitrogen source (a) and nitrogen source concentration (b) on cell growth and pullulanase production. DCW (black), pullulanase activity (white). BE beef extract, PO polypeptone, PE peptone, YE yeast extract, SO soytone, CO cottonseed meal, ME meat extract, BE+YE 10 g/L beef extract and 5 g/L yeast extract, BE+CO 10 g/L beef extract and 5 g/L cottonseed meal; 10B5C, 10 g/L beef extract and 5 g/L cottonseed meal; 20B5C, 20 g/L beef extract and 5 g/L cottonseed meal; 30B5C, 30 g/L beef extract and 5 g/L cottonseed meal; 40B5C, 40 g/L beef extract and 5 g/L cottonseed meal; 30B2C, 30 g/L beef extract and 2 g/L cottonseed meal; 30B10C, 30 g/L beef extract and 10 g/L cottonseed meal. Reused with permission from J Ind Microbiol Biotechnol. Zou C, Duan X, Wu J. 2015

soytone at a concentration of 15 g/L. After shake-flask cultivations, the inorganic nitrogen sources (NH4)2SO4 and NH4Cl cannot sustain the growth of bacteria. The pullulanase activity of beef extract was the highest (57.8 U/mL), while the DCW was much lower (1.5 g/L). The pullulanase activities of yeast extract and cottonseed meal were not very high, while their DCW values were relatively high, 2.8 and 2.9 g/L, respectively. Then, the beef extract combined with yeast extract and cottonseed meal could enhance both the activity and DCW. When the concentration of cottonseed meal was fixed at 5 g/L, the pullulanase activity increased with increasing concentrations of yeast extract up to 30 g/L. When the concentration of yeast extract was fixed at 30 g/L, the pullulanase activity was not significantly influenced by the concentration of cottonseed meal, while the DCW was significantly influenced by the concentration of cottonseed meal, which shows the optimal cottonseed meal concentration of 5 g/L. Overall, the pullulanase activity was the highest (109.7 U/ mL) by 30 g/L yeast extract and 5 g/L cottonseed meal (Fig. 6.4).

6.2.2.4

Metal Ion Optimization

The effects of metal ions on the expression of pullulanase were investigated at a concentration of 10 mmol/L. After shake-flask cultivations, Na+ and K+ had no significant influence on cell growth and pullulanase activity. Except for Cu2+, which significantly influenced cell growth, and Zn2+ and Fe2+, which slightly influenced cell growth, the other metal ions had no significant influence on cell growth. Ca2+, Cu2+, and Zn2+ show negative pullulanase activity, while Mg2+, Mn2+, and Fe2+

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Fig. 6.5 Effects of metal ion (a) and MgCl2
6H2O concentration (b) on cell growth and pullulanase production. DCW (black), pullulanase activity (white). Reused with permission from J Ind Microbiol Biotechnol. Zou C, Duan X, Wu J. 2015 Table 6.1 Variables and levels of CCD. Reused with permission from J Ind Microbiol Biotechnol. Zou C, Duan X, Wu J. 2015

Factor (g/L) Glucose Yeast extract MgCl2
6H2O

Code A B C

Level -1.68 3.18 13.18 3.27

-1 10 20 6

0 20 30 10

1 30 40 14

1.68 36.82 46.82 16.73

show positive pullulanase activity, especially Mg2+. The concentration of MgCl2
6H2O was optimized to further improve pullulanase activity at 2, 6, 10, 14, and 18 g/L. After shake-flask cultivations, 10 g/L MgCl2
6H2O showed the highest pullulanase activity of 386.2 U/mL. The positive effect of Mg2+ on the higher pullulanase activity was not due to a higher expression level because of the higher specific activity, whose mechanism will be discussed in the next section (Fig. 6.5).

6.2.2.5

Key Factor Optimization Using Response Surface Methodology

Based on the above single-factor optimization, the concentrations of glucose, yeast extract, and MgCl2
6H2O were further optimized using response surface methodology. As shown in Table 6.1, optimization was performed in the form of three factors and five separate levels. The design and experimental results are shown in Table 6.2. Using Design-Expert 8.0.5b software to analyze the results, a quadratic model was obtained: Y = 388.23 + 54.79(A) – 22.62(B) + 44.82(C) + 5.64(AB) + 8.49(AC) – 11.29 (BC) – 38.6(A2) – 61.02(B2) – 48.7(C2), where Y represents pullulanase activity. The adequacy of the model was tested by analysis of variance (ANOVA); the results are shown in Table 6.3. The P value of the model was F lauric acid > myristic acid > palmitic acid > stearic acid, and a linear correlation occurred between the carbon numbers and CI values (R2 = 0.94). The possible explanation was considered to be the increased steric hindrance and decreased solubility in water as the length of the carbon chain increased, as well as the lower molecular weight and length of the helical structure formed by debranched starch than that of amylose. Compared to debranched waxy-maize starch, the relative crystallinity of the complexes increased 2–3 times and was positively related to the carbon chain length of the fatty acids. The RS contents of complexes of long-chain fatty acids were also higher than those of short-chain fatty acids, increasing from 33.06% of hexanoic acid to 47.43% of stearic acid [177]. However, some different trends were observed. Debranched wheat starch (DWS) was prepared by pullulanase from Klebsiella and formed complexes with four saturated fatty acids (FAs) with different chain lengths, lauric acid, myristic acid, palmitic acid, and stearic acid. In all cases, the FA contents in the complexes increased after debranching, and FA with a long-chain length was

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more able to form complexes. The highest FA content was obtained from the complex of debranched wheat starch and stearic acid. These differences might be attributed to the complex preparation methods, such as reaction time and temperature, and washing twice with 50% (v/v) ethyl alcohol [192]. Aliphatic alcohols are important flavoring substances and have a wide range of applications in many fields. Recently, it was first reported that debranched starch could form V-type complexes with six aliphatic alcohols, n-hexanol, n-octanol, n-decanol, lauryl alcohol, tetradecyl alcohol, and cetanol. With the increase in the carbon chain length of aliphatic alcohols, the peak melting temperature and relative crystallinity of the complexes gradually increased, but the complexing index value decreased with a linear correlation (R2 = 0.93) [172]. In addition to fatty acids and aliphatic alcohols, starch also formed complexes with other guest molecules. Starch-ascorbyl palmitate (AP) inclusion complexes were prepared, and pullulanase debranching of high-amylose maize starch (HAMS) facilitated formation as the debranching time increased from 0 to 8 h. The loading efficiency reached a maximum of 71.06% when debranching for 8 h. The enthalpy change and crystallinity increased, and the retrogradation of starch was also restricted. However, excessive debranching (16 h) produced excessively short chains and mobility and inhibited complexation. Starch digestion was more restrained (the RS content from 29.37% to 44.59%) with increasing debranching level. The stability of AP in complexes was also enhanced against light, heat, and oxidation [193]. Debranched starch/phosphatidylcholine inclusion complexes were prepared by hydrophobic interactions between the alkyl chain of phosphatidylcholine and the debranched starch helix cavity. Under the optimal conditions, the phosphatidylcholine payload and inclusion rate prepared were 106 mg/g and 84.8%, respectively. Complexation with debranched starch significantly improved the stability of phosphatidylcholine, and the encapsulated phosphatidylcholine was gradually released with pancreatin treatment [173]. A novel emulsion was developed to encapsulate curcumin using debranched starch with a curcumin encapsulation efficiency of 71.11% and a loading rate of 12.07%. The prepared emulsions showed better stability and solubility of curcumin [194]. Similarly, debranched starch nanoparticles (DBS-NPs) were loaded with curcumin, and the maximum encapsulation efficiency reached 92.49%. DBS-NPs significantly enhanced the antioxidant and antibacterial activities and antioxidant stability of curcumin [195]. In addition, tea polyphenol (TPP)-loaded debranched corn starch-xanthan gum-based microcapsules were obtained for the sustained release of TPP to improve its bioavailability and halflife. The encapsulation efficiency was above 80%, and microcapsules with a 34% debranching degree presented suitable preparation performance, exhibiting 30% and 80% release in simulated gastric and intestinal fluids, respectively [196].

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Applications of Starch Debranching Enzymes in Starch Processing

7.7.2

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Production of Starch Complexes in Combination with Debranching Enzymes and Other Treatments

Three sources, potato, common corn, and high-amylose corn starches (Hylon VII), were used for complexation with stearic acid by debranching alone and in combination with an additional β-amylase treatment. Within the debranched starch complexes, the amount of stearic acid in the Hylon VII starch complex was highest, followed by potato starch and common corn starch. An additional β-amylase treatment significantly improved the complexation yield of potato and common corn starch because more suitable starch chains were produced for complexation with stearic acid. However, the Hylon VII starch complexes slightly decreased. The amount of stearic acid in the debranched and β-amylase-treated potato starch was highest, 55.9 mg/g starch complex. The results of X-ray diffraction showed a mixture of B- and V-type patterns in all starch–stearic acid complexes and a more V-type pattern in the debranched and β-amylase-treated starch complexes [197]. Nevertheless, the number of branched chains that can be debranched to take part in the formation of the V-amylose complex is limited in native starch. Maltogenic amylase (MAL) could hydrolyze the α-1,4 glycosidic bonds and transglycosidation to form new α-1,6 linkages, thereby producing amylopectin clusters with higher branch density, which was further debranched by pullulanase (PUL) to generate more linear starch chains with a chain length suitable for the formation of complex with lauric acid (Lau). The physicochemical properties of the starch–Lau complex were found to be significantly influenced by the level of MAL. Crystalline complexes were mainly present in the dual-enzyme-modified samples with MAL levels between 4 and 12 U/g, while amorphous complexes were mainly present in the untreated sample and PUL- and MAL (16 U/g)–PUL–starch–Lau complex. Crystalline complexes are more resistant to digestive enzymes than amorphous complexes [179]. In addition, physical treatments have also been used for the preparation of these complexes [198, 199]. Yixin Zheng et al. used pullulanase pretreatment in combination with ultrasound-microwave synergistic processing to produce lotus seed starch–glycerin monostearin complexes. Pullulanase pretreatment could form more amylose single helices and improve the complex index of the inclusion complexes. The structural properties of lotus seed starch were also determined. Moreover, pullulanase debranching contributed to an increase in slowly digestible starch content [175].

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