Production, Isolation and Purification of Industrial ...

Current Developments in Biotechnology and Bioengineering

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Current Developments in Biotechnology and Bioengineering Production, Isolation and Purification of Industrial Products

Edited by

Ashok Pandey, Sangeeta Negi, Carlos Ricardo Soccol

AMSTERDAM l BOSTON l HEIDELBERG l LONDON l NEW YORK PARIS l SAN DIEGO l SAN FRANCISCO l SINGAPORE l SYDNEY

l l

OXFORD TOKYO

Elsevier Radarweg 29, PO Box 211, 1000 AE Amsterdam, Netherlands The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States Copyright © 2017 Elsevier B.V. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-0-444-63662-1 For information on all Elsevier publications visit our website at https://www.elsevier.com/

Publisher: John Fedor Acquisition Editor: Kostas Marinakis Editorial Project Manager: Anneka Hess Production Project Manager: Vijayaraj Purushothaman Designer: Greg Harris Typeset by TNQ Books and Journals

Contents List of Contributors

xxi

About the Editors

xxvii

Preface

xxxi

Part 1 Industrial and Therapeutic Enzymes

1

1. a-Amylases

3

R. Sindhu, P. Binod, A. Pandey 1.1 Introduction

3

1.2 Sources of a-Amylase

5

1.3 Production of a-Amylase

6

1.4 Assay of a-Amylases

11

1.5 a-Amylase Inhibitors

11

1.6 Strain Improvement

11

1.7 Purification and Characterization of a-Amylases

15

1.8 Applications of a-Amylase

17

1.9 Conclusion and Perspectives

19

References

19

2. Amylolytic Enzymes: Glucoamylases

25

S. Negi, K. Vibha 2.1 Introduction

25

2.2 Sources of Glucoamylase

26

2.3 Glucoamylase Production

28

2.4 Purification and Characterization

31

2.5 Enzyme Assay

36

v

vi

Contents


37

2.7 Commercially Available Glucoamylases

38

2.8 Conclusion and Perspective

40

References

41

3. Pectinolytic Enzymes

47

Hećtor A. Ruiz, Rosa M. Rodrı´guez-Jasso, Ayerim HernandezAlmanza, Juan C. Contreras-Esquivel, Cristo´bal N. Aguilar 3.1 Introduction

47

3.2 Pectic Substances

48

3.3 Pectinase Classification

48

3.4 Pectinase Assays

49

3.5 Pectinase Production Processes

51

3.6 Downstream and Purification Methods

58

3.7 Technoeconomic Analysis of Pectinase Production

61

3.8 Conclusions and Perspectives

63

References

64

4. Cellulases

73

Reeta R. Singhania, M. Adsul, A. Pandey, A.K. Patel 4.1 Introduction

73

4.2 Sources

74

4.3 The Cellulase System

76

4.4 Regulation of Cellulase Expression

79

4.5 Research on Bioprocesses for Improved Cellulase Production

80


85

4.7 Cellulase Global Market

89

4.8 Protocol for Assay of Filter Paper Activity

91

Contents

4.9 Challenges for Enzymatic Biomass Conversion

vii

93

4.10 Future Perspectives

94

4.11 Conclusion

96

References

97

5. Industrial Enzymes: b-Glucosidases

103

Reeta R. Singhania, A.K. Patel, R. Saini, A. Pandey 5.1 Introduction

103

5.2 Classification of b-Glucosidases

104

5.3 Mechanism of Action

105

5.4 Sources of b-Glucosidases

107

5.5 Production of b-Glucosidases

107

5.6 Assay of b-Glucosidases

111


112

5.8 Applications of b-Glucosidases

116


119

References

120

6. Industrial Enzymes: Xylanases

127

L. Thomas, A. Joseph, Reeta R. Singhania, A.K. Patel, A. Pandey 6.1 Introduction

127

6.2 Sources of Xylanases

128

6.3 Production of Xylanases

129

6.4 Purification and Characterization of Xylanases

132

6.5 Xylanase Assays

134


139


140

References

141

viii Contents

7. Proteolytic Enzymes

149

A. Dhillon, K. Sharma, V. Rajulapati, A. Goyal 7.1 Introduction

149

7.2 Sources of Proteolytic Enzymes

150

7.3 Production of Proteolytic Enzymes

152

7.4 Industrial Production Scenario

155

7.5 Purification and Characterization of Proteolytic Enzymes

156

7.6 Assay of Proteolytic Enzymes

158

7.7 Properties of Proteolytic Enzymes

159


160


164

References

165

8. Lipolytic Enzymes

175

R. Gaur, R. Hemamalini, S.K. Khare 8.1 Introduction

175

8.2 Microbial Sources of Enzymes

177

8.3 Production of Enzyme

179

8.4 Purification and Characterization of Lipases

183

8.5 Studies on Pseudomonas aeruginosa Lipase Genes

188

8.6 Assay of Enzyme Lipolytic Activity

189


190

References

190

9. Laccases

199

L.R.C. Guimara˜es, A.L. Woiciechowski, S.G. Karp, J.D. Coral, A. Zandona´ Filho, C.R. Soccol 9.1 Introduction

199

9.2 Possible Substrates for Laccases

199

Contents

ix

9.3 Laccase Action Mechanism

203

9.4 The Ideal Physicochemical Conditions for Laccases

205

9.5 Sources of Laccases

206

9.6 Laccase Applications

207

9.7 Future Trends in Uses and Applications

210


211

References

211

10. Peroxidases

217

J.D.C. Medina, A.L. Woiciechowski, L.R.C. Guimara˜es, S.G. Karp, C.R. Soccol 10.1 Introduction

217

10.2 Lignin Structure

218

10.3 Peroxidases

219

10.4 Lignin Peroxidase

220

10.5 Sources of Peroxidases

221

10.6 Enzymes as Biocatalysts

224

10.7 Peroxidase Applications

225

10.8 Trends and Future of Peroxidases

228


228

References

229

11. Therapeutic Enzymes: L-Glutaminase

233

N. Vijayan, T.S. Swapna, M. Haridas, A. Sabu 11.1 Introduction

233

11.2 Isozymes of Glutaminase

233

11.3 Glutaminase as a Therapeutic Agent

234

11.4 Glutaminase in the Food Industry

234

11.5 Sources of L-Glutaminases

235

11.6 Production of L-Glutaminase

237

x Contents

11.7 Enzyme Characteristics

241

11.8 Glutaminase Assay

243


243


244

References

244

12. Therapeutic Enzymes: L-Asparaginases

249

J. Vidya, S. Sajitha, M.V. Ushasree, P. Binod, A. Pandey 12.1 Introduction

249

12.2 Sources of Asparaginases

250

12.3 Production of L-Asparaginase

251

12.4 Purification and Characterization of L-Asparaginases

254

12.5 Assays for L-Asparaginase

255


256

12.7 Commercially Available L-Asparaginases

259

12.8 Carriers of Asparaginase for Better Delivery Methods

259

12.9 Conclusions and Future Perspectives

260

References

260

13. Industrial and Therapeutic Enzymes: Penicillin Acylase

267

A. Illanes, P. Valencia 13.1 Introduction: Industrial Applications of Penicillin Acylase

267

13.2 Sources of Penicillin Acylase

271

13.3 Improvement of Penicillin Acylase-Producing Strains

274

13.4 Improvement of Penicillin Acylase Catalysts for Industrial Use

277

13.5 Production of Penicillin Acylase

278

13.6 Purification of Penicillin Acylase

280

13.7 Characterization of Penicillin Acylase

286

Contents

xi

13.8 Assays for Penicillin Acylase Activity

288


291

References

291

Other Enzymes

307

14. Other Enzymes: Phytases

309

M.V. Ushasree, J. Vidya, A. Pandey 14.1 Introduction

309

14.2 Phytase Sources

310

14.3 Phytase Market and Production

314


318

14.5 Phytase Assay

320


321

14.7 Summary and Conclusions

323

References

324

15. Chitinases

335

N. Karthik, P. Binod, A. Pandey 15.1 Introduction

335

15.2 Sources of Chitinases

337

15.3 Production of Microbial Chitinases

339

15.4 Assay of Chitinases

345


348

15.6 Purification and Characterization of Chitinases

350

15.7 Biocontrol Properties of Chitinases

355


357

Acknowledgments

358

References

358

xii

Contents

16. a-Galactosidases

369

G.S. Anisha 16.1 Introduction

369

16.2 Substrates for a-Galactosidases

370

16.3 Sources of a-Galactosidases

373

16.4 Production of a-Galactosidases

374


376

16.6 Purification and Characterization of a-Galactosidases

378


386


387

References

387

17. b-Galactosidases

395

G.S. Anisha 17.1 Introduction

395

17.2 Assay of b-Galactosidase Activity

396

17.3 Sources of b-Galactosidase

396

17.4 Production of b-Galactosidase

398

17.5 Extraction of b-Galactosidase

401


402


407

17.8 Industrial Scenario

412


414

References

415

18. Inulinases

423

R.S. Singh, R.P. Singh 18.1 Introduction

423

18.2 Inulinase Production

424

18.3 Purification and Characterization of Inulinases

429

18.4 Inulinase Assay

433

Contents

xiii


433

18.6 Applications of Inulinases

434


436

References

437

19. Keratinases

447

D. Kothari, A. Rani, A. Goyal 19.1 Introduction

447

19.2 Sources

448

19.3 Production of Keratinase

450


452

19.5 Purification of Keratinase

453

19.6 Biochemical Characterization of Keratinases

455

19.7 Assays of Keratinase

457

19.8 Strain Improvement Techniques

458


461

References

461

20. Tannases

471

M.L. Cha´vez Gonza´lez, J. Buenrostro-Figueroa, L.V. Rodrı´guez Durań, P.A. Za´rate, R. Rodrı´guez, Rosa M. Rodrı´guez-Jasso, Hećtor A. Ruiz, Cristo´bal N. Aguilar 20.1 Introduction

471

20.2 Chemical Structure of Tannins

472

20.3 Methods for Tannases Assay

473

20.4 Production of Tannases

474

20.5 Downstream Processing

480

20.6 Scientific and Technological Perspectives

483

20.7 Conclusions

483

References

483

xiv

Contents

21. Microbial Aminopeptidases

491

A. Nandan, K.M. Nampoothiri 21.1 Introduction

491

21.2 Classification and Nomenclature of Aminopeptidases

492

21.3 Catalytic Mechanism of Aminopeptidases

493

21.4 Localization of Aminopeptidase

495

21.5 Physiological Role of Bacterial Aminopeptidases

496

21.6 Fermentative Production of Aminopeptidases

498

21.7 Commercial Aminopeptidases

499

21.8 Industrial Applications of Aminopeptidases

500


502

References

503

22. Nattokinases

509

M.G.B. Pagnoncelli, M.J. Fernandes, C. Rodrigues, C.R. Soccol 22.1 Introduction

509

22.2 Fibrinolytic Enzymes

511

22.3 Nattokinase Production

513

22.4 In Vitro and In Vivo Tests

517


519

22.6 Enzyme Assay

520


521

References

522

23. Polysaccharide Lyases

527

S. Chakraborty, A. Rani, A. Dhillon, A. Goyal 23.1 Introduction

527

23.2 Sources of Polysaccharide Lyase

528

23.3 Production of Polysaccharide Lyases

530

23.4 Enzyme Assay

530

Contents

xv

23.5 Industrial Scenario

531


531

23.7 Biochemical Characterization of Polysaccharide Lyase

532


533


535

References

536

Part 2 Organic Acids

541

24. Production and Application of Lactic Acid

543

C. Rodrigues, L.P.S. Vandenberghe, A.L. Woiciechowski, J. de Oliveira, L.A.J. Letti, C.R. Soccol 24.1 Introduction

543

24.2 Production of Lactic Acid

544

24.3 Metabolic Pathways of Lactic Acid Synthesis

547

24.4 Recovery Processes

550

24.5 Demand and Products

551


552

References

553

25. Production and Application of Citric Acid

557

L.P.S. Vandenberghe, C. Rodrigues, J.C. de Carvalho, A.B.P. Medeiros, C.R. Soccol 25.1 Introduction

557

25.2 Production

558

25.3 Metabolic Pathways

562

25.4 Factors Affecting Citric Acid Production

564

25.5 Citric Acid Recovery Processes

568

25.6 Demand and Products

570


571

References

573

xvi

Contents

26. Gluconic Acid

577

S. Ramachandran, S. Nair, C. Larroche, A. Pandey 26.1 Introduction

577

26.2 History

578

26.3 Properties

579

26.4 Applications

581

26.5 Sources of Gluconic Acid

582

26.6 Glucose Oxidase

582

26.7 Production of Gluconic Acid

587

26.8 Downstream Processing

594

References

595

27. Production and Applications of Succinic Acid

601

R.K. Saxena, S. Saran, J. Isar, R. Kaushik 27.1 Introduction

601

27.2 Market and Companies Involved in Succinic Acid Production

603

27.3 Microorganisms Involved in the Production of Succinic Acid

605

27.4 Synthesis and Enzyme Regulation of Succinic Acid

607

27.5 Estimation of Enzymes Involved in Succinic Acid Production

610

27.6 Enzymatic Regulation of Succinic Acid Production

612

27.7 Production and Regulation of Succinic Acid

613

27.8 Recovery Systems for Succinic Acid

616

27.9 Applications and Uses of Succinic Acid

621


622

Acknowledgment

624

References

624

Contents

xvii

Part 3 Biopolymers and Other Products

631

28. Production and Application of Polylactides

633

J. de Oliveira, L.P.S. Vandenberghe, S.F. Zawadzki, C. Rodrigues, J.C. de Carvalho, C.R. Soccol 28.1 Introduction

633

28.2 Properties of Polylactide

634

28.3 Synthesis of Polylactide

635

28.4 Applications of Polylactide

642

28.5 Polylactide Global Market

646


646

References

647

29. Production of Polyhydroxyalkanoates

655

D. Tan, J. Yin, G.-Q. Chen 29.1 Introduction to Polyhydroxyalkanoates

655

29.2 Applications of Polyhydroxyalkanoates

661

29.3 Laboratory Production of Polyhydroxyalkanoates

664

29.4 Industrial Production of Polyhydroxyalkanoates

675

29.5 Open and Continuous Production of Polyhydroxyalkanoates by Halophilic Microorganisms

680

29.6 Extraction and Purification of Polyhydroxyalkanoates

682

29.7 Conclusions

683

References

684

30. Production and Application of Poly-g-glutamic Acid

693

Q. Wang, X. Wei, S. Chen 30.1 Introduction

693

30.2 Poly-g-glutamic Acid Biosynthesis by Microorganisms

693

xviii

Contents

30.3 Fermentative Production of Poly-g-glutamic Acid

698

30.4 Purification and Characterization of Poly-g-glutamic Acid

703

30.5 Applications of Poly-g-glutamic Acid

706


711

References

712

31. Production and Applications of 1,3-Propanediol

719

N. Vivek, A. Pandey, P. Binod 31.1 Introduction

719

31.2 Properties of 1,3-Propanediol

720

31.3 Substrates and Cosubstrates for 1,3-Propanediol Production

720

31.4 Chemical Synthesis of 1,3-Propanediol

720

31.5 Biological Production of 1,3-Propanediol

721

31.6 Downstream Process

731

31.7 Problems in Biotransformation

732

31.8 Applications of 1,3-Propandiol

733


733

Acknowledgments

734

References

734

32. Biodegradation of Biopolymers

739

N.R. Nair, V.C. Sekhar, K.M. Nampoothiri, A. Pandey 32.1 Introduction

739

32.2 Types of Biopolymers

740

32.3 Mechanisms Involved and Factors Affecting Biodegradability

741

32.4 Biodiversity of Biopolymer-Degrading Microorganisms

742

32.5 Biodegradation of Bio-Based Plastics

742

32.6 Starch-Based Polymers

743

Contents

xix

32.7 Cellulose-Based Polymers

745

32.8 Aliphatic Polyesters of Microbial Origin

745

32.9 Polymers From Bio-Derived Monomers: Polylactic Acid

747

32.10 Polymers From Petrochemical Products

748

32.11 Conclusion and Future Perspectives

751

References

751

33. Production of Fungal Spores for Biological Control

757

F. Miranda-Hernańdez, A. Angel-Cuapio, O. Loera-Corral 33.1 Introduction

757

33.2 Insect Pests and Their Effects on Agricultural Production

757

33.3 Chemical Pesticides

758

33.4 Current Alternatives for Pest Control

758

33.5 Biological Control: Concept and Agents

759

33.6 Entomopathogenic Fungi

760

33.7 Substrates Used to Produce Entomopathogenic Fungi

764

33.8 Production Types

767

33.9 Quality Evaluation

770

33.10 Unsolved Problems and Perspectives

773

References

773

Part 4 Products Isolation and Purification

781

34. Approaches for the Isolation and Purification of Fermentation Products

783

J.C. de Carvalho, A.B.P. Medeiros, L.P.S. Vandenberghe, A.I. Magalha˜es, C.R. Soccol 34.1 Introduction

783

34.2 From Molecule Properties to Process Selection

786

xx

Contents

34.3 Separation Principles

786

34.4 Process Development for Bioseparations

788

34.5 Typical Separation Steps for Selected Classes of Biomolecules

789

34.6 Alternative Separations

789

34.7 Sizing Guide for Operations

791

34.8 Niche Operations and Single-Use Systems

800

34.9 Selected Guidelines for Process Development

802


803

References

804

35. Cell Disruption and Isolation of Intracellular Products

807

J.C. de Carvalho, A.B.P. Medeiros, L.A.J. Letti, P.C.S. Kirnev, C.R. Soccol 35.1 Introduction

807

35.2 Cell Disruption Methods

808

35.3 Choosing Cell-Disruption Equipment

817

35.4 Selected Microorganisms and Disruption Conditions

817

35.5 Product Isolation Rationale

819


820

Acknowledgments

820

References

820

Index

823

List of Contributors M. Adsul DBT-IOC Centre for Advanced Bioenergy Research, IndianOil Corporation Limited

Cristóbal N. Aguilar

Food Research Department, School Autonomous University of Coahuila, Saltillo, Coahuila, México

A. Angel-Cuapio

of

Chemistry,

Universidad Autónoma Metropolitana-Iztapalapa, Mexico City,

DF, Mexico

G.S. Anisha

Government College for Women, Trivandrum, Kerala, India

P. Binod CSIR-National Institute for Interdisciplinary Science and Technology (NIIST), Trivandrum, India J. Buenrostro-Figueroa

Department of Biotechnology, Division of Health and Biological Sciences, Metropolitan Autonomous University, Iztapalapa, México

S. Chakraborty

Indian Institute of Technology Guwahati, Guwahati, Assam, India

M.L. Chávez González Food Research Department, School of Chemistry, Autonomous University of Coahuila, Saltillo, Coahuila, México

G.-Q. Chen S. Chen

Tsinghua University, Beijing, China

Hubei University, Wuhan, PR China

Juan C. Contreras-Esquivel

Food Research Department, School of Chemistry, Autonomous University of Coahuila, Saltillo, Coahuila, México

J.D. Coral

Bioprocess Engineering and Biotechnology Department, Federal University of Paraná (UFPR), Curitiba, PR, Brazil

J.C. de Carvalho


xxi

xxii

List of Contributors

J. de Oliveira


A. Dhillon


M.J. Fernandes Bioprocess Engineering and Biotechnology Department, Federal University of Paraná (UFPR), Curitiba, PR, Brazil

R. Gaur

Indian Oil Corporation Limited, R&D Centre, Faridabad, India

A. Goyal


L.R.C. Guimarães


M. Haridas

Kannur University, Kannur, India

R. Hemamalini

Indian Institute of Technology Delhi, New Delhi, India

Ayerim Hernandez-Almanza


A. Illanes J. Isar

Pontificia Universidad Católica de Valparaíso, Valparaíso, Chile

University of Delhi South Campus, New Delhi, India

A. Joseph


S.G. Karp


N. Karthik

CSIR-National Institute for Interdisciplinary Science and Technology (NIIST), Trivandrum, India

R. Kaushik


S.K. Khare

Indian Institute of Technology Delhi, New Delhi, India

P.C.S. Kirnev


D. Kothari



C. Larroche

xxiii

Blaise Pascal University, Aubière Cedex, France

L.A.J. Letti Bioprocess Engineering and Biotechnology Department, Federal University of Paraná (UFPR), Curitiba, PR, Brazil

O. Loera-Corral

Universidad Autónoma Metropolitana-Iztapalapa, Mexico City, DF,

Mexico

A.I. Magalhães, Jr. Bioprocess Engineering and Biotechnology Department, Federal University of Paraná (UFPR), Curitiba, PR, Brazil

A.B.P. Medeiros


J.D.C. Medina


F. Miranda-Hernández

Universidad Autónoma Metropolitana-Iztapalapa, Mexico

City, DF, Mexico

N.R. Nair


S. Nair

Dow Chemicals GmBH, Dubai, UAE

K.M. Nampoothiri


A. Nandan


S. Negi

Motilal Nehru National Institute of Technology, Allahabad, India

M.G.B. Pagnoncelli

Bioprocess Engineering and Biotechnology Department, Federal University of Paraná (UFPR), Curitiba, PR, Brazil; Federal Technological University of Parana, Dois Vizinhos, Brazil

A. Pandey

Center of Innovative and Applied Bioprocessing, (a national institute under Dept of Biotechnology, Ministry of S&T, Govt of India), Mohali, Punjab, India

A.K. Patel Limited

DBT-IOC Centre for Advanced Bioenergy Research, IndianOil Corporation

xxiv


V. Rajulapati


S. Ramachandran A. Rani

Insight Professional Institute, Dubai, UAE


C. Rodrigues


Rosa M. Rodríguez-Jasso


R. Rodríguez


L.V. Rodríguez Durán

Department of Biotechnology, Division of Health and Biological Sciences, Metropolitan Autonomous University, Iztapalapa, México

Héctor A. Ruiz


A. Sabu


R. Saini

DBT-IOC Centre for Advanced Bioenergy Research, IndianOil Corporation

Limited

S. Sajitha CSIR-National Institute for Interdisciplinary Science and Technology (NIIST), Trivandrum, India S. Saran


R.K. Saxena University of Delhi South Campus, New Delhi, India V.C. Sekhar


K. Sharma


R. Sindhu


R.P. Singh

Punjabi University, Patiala, Punjab, India


xxv

R.S. Singh Punjabi University, Patiala, Punjab, India Reeta R. Singhania

DBT-IOC Centre for Advanced Bioenergy Research, IndianOil

Corporation Limited

C.R. Soccol


T.S. Swapna Government Victoria College, Palakkad, India D. Tan Xían Jiaotong University, Xían, China L. Thomas


M.V. Ushasree


P. Valencia

Universidad Técnica Federico Santa María, Valparaíso, Chile

L.P.S. Vandenberghe


K. Vibha

Motilal Nehru National Institute of Technology, Allahabad, India

J. Vidya


N. Vijayan


N. Vivek


Q. Wang X. Wei



A.L. Woiciechowski Bioprocess Engineering and Biotechnology Department, Federal University of Paraná (UFPR), Curitiba, PR, Brazil

J. Yin

Tsinghua University, Beijing, China

xxvi


A. Zandoná Filho


P.A. Zárate


S.F. Zawadzki


About the Editors Ashok Pandey Professor Ashok Pandey is Eminent Scientist at the Center of Innovative and Applied Bioprocessing, Mohali (a national institute under the Department of Biotechnology, Ministry of Science and Technology, Government of India), and former chief scientist and head of the Biotechnology Division at the CSIR’s National Institute for Interdisciplinary Science and Technology at Trivandrum. He is an adjunct professor at Mar Athanasios College for Advanced Studies Thiruvalla, Kerala, and at Kalasalingam University, Krishnan Koil, Tamil Nadu. His major research interests are in the areas of microbial, enzyme, and bioprocess technology, which span various programs, including biomass to fuels and chemicals, probiotics and nutraceuticals, industrial enzymes, solid-state fermentation, etc. He has more than 1100 publications and communications, which include 16 patents, 50+ books, 125 book chapters, and 425 original and review papers, with an h index of 75 and more than 23,500 citations (Google Scholar). He has transferred several technologies to industries and has been an industrial consultant for about a dozen projects for Indian and international industries. Professor Pandey is the recipient of many national and international awards and fellowships, which include Elected Member of the European Academy of Sciences and Arts, Germany; Fellow of the International Society for Energy, Environment and Sustainability; Fellow of the National Academy of Science (India); Fellow of the Biotech Research Society, India; Fellow of the International Organization of Biotechnology and Bioengineering; Fellow of the Association of Microbiologists of India; honorary doctorate degree from the Universite´ Blaise Pascal, France; Thomson Scientific India Citation Laureate Award, United States; Lupin Visiting Fellowship; Visiting Professor at the Universite´ Blaise Pascal, France, the Federal University of Parana, Brazil, and the Ećole Polytechnique Fe´de´rale de Lausanne, Switzerland; Best Scientific Work Achievement Award, Government of Cuba; UNESCO Professor; Raman Research Fellowship Award, CSIR; GBF, Germany, and CNRS, France fellowships; Young Scientist Award; and others. He was chairman of the International Society of Food, Agriculture and Environment, Finland (Food & Health) during 2003e04. He is the Founder President of the Biotech xxvii

xxviii

About the Editors

Research Society, India (www.brsi.in); International Coordinator of the International Forum on Industrial Bioprocesses, France (www.ifibiop.org); chairman of the International Society for Energy, Environment & Sustainability (www.isees.org); and vice president of the All India Biotech Association (www.aibaonline.com). Professor Pandey is editor-in-chief of Bioresource Technology, Honorary Executive Advisor of the Journal of Water Sustainability and Journal of Energy and Environmental Sustainability, subject editor of the Proceedings of the National Academy of Sciences (India), and editorial board member of several international and Indian journals, and also a member of several national and international committees. Sangeeta Negi Dr. Sangeeta Negi is an assistant professor in the Department of Biotechnology at the Motilal Nehru National Institute of Technology, India. She has a First Class Master’s degree in biochemistry and a PhD in biotechnology from the Indian Institute of Technology, Kharagpur. She has also worked as an academic guest at the Biological Engineering Department, Polytech Clermont-Ferrand; the Universite´ Blaise Pascal, France; and the Bioenergy and Energy Planning Research Group, Swiss Federal Institute of Technology, Lausanne, Switzerland. Dr. Negi’s current research interests are in the areas of biofuels, industrial enzymes, and bioremediation. She is an editorial board member of the Journal of Waste Conversion, Bioproducts and Biotechnology and the Journal of Environmental Science and Sustainability. She has been awarded as Outstanding Reviewer by Elsevier and has won the Young Scientist Award by DST at the National Seminar on Biological and Alternative Energies Present and Future, organized by Andhra University, Visakhapatnam, in 2009. She has also won the Best Poster Award at the International Congress on Bioprocesses in Food Industries (2008) at Hyderabad. Dr. Negi has contributed to nearly 70 publications, including review articles, original papers, and conference communications.

About the Editors

xxix

Carlos Ricardo Soccol Professor Carlos Ricardo Soccol is the research group leader of the Department of Bioprocesses Engineering and Biotechnology at the Federal University of Parana´ (UFPR), Brazil, with 20 years of experience in biotechnological research and development of bioprocesses with industrial application. He graduated with a BSc in chemical engineering (UFPR, 1979), Master’s in food technology (UFPR, 1986), and PhD in Geńie Enzymatique, Microbiologie et Bioconversion (Universite´ de Technologie de Compie`gne, France, 1992). He did his postdoctoral work at the Institut ORSTOM/IRD (Montpellier, 1994 and 1997) and at the Universite´ de Provence et de la Me´diterraneé (Marseille, ´ cole d’Ingeńieurs Supe´riure of Luminy, 2000). He is an HDR Professor at the E MarseilleeFrance. He has experience in the areas of science and food technology, with emphasis on agro-industrial and agro-alimentary biotechnology, acting in the following areas: bioprocess engineering and solid-state fermentation, submerged fermentation, bioseparations, industrial bioprocesses, enzyme technology, tissue culture, bioindustrial projects, and bio-production. He is currently the Coordinator of Master BIODEV-UNESCO, associate editor of five international journals, and editor-in-chief of the journal Brazilian Archives of Biology and Technology. Professor Soccol has received several national and international awards, which include the Science and Technology Award of the Government of Parana´ (1996); Scopus/Elsevier Award (2009); Dr. Honoris Causa, Universite´ Blaise Pascal, France (2010); Outstanding Scientist at the 5th International Conference on Industrial Bioprocesses, Taipei, Taiwan (2012); and Elected Titular Member of the Brazilian Academy of Sciences (2014). He is a technical and scientific consultant for several companies, agencies, and scientific journals in Brazil and abroad. He has supervised and mentored 96 Master of Science students, 48 PhD students, and 14 postdoctoral students. He has 995 publications and communications, which include 17 books, 107 book chapters, 270 original research papers, and 557 research communications in international and national conferences and has registered 44 patents. His research articles as of this writing have been cited (Scopus database) 5600 times with an h index of 36.


Preface This book is a part of the comprehensive series Current Developments in Biotechnology and Bioengineering, comprising nine volumes (Editor-in-chief: Ashok Pandey), and deals with the production, isolation, and purification of industrial products produced by biotechnological processes. This book covers recent technological advances of a great number of biotechnological products and is divided into four different parts: Production of Industrial and Therapeutic Enzymes, Organic Acids, Biopolymers and Other Products, and Products Isolation and Purification. Part 1 is devoted to the production of industrial and therapeutic enzymes. The first chapter describes the current and future trends of production, application, and strain improvement of a-amylases, one of the most important enzymes used in industry. a-Amylases find application in several industrial processes, such as starch liquefaction, desizing of textiles, detergents, baking, bioethanol production, etc. Glucoamylase is another enzyme extensively used in the food and fermentation industries, mainly for the saccharification of starch, brewing, and production of high-fructose syrup, which are discussed in Chapter 2. Cellulases, b-glucosidases, and xylanases are the second most used enzymes in industry by sales volume, with an increasing demand since 1995 in several industrial applications, comprising detergents and textiles, animal feed, food, paper, and biofuels. These enzymes are discussed in Chapters 4, 5, and 6 of this book. Chapter 7 discusses proteolytic enzymes, also known as “proteases,” which are used to cleave the peptide bonds connecting two amino acids. They are produced mainly by microorganisms and have great commercial value, being used in food, dairy, detergents, and leather processing. Lipolytic enzymes are hydrolases comprising 15 families of lipases, as shown in Chapter 8 of this book through a study of the industrial applications and other important aspects of these enzymes. The purpose of Chapters 9 and 10 is to present an overview of laccases and peroxidases, covering their production and use in the pretreatment of lignocellulosic biomass and biopulping, and also projecting new perspectives on improving such processes and products using these enzymes. Sources of production, strategies, characteristics, applications, and industrial importance of therapeutic enzymes, such as L-glutaminase, L-asparaginase, and penicillin acylase, are presented and discussed in Chapters 11, 12, and 13. Other enzymes, such as phytases, chitinases, keratinases, tannases, aminopeptidases, nattokinases, and polysaccharide lyases, are reviewed in Chapters 14 to 23, covering recent advances, production methods, potential applications, and the global market. The second part of the book is dedicated to organic acids. In Chapters 24 and 25, lactic acid and citric acid production, synthesis (covering factors that affect biochemical pathways), and recovery are addressed. Chapter 26 reviews the microbial production of gluconic acid, properties of glucose oxidase, production, recovery, and applications. Succinic acid is an important platform molecule, used as an intermediate in the production of numerous everyday products, among which are pharmaceuticals and adhesives, representing a total immediate addressable market of more than $7.2 billion. Chapter 27 presents an analysis of the current market, biological-based production processes, enzymatic regulation, and recovery systems of succinic acid.

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Preface

Part 3 discusses polymer production and other products. Polylactide (PLA), derived from lactic acid, a biodegradable polyester, has applications in packaging, textiles, and the biomedical and pharmaceutical industries. Chapter 28 reviews the properties and applications of PLA, focusing on recent technologies and improvement of production techniques. Polyhydroxyalkanoates (PHAs), a family of environmentally friendly polyesters that can be synthesized by a wide range of microorganisms as carbon and energy reserves, have been considered an alternative to petroleum-based chemicals. The composition and structural diversity of PHAs have led to various properties and endless applications to form a PHA value chain. Chapter 29 briefly introduces their production and application, highlighting the laboratory production by the microbial strains developed using genetic and/or metabolic engineering or synthetic biology techniques. Industrial production, recent technologies, and improvement of PHA production are also discussed. Poly-g-glutamic acid (g-PGA) is a natural polymer, synthesized by various strains of Bacillus spp., that is used in food, cosmetics, agriculture, and the wastewater industry. Chapter 30 provides updated information on the biosynthesis, fermentation, purification, and application of g-PGA. In Chapter 31, recent developments in the biological production of 1,3-propanediol by various natural and genetically engineered microorganisms, nonnative 1,3-propanediol producers, as well as mixed cultures, are discussed. Important aspects of downstream processing and various methods and steps involved in the extraction and purification of 1,3-propanediol from the fermentation broth are also covered in this chapter. The production of petroleum-based plastics is a challenging environmental problem, causing the production and consumption of biodegradable plastics to receive considerable attention nowadays. Chapter 32 provides an overview of the degradation mechanisms of biodegradable polymers, with particular emphasis on the main parameters affecting the degradation of these polymeric biomaterials. In Chapter 33 the potential of biological control is presented and discussed as a promising alternative to chemical pesticides. The final two chapters of this book, Chapters 34 and 35, present the most relevant downstream processes to extract, isolate, purify, and refine fermentation products. We are confident that this book will be profitable to students, professors, researchers, and professionals interested in studying biotechnology and bioengineering. We thank Dr. Kostas Marinakis, Book Acquisition Editor; Ms. Anneka Hess; and entire production team at Elsevier for their help and support in bringing out this volume. Editors Ashok Pandey Sangeeta Negi Carlos Ricardo Soccol

PART

1 Industrial and Therapeutic Enzymes


a-Amylases

1 R. Sindhu1, *, P. Binod1, A. Pandey2

1

CSIR-NATIONAL INSTITUTE F OR INTERDISCIPLINARY SCIENCE AND TECHNOLOGY (NIIST), TRIVANDRUM, INDIA; 2 CE NTER OF INNOVATIVE AND APPLIED BIOPROCESSING, (A NATIONAL INSTITUTE UNDER DEPT OF BIOTECHNOL OGY, MINISTRY OF S&T, GOVT OF INDIA), MOHALI, PUNJAB , INDIA

1.1 Introduction 1.1.1

Starch

Starch is the major polysaccharide food reserve in nature after cellulose. It serves as an important source of nutrition for other living organisms [1]. It is synthesized in the plastids present in leaves and accumulates as insoluble granules in higher and lower plants. Starch is composed of a large number of glucose units joined by glycosidic bonds. It consists of two types of molecules: amylose and amylopectin. Amylose is a linear, water-insoluble polymer of glucose joined by a-1,4 bonds, whereas amylopectin is a branched, water-soluble polysaccharide with short a-1,4-linked linear chains of 10e60 glucose units and a-1,6-linked side chains with 15e45 glucose units. The levels of amylase and amylopectin vary among different starches. Generally, starch is composed of amylose and amylopectin in the range 25e28% and 72e75%, respectively.

1.1.2

Amylases

Amylases are the enzymes that break down starch, or glycogen. These enzymes are produced by a variety of living organisms, ranging from bacteria to plants to humans. Though amylases are produced by several microorganisms, those produced by fungi and bacteria have dominated applications in the industrial sector [2]. Bacteria and fungi secrete amylases to the outside of their cells to carry out extracellular digestion, which breaks down the insoluble starch, and then the soluble end products (such as glucose or maltose) are absorbed into the cells. Amylases constitute a class of industrial enzymes occupying about 25% of the enzyme market. Because of the increasing demand for these enzymes in various industries, there is enormous interest in developing them with better properties, such as raw starchdegrading amylases suitable for industrial applications, and cost-effective production *

Corresponding Author.

Current Developments in Biotechnology and Bioengineering: Production, Isolation and Purification of Industrial Products http://dx.doi.org/10.1016/B978-0-444-63662-1.00001-4 Copyright © 2017 Elsevier B.V. All rights reserved.

3

4 CURRENT DEVELOPMENTS IN BIOTECHNOLOGY AND BIOENGINEERING

techniques. Although amylases can be derived from several sources, including plants, animals, and microorganisms, microbial enzymes generally meet industrial demands. A large number of microbial amylases are available commercially and they have almost completely replaced the chemical hydrolysis of starch in the starch processing industry [3]. One of the most important advantages of using microbes for the production of amylases is the bulk production capacity and the fact that microbes can be genetically modified to produce enzymes with desired characteristics [4]. These enzymes are of great significance in biotechnology, with various applications ranging from food, fermentation, and textiles to the paper industry. Each application of a-amylase requires unique properties with respect to specificity, stability, and temperature and pH dependence. Modern technologies such as computational packages and online servers are the current tools used in protein sequence analysis and characterization. The physicochemical and structural properties of these proteins are well understood with the use of computational tools. The protein sequence profile, such as number of amino acids and sequence length, and the physicochemical properties of the protein, such as molecular weight, atomic composition, extinction coefficient, aliphatic index, instability index, etc., can be computed by ProtParam, and the secondary structure prediction, sequence similarity, evolutionary relationships, and 3-D structure of various proteins can be computed using the ESyPred3D server [5].

1.1.3

Classification of Amylases

Based on the mechanism of breakdown of starch, the molecules are classified into three types: a-amylase, b-amylase, and amyloglucosidase. a-Amylase reduces the viscosity of starch by breaking down the bonds at random, thereby producing variably sized chains of glucose. b-Amylase enzyme breaks the glucoseeglucose bonds by removing two glucose units at a time, thereby producing maltose. Amyloglucosidase is the enzyme that breaks successive bonds from the nonreducing end of the straight chain, producing glucose. Many microbial amylases usually contain a mixture of these amylases. This chapter focuses only on a-amylases. a-Amylases (EC 3.2.1.1) are starch-degrading enzymes that catalyze the hydrolysis of internal a-1,4-O-glycosidic bonds in the polysaccharides with the retention of the a-anomeric configuration in the products. Most of the a-amylases are metalloenzymes, which require calcium ions (Ca2þ) for their activity, structural integrity, and stability. They belong to family 13 (GH-13) of the glycoside hydrolase group of enzymes [6,7]. Based on the end-product formation a-amylases are classified as saccharifying and liquefying amylases. The saccharifying a-amylases are further classified as maltose forming, maltotetraose forming, maltopentaose forming and maltohexaose forming based on the end products formed [1]. The a-amylase family is the largest family of glycoside hydrolases, transferases, and isomerases, comprising 30 different enzyme specificities. These enzymes are classified into four groups: endoamylases, exoamylases, debranching enzymes, and transferases. Endoamylases are enzymes that cleave internal a-1,4 bonds resulting in a-anomeric

Chapter 1 a-Amylases

5

products. Exoamylases are enzymes that cleave a-1,4, or a-1,6 bonds of the external glucose residues resulting in a- or b-anomeric products. Debranching enzymes are enzymes that hydrolyze a-1,6 bonds leaving linear polysaccharides. Transferases are enzymes that cleave a-1,4 glycosidic bonds of the donor molecule and transfer part of the donor molecule to a glycosidic acceptor, forming a new glycosidic bond [7].

1.2 Sources of a-Amylase a-Amylases are universally distributed throughout the plant, animal, and microbial kingdoms. The enzymes from microbial sources have dominated applications in industrial processes [2]. Though a-amylases have been derived from several microbial sources, including bacteria, fungi, yeast, and actinomycetes, the enzymes produced from bacterial and fungal sources have dominated applications in industrial sectors. Because of their short growth period, their biochemical diversity, and the ease with which enzyme concentrations might be increased by environmental and genetic manipulation, the enzymes from microbial sources generally meet industrial demands.

1.2.1

Plant a-Amylases

Plants store carbon predominantly as starch and the metabolism of starch is essential to all life. Family 1 a-amylases are characterized by having a secretary signal peptide. This plays an important role in the degradation of extracellular starch in cereal grain endosperms. Family 2 a-amylases are characterized by having no predicted targeted peptide and are localized in the cytoplasm. These amylases have been identified from monocotyledons, dicotyledons, and gymnosperms. They become most active when the plastidial starch reserves of leaves are more depleted. They are involved in general stress responses. Family 3 a-amylases are characterized by having a large N-terminal domain, which contains a large predicted chloroplast transit peptide. These enzymes are responsible for degrading plastid-bound starch in storage tissues and leaves [8].

1.2.2

Bacterial a-Amylases

a-Amylases are produced from various bacterial sources, including Bacillus, Brevibacterium, Clostridium, Halomonas, Naxibacter, Nesterenkonia, Paenibacillus, Pseudomonas, Streptomyces sp., etc. Among the bacterial sources, Bacillus sp. is widely used, especially for the production of thermostable a-amylases. Bacillus subtilis, Bacillus stearothermophilus, Bacillus amyloliquefaciens, Bacillus licheniformis, Bacillus acidocaldarius, Bifidobacterium bifidum, and Bifidobacterium acerans are important sources used for a-amylase production [9]. Alkaline and thermotolerant amylases have been reported from Bacillus sp., B. licheniformis, and Bacillus halodurans [10]. Other bacteria producing a-amylase include Anoxybacillus beppuensis [11], Bacillus laterosporus [12], Bacillus acidicola [13], Chryseobacterium taeanense [14], Clostridium sp. [15], Microbacterium foliorum [16], Nesterenkonia sp. [17], Thermococcus sp. [18], Anoxybacillus flavithermus [19] etc.


1.2.3

Fungal a-Amylases

Several fungal species also produce a-amylases, including Acremonium, Aspergillus, Penicillium, Mucor, Neurospora, and Thermomyces sp. Among the fungal sources, the genus Aspergillus has been widely used for the production of a-amylases. Aspergillus niger, Aspergillus flavus, and Aspergillus oryzae are important sources used among the fungal sources [20,21]. Other fungal strains producing a-amylase include Thermomyces lanuginosus [22].

1.3 Production of a-Amylase 1.3.1

Production Methods

To meet the industrial demand, it is essential to develop a low-cost medium for the production of a-amylase. It can be produced by submerged fermentation (SmF) and solid-state fermentation (SSF). The production is affected by a variety of physiological factors, which include pH, temperature, aeration, inoculum concentration, inoculum age, composition of the growth medium, surfactants, carbon source, nitrogen source, etc. [23]. Interactions of these parameters have a significant influence on the production of the enzyme. Generally, SmF is carried out using synthetic media, incorporating medium constituents such as nutrient broth and soluble starch, as well as other components, which are very expensive. Replacement of such constituents by cheaper carbon and nitrogen sources as well as nutrients would benefit the process in cost reduction. Agricultural by-products offer potential benefits in this regard [7]. SSF is defined as the process in which the growth of microorganisms is carried out on solid substrates with negligible free water, or free-flowing water [24]. SSF plays an important role in the production of enzymes. Agro-industrial substrates are considered the best substrates for SSF processes. It is of special interest in those processes in which a crude fermented product may be used directly as an enzyme source. The common substrates used for SSF processes are wheat bran, rice bran, cassava waste, palm oil waste, banana waste, tea waste, coconut oil cake, coir pith, corn cobs, etc. In SSF, it is important to provide optimized water content and to control the water activity of the fermenting substrate. At times, SSF is preferred to SmF because of its simple technique, low capital investment, lower levels of catabolite repression and end-product inhibition, low wastewater output, better product recovery, and high-quality production [25]. Continuous and fed-batch studies are more effective for the production of a-amylase. The study conducted by Lee and Parulekar [26] revealed that the a-amylase production by B. subtilis TN 106 was enhanced when batch cultivation was extended with fed-batch cultivation, and the enzyme activity was 54% higher in a two-stage fed-batch operation compared to a single-stage batch culture. Mishra and Maheswari [27] reported a-amylase from a thermophilic fungus, T. lanuginosus; the enzyme was a dimeric protein with a molecular mass of 42 kDa with optimum pH and temperature of 5.6 and 65 C,


7

respectively. The enzyme produced high levels of maltose from potato starch, suggesting its usefulness in the commercial production of maltose and glucose syrups. The study conducted by Krishna and Chandrasekharan [28] revealed that banana peel could be utilized as a potential substrate for a-amylase production by A. niger. Saxena and Singh [29] screened various agro-industrial residues for amylase production from Bacillus sp. and found mustard oil cake to be the best substrate. The strain produced 5400 U/g of amylase at 1:3 moisture content, 20% inoculum, and an incubation period of 72 h. Yang and Wang [30] reported a-amylase production by Streptomyces rimosus TM 55 using sweet potato residue and peanut meal residue as a substrate. The strain produced 1903 U of a-amylase after 96 h of incubation. Ramachandran et al. [20] used coconut oil cake (COC), a by-product of oil extraction from dried copra, as a substrate for the production of a-amylase from fungi. COC supplemented with 0.5% starch and 1% peptone enhanced a-amylase production by A. oryzae. COC serves as a source of soluble proteins and lipids thus providing essential nutrients for the growth of and enzyme synthesis by the organism. Production of a-amylase by B. amyloliquefaciens under SSF using corn gluten meal (CGM) was reported by Saban et al. [31]. The study revealed that a-amylase production in a medium with CGM was five times higher than that in a medium containing starch and other components. Utilization of CGM as a substrate makes the process economically viable because CGM is a by-product of starch-based industries. Production and optimization of a-amylase from A. oryzae CBS 819 using a by-product of wheat grinding (gruel) as the sole carbon source was done by Kammoun et al. [32]. Various process parameters affecting the production were optimized by adopting a BoxeBehnken design, which increased the enzyme production from 40.1 to 151.1 U/mL. Murthy et al. [33] reported coffee by-products as suitable substrate for the production of a-amylase under SSF. Coffee waste was converted into value-added products by fermentation using Neurospora crassa CFR 308. The optimum conditions for a-amylase production were moisture content of 60%, pH 4.5, incubation temperature of 27 C, particle size of 1 mm, and incubation time of 5 days. Under optimized conditions the strain produced 7084 U/gds of a-amylase. Syed et al. [34] reported extracellular amylase production by Streptomyces gulbargensis DAS 131 by SmF. The highest amylase production was observed when the medium was supplemented with 1% starch. The enzyme was thermotolerant and stable at pH 9.0. Starch and peptone were good sources of carbon and nitrogen. Sharma and Satyanarayana [13] reported enhanced production of acidic high-maltose-forming and Ca2þ-independent a-amylase by B. acidicola; a maximum enzyme titer of 366 IU/L was attained after 36 h of fermentation at pH 4.5, 33 C, with 0.5 vvm aeration. The enzyme titer was 10,100 IU/L in fed-batch fermentation. One of the main advantages of fedbatch fermentation over the batch fermentation is that the concentration of limiting substrate is maintained at low levels, thus avoiding the repressing effect of high substrate concentration and thereby minimizing the accumulation of inhibitory metabolites.


A highly thermostable and calcium-independent a-amylase from A. beppuensis TSSC-1 was reported by Kikani and Singh [11]. This organism produced a monomeric a-amylase with optimal pH and temperature of 7.0 and 55 C, respectively. The key findings of this study were cost-effective purification, high thermostability, and broad pH stability. The enzyme exhibited Ca2þ independence and resistance to chemical denaturation, which could make it suitable for many industrial applications. Another agro-industrial residue, date waste, has also been used as the substrate for the production of a-amylase using yeast, Candida guilliermondii CGL-A10 [35]. Maximum enzyme production was attained in SmF (2056 mmol/L/min). Rajagopalan et al. [15] used sugarcane bagasse hydrolyzate for the production of a-amylase produced by a solventogenic Clostridium sp. BOH3. The strain used starch directly without any pretreatment and produced extracellular amylase (7.15 U/mg protein) and butanol almost equivalent to 90% of the yield equivalent to glucose. Sugarcane bagasse was used by Roohi and Kuddus [16] to produce a cold-active a-amylase from M. foliorum GA2. Maximum enzyme production (6610 U) was observed when fermentation was carried out in a medium containing 40% bagasse, 0.0003 M lactose, at pH 8.0, with incubation temperature of 20 C for 5 day at static conditions. This was the first report on coldactive a-amylase production from M. foliorum GA2. Table 1.1 shows various microorganisms used for the production of a-amylase.

1.3.2

Factors Influencing the Production of a-Amylase

Production of a-amylase by SSF and SmF is affected by a variety of physicochemical factors [3]. These include media composition, incubation temperature, inoculum age, carbon source, nitrogen source, pH, phosphate concentration, aeration, and others. Table 1.1

Strains and Strategies Adopted for a-Amylase Production

Microorganism

Method of Production

Bacillus subtilis TN 106 Streptomyces rimosus TM55

Fed batch SSF

Aspergillus oryzae Aspergillus oryzae OBS819 Neurospora crassa CFR 308 Streptomyces gulbargensis DAS 131 Bacillus acidicola Anoxybacillus beppuensis TSSC-1 Candida guilliermondii CGL-A10 Clostridium sp. BOH3

SSF SmF SSF SmF SmF SmF SmF

Microbacterium foliorum GA2

SmF

SmF, submerged fermentation; SSF, solid-state fermentation.

Substrate

Enzyme Yield

References

Sweet potato residue/ peanut residue Oil cake

2642.7 U/gds

[26] [30]

Coffee waste Starch

9196 U/gds 151.1 U/mL 7084 U/gds

2056 mmol/L/min 7.15 U/mg protein

[20] [32] [33] [34] [13] [11] [35] [15]

6610 U/mL

[16]

366 IU/L

Sugarcane bagasse hydrolyzate Bagasse


9

1.3.2.1 Incubation Temperature The effect of temperature on a-amylase production is related to the growth of the organism. Temperature control is very important in fermentation processes because growth and production of enzymes are sensitive to temperature. Hence, the optimum temperature varies with the culture. a-Amylases have been produced by various microbes over a wide range of temperature. Productions in SSF as well as in SmF are usually carried out in the range 25e37 C. However, psychrophilic and thermophilic temperatures have also been reported for the production. For example, a-amylase production was attained at 55 C by the thermophilic fungi Thermomonospora fusca [36] and T. lanuginosus [27] and at 80 C by a hyperthermophilic bacterium, Thermococcus profundus [36]. A psychrophilic bacterium, Alteromonas haloplanktis, produced a-amylase at 4 C [37].

1.3.2.2 pH The pH of the fermentation medium plays an important role in enzyme production. It induces morphological changes in the organisms as they are sensitive to the concentration of hydrogen ions present in the medium. A pH change in the medium affects the growth as well as the product stability. Unlike SmF, in which pH control is almost mandatory for a-amylase production, in SSF processes, generally there is no need to set, or control, the pH, as the substrates (agro-industrial residues) mostly possess excellent buffering capacity and keep the pH favorable for the growth and activity of the culture. Most of the Bacillus strains used commercially for the production of a-amylases have an optimum pH of 6.0 or 7.0. Some of the medium components eliminate the need for pH control. Yabuki et al. [38] reported that A. oryzae 557 accumulated a-amylase in the mycelia when grown in phosphate, or sulfate-deficient, medium and it was released when the mycelia were placed in a medium with pH above 7.2. Based on the optimal pH for activity, a-amylases are classified as acidic, neutral, and alkaline [1].

1.3.2.3 Carbon Sources a-Amylase production could be either constitutive or inducible. Galactose, inulin, and glycogen are suitable substrates for a-amylase production in SmF. Supplementation with lactose; an analog of maltose, a-methyl-D-glucoside; and yeast extract induces the production [7]. Several agro-residues such as wheat bran, rice bran, vegetable peels, fruit peels, cassava bagasse, and vegetable-oil-extracted residues are used as substrates for a-amylase production in SSF. Most studies on a-amylase production by A. oryzae suggest that the general inducer molecule is maltose. Eratt et al. [39] observed a 20-fold increase in enzyme activity when maltose and starch were used as inducers in A. oryzae NRC 401013. Xylose and fructose support good growth, but they are strongly repressive [40].

1.3.2.4 Nitrogen Sources Organic as well inorganic nitrogen sources are used for the production of a-amylases, although organic sources have been preferred over inorganic nitrogen sources. Commonly


used nitrogen sources include bactopeptone, ammonium sulfate, ammonium nitrate, Vogel salts, casein, meat extract, beef extract, yeast extract, corn steep liquor, and soybean flour. There are reports on the use of several other nitrogenous sources for a-amylase production. For example, L-asparagine was reported as the better nitrogen source for enzyme production by T. lanuginosus; casein hydrolyzate and yeast extract improved a-amylase production several fold and by 110e156%, respectively, by A. oryzae [41]. Complex nitrogen sources in the medium influence the production of a-amylases. Studies carried out by Dettori et al. [42] revealed that the supplementation of two organic nitrogen sources enhanced amylase production and this was better than a single organic nitrogen source.

1.3.2.5 Metal Ions Supplementation of metal ions in the fermentation medium promotes microbial growth, which in turn accelerates the enzyme production. Most a-amylases are known to be metal dependent for divalent ions, e.g., Ca2þ, Mg2þ, Mn2þ, and Zn2þ [2]. Supplementation with Ca2þ is generally required for an increased in a-amylase production by several bacteria. Ca2þ imparts thermostability of the enzyme due to salting out of hydrophobic residues by Ca2þ in the protein. The production was reduced to 50% when Mg2þ was omitted from the medium; Naþ and Mg2þ showed coordinated stimulation of enzyme production by Bacillus sp. CRP strain [43]. However, some metal ions could have a negative impact on the microbes for a-amylase production, e.g., Li2þ and Hg2þ have negative effects on a-amylase production. Mg2þ also plays an important role in a-amylase production.

1.3.2.6 Surfactants Addition of surfactants to the fermentation medium is generally known to increase the secretion of proteins by increasing cell membrane permeability. The commonly used surfactants are Tween 80, Tween 40, Triton X-100, sodium dodecyl sulfate (SDS), polyethylene glycol, and glycerol. These surfactants are reported to increase cell permeability, thereby enhancing enzyme yield. Arneson et al. [44] reported a twofold increase in a-amylase production by T. lanuginosus. Goes and Sheppard [45] reported a significant advantage in using the bio-surfactant surfactin to enhance the production of a-amylase by B. subtilis in SSF. In addition to increasing the enzyme activity, surfactin offers other advantages, including eco-friendliness, less sensitivity to extremes of temperature and pH, and being a potential fungicide, thereby eliminating contamination of the exposed substrate, compared to synthetic surfactants.

1.3.2.7 Agitation Agitation influences the mixing as well as the oxygen transfer rate in most fermentations and thus influences cell morphology and product formation [46,47]. It is generally believed that higher agitation is detrimental to cell growth, which in turn could decrease enzyme production. Agitation intensities up to 300 rpm are normally employed for the production of a-amylase in SmF from various microorganisms.


11

1.4 Assay of a-Amylases Activity of a-amylases is quantified by measuring either the end products, like glucose or maltose, or the amount of substrate that remains after enzymatic hydrolysis. a-Amylases are assayed using soluble starch or modified starch as the substrate. They catalyze the hydrolysis of a-1,4 glycosidic linkages in starch to produce glucose, dextrins, and limit dextrins. The reaction is monitored by an increase in the reducing sugar levels or a decrease in the iodine color of the treated substrate. Various methods are available for the determination of a-amylase activity [48]. These are based on a decrease in starcheiodine color intensity, increase in reducing sugars, degradation of color-complexed substrate, and decrease in viscosity of the starch suspension [3]. The common methods employed for the determination of a-amylase activity are the iodine method [49]; dextrinizing activity [50]; Sandstedt, Kneen, and Blish method [51]; dinitrosalicylic acid method [52]; and degradation of color-complexed substrate [53,54]. The dinitrosalicylic acid (DNS) method [52] is among the most commonly used methods for estimating the reducing sugars. The DNS reacts with reducing sugar under boiling and turns to red from yellow. In the method of Fuwa et al. [50], the starch reacts with iodine and forms a blue solution and the intensity of the color is directly proportional to the starch concentration. Boron dipyrromethene-labeled substrate releases a fluorescent fragment upon digestion with the enzyme and has been developed for determining a-amylase activity in foods [55].

1.5 a-Amylase Inhibitors Proteinaceous a-amylase inhibitors have been isolated from plants and microorganisms [56]. These inhibitors control endogenous a-amylase activity or work in defense against pests and pathogens; some inhibitors are antinutritional factors. a-Amylase inhibitors belong to seven different protein structural families. Six types are from higher plants and one is from Streptomyces sp. High-resolution structures are available for target a-amylase and these structures indicate major diversity and some similarities in the structural basis of a-amylase inhibition. Various types of inhibitors include Streptomyces inhibitors, knottins, g-thionins, CM proteins, and kunitz-type, thaumatin-like, and lectin-like inhibitors. Some a-amylase inhibitors have adverse effects on nutrition due to their inhibition of digestive enzymes in humans and animals. a-Amylase inhibitors find application in obesity and diabetic therapy.

1.6 Strain Improvement Strain improvement is usually carried out to increase production as well as to improve the properties of the enzyme. The catalytic properties of enzymes are determined by their 3-D structure. Hence, enzyme properties can be altered by site-directed mutagenesis. Using this method, the properties of an enzyme can be improved, by making


Table 1.2

Some Strategies Adopted for Strain Improvement/Properties of a-Amylase

Microorganism

Improved Property

References

Bacillus subtilis BR151 Alternaria tenuissima FCBP 252 Thermobifida fusca NTU22 Bacillus amyloliquefaciens Anoxybacillus sp.

Thermostability 2.39-fold increased production Increased production Increased production (1.4-fold) High stability in absence of Ca2þ ions at 60 C and high levels of maltose production Increased production (2.1-fold) High rate of maltose production Self-inducible, catabolite repression free, and glucose-activated expression system Increased production (7-fold) and high stability in absence of Ca2þ Oxidative stability Improved protein stability and catalytic efficiency Increased production

[60] [62] [63] [71] [66]

Aspergillus oryzae IIB 30 Paenibacillus sp. Bacillus licheniformis MSG B. subtilis ASO1a Thermotoga maritima B. subtilis Bacillus sp. AAH-31

[70] [67] [68] [69] [72] [73] [74]

it thermostable, reducing its dependence on cofactors, or increasing its activity at low temperature. Studies on the cloning of the a-amylase gene have been extensively carried out for hyperproduction [7]. Table 1.2 presents some strategies that have been adopted for strain improvement of a-amylase. a-Amylases have been engineered for the improvement of properties such as pH tolerance, thermotolerance, etc. [57e59]. Barnett et al. [58] found that the introduction of disulfide bonds in the enzymes and alteration of amino acids prone to oxidation by an amino acid resistant to oxidizing agents improved the stability of the enzyme. Suzuki et al. [57] constructed hybrids of homologous strains of the B. licheniformis and B. amyloliquefaciens with improved thermostability. Ozcan and Ozcan [60] introduced the thermostable plasmid pC194Amy, harboring a 5.2-kb DNA fragment encoding a gene of B. stearothermophilus, into B. subtilis BR151 by electroporation. The recombinant strains produced more thermostable a-amylase compared to the wild-type strain. A new strain of B. licheniformis CBBD302, carrying a recombinant plasmid, pHY-amyL, for B. licheniformis a-amylase (BLA) production, was constructed by Niu et al. [61]. The combination of target-directed screening and genetic recombination led to an approximately 26-fold improvement in BLA production and export in B. licheniformis. Shafique et al. [62] reported the production of an extracellular amylase from Alternaria tenuissima FCBP 252 in SSF. Chemical mutagenesis using ethyl methanesulfonate (EMS) produced mutants with a high level of a-amylase activity (2.39-fold) compared to the parental strain. Genetic characterization of the mutants using random amplified polymorphic DNA PCR revealed that the expression patterns of the mutants were isogenic variants of the parent strain. Yang et al. [63] expressed


13

an a-amylase gene from Thermobifida fusca NTU22 in Pichia pastoris X33 because of its potential application as a food supplement. Recombinant expression resulted in higher levels of extracellular enzyme production (510 U/L), indicating constitutive expression and secretion of the protein. The amount of extracellular protein in the culture of P. pastoris transformants was less than that in the cell-free extract of Escherichia coli transformants, hence facilitating the application of crude amylase in industry without purification. The gene encoding the a-amylase enzyme in B. subtilis PY22 was amplified by PCR, sequenced, and cloned into P. pastoris KM71H strain using the vector Ppicz A, allowing methanol-induced expression and secretion of the protein [64]. Recombinant expression resulted in high levels of extracellular amylase production (22 mg/L). The presence of Ca2þ ions in the medium resulted in a 41% increase in a-amylase activity. Expression in P. pastoris not only increased the yield of production but also potentially helped facilitate purification. Gene cloning and heterologous expression of the high-maltoseproducing a-amylase of Rhizopus oryzae showed successful expression of R. oryzae a-amylase in P. pastoris at a high level (382 mg/L) [65]. The enzyme had an extremely high affinity for maltotriose and no maltotriose remained after hydrolysis. Chai et al. [66] cloned two genes that encoded a-amylases from Anoxybacillus sp. and expressed them in E. coli. The enzymes produced by the recombinant strains were highly stable even in the absence of calcium at 60 C for 48 h and they produced high levels of maltose. Protein sequencing revealed that the recombinant a-amylase differed in 17 amino acids compared to the amylase produced by the wild-type strain. A gene encoding a-amylase from the genomic DNA of Paenibacillus sp. and the heterologous expression of recombinant Amy1 in E. coli BL21 (DE3) facilitated the recovery of this protein in soluble form. The high rate of maltose production due to the action of Amy1 could be exploited for the production of simple sugars as a by-product in food waste processing [67]. The use of an expression system to overcome catabolite repression opens up an avenue for exploiting cheap carbon sources for the production of recombinant enzyme. Nathan and Nair [68] developed a repression-free catabolite-enhanced expression system for a thermophilic a-amylase from B. licheniformis MSG. A self-inducible, catabolite repression-free, and glucose-activated expression system was developed using a thermophilic a-amylase as a model. The a-amylase gene from B. licheniformis MSG without any 50 cre operator produced unimpeded glucose-enhanced expression when fused to the phosphate starvation-inducible strong pst promoter with optimum translation signals in a protease-deficient B. subtilis. The yield was 18.5-fold higher than that of native promoter. Roy et al. [69] cloned and overexpressed a raw-starch-digesting a-amylase gene (AmyBS-I) from B. subtilis strain ASO1a in E. coli BL21. The gene also included its signal peptide sequence for the efficient extracellular expression of recombinant a-amylase in correctly folded form. The extracellular secretion of AmyBS-I was sevenfold higher and it did not require Ca2þ ions for its a-amylase activity/thermostability, which was an added advantage for its use in the starch industry.


Random mutagenesis has also been used for enhanced production of a-amylase. A strain of A. oryzae IIB 30 was subjected to physical (using UV light) and chemical mutagenesis (using nitrous acid and EMS). Mutation using EMS-20 showed a 2.1-fold increased amylase activity compared to the wild-type strain [70]. An identical observation was earlier reported for a B. amyloliquefaciens strain in which mutation using EMS improved enzyme activity by 1.4-fold higher than that of the parental strain [71]. Ozturk et al. [72] reported site-directed mutagenesis of methionine residues for improving the oxidative stability of a-amylase from Thermotoga maritima. The oxidative stability of a-amylase (AmyC) was improved by mutating the methionine residues at positions 43 and 44, and 55 and 62, to oxidative-resistant alanine residues. The mutant exhibited improved oxidative properties. The engineered AmyC could be a potential candidate for industrial applications, especially in the presence of oxidizing agents. This is the first protein engineering attempt for AmyC from T. maritima. Yang et al. [73] carried out structural engineering of histidine residues in the catalytic domain of a-amylase from B. subtilis for improved protein stability and catalytic efficiency under acidic conditions by site-directed mutagenesis. The four histidine residues His222, His275, His293, and His310 in the catalytic domain were selected as the mutation sites and were further replaced with acidic aspartic acid, respectively yielding four mutants H222D, H275D, H293D, and H310D. The acidic stability of the enzyme was significantly enhanced after mutation, and 45e92% of the initial activity of the mutants was retained after incubation at pH 4.5 and 25 C for 24 h, whereas that for the wild type was only 39.5%. As revealed by the structure models of the wild-type and mutant enzymes, the hydrogen bonds and salt bridges were increased after mutation, and an obvious shift of the basic limb toward acidity was observed for the mutants. These changes around the catalytic domain contributed to the significantly improved protein stability and catalytic efficiency at low pH. This work provided an effective strategy to improve the catalytic activity and stability of a-amylase under acidic conditions, and the results indicated potential application for the improvement of acid resistance of other enzymes. The hydrolytic activity of thermophilic, alkalophilic a-amylase could also be enhanced through the optimization of amino acid residues surrounding the substrate binding site [74]. Twenty-four selected amino acid residues were replaced with Ala, and Gly429 and Gly550 were altered to Lys and Glu, respectively, based on comparison of AmyL’s amino acid sequence with related enzymes. Y426A, H431A, I509A, and K549A showed higher activity than the wild type at 162e254% of wild-type activity. Tyr426, His431, and Ile509 were predicted to be located near subsite 2, and Lys549 was near subsite þ2. Ser, Ala, Ala, and Met were the best amino acid residues for the positions of Tyr426, His431, Ile509, and Lys549, respectively. Combinations of the optimized single mutations at distant positions were effective in enhancing catalytic activity. The doublemutant enzymes Y426S/K549M, H431A/K549M, and I509A/K549M, combining two of the selected single mutations, showed 340%, 252%, and 271% of wild-type activity, respectively. Triple- and quadruple-mutant enzymes of the selected mutations did not show higher activity than the best double mutant, Y426S/K549M.


15

1.7 Purification and Characterization of a-Amylases a-Amylases produced by fermentation are relatively crude preparations. Most of the commercial use of a-amylase does not require 100% purification of the enzyme. But, high-purity enzymes are required when they are used in clinical and pharmaceutical sectors. The first steps in the purification involve the isolation of crude enzyme after the fermentation. In SmF, this is usually done by centrifuging the fermented medium and taking the supernatant as the source of crude enzyme; in the case of SSF, the fermented matter is usually mixed with water or buffers, and after suitable mixing the contents are filtered, whereby the filtrate contains the crude enzyme. Then, the enzyme is concentrated (in the supernatant/filtrate), precipitated (using salts/solvents), and purified using various chromatographic techniques such as ion-exchange chromatography, gel filtration, isoelectric focusing, etc. Table 1.3 presents strategies adopted for the purification of a-amylase from various microorganisms. There are a large number of reports on the purification and characterization of a-amylases produced by bacterial or fungal sources in SmF and SSF [75e87]. An enzyme produced in SSF was partially purified by ammonium sulfate fractionation. The enzyme was optimally active at pH 5.0 and 50 C with a molecular mass of 66 kDa. The presence of Mn2þ and Fe2þ enhanced the enzyme activity, whereas in the presence of Hg2þ and Cu2þ the activity was reduced [76]. A partially purified a-amylase from Streptomyces erumpens MTCC 7317 showed a molecular mass of 54,500 Da [77]. a-Amylase from B. subtilis KIBGE-HAS was purified by ultrafiltration and ammonium sulfate precipitation with 19.2fold purification and specific activity of 4195 U/mg. The enzyme was highly stable in the presence of various surfactants and detergents. Metal ions such as Mn2þ, Ca2þ, Mg2þ, Kþ, Co2þ, and Fe3þ activated the enzyme, whereas Ba2þ, Cu2þ, Naþ, and Al3þ strongly inhibited the activity. A highly efficient raw-starch-digesting a-amylase from B. licheniformis ATCC 9945a was purified by gel-filtration chromatography with a sixfold increase in specific activity and recovery of 38% with a molecular mass of 31 kDa [82]. The purified enzyme showed an optimum pH and temperature of 6.5 and 90 C, respectively. Co2þ, Ni2þ, and Ca2þ slightly stimulated, whereas Hg2þ completely inhibited, a-amylase activity. An a-amylase from Brevibacterium linens DSM 20158, purified by ion-exchange chromatography on a DEAEeSephadex column, showed a 7.88-fold increase in purity with a 16.80% yield, and it appeared homogeneous on SDSepolyacrylamide gel electrophoresis with a molecular mass of 58 kDa. EDTA and Hg2þ inhibited the enzyme activity, whereas Mn2þ and Ca2þ enhanced the enzyme activity [83]. A novel SDS- and surfactant-stable, raw-starch-digesting, and halophilic a-amylase was purified from Nesterenkonia sp. [17]. The extracellular a-amylase was purified to homogeneity by 80% ethanol precipitation, Q-Sepharose anion-exchange chromatography, and Sephacryl S-200 gel-filtration chromatography. The optimum temperature and pH were 45.8 C and 7.5, respectively. The molecular mass was estimated as 100 kDa. The enzyme was inhibited by EDTA, but was not inhibited by phenylmethanesulfonyl fluoride and b-mercaptoethanol. Ca2þ stimulated enzyme activity, whereas the enzyme

Table 1.3

Strategies Adopted for Purification of a-Amylase from Various Microorganisms

Bacillus subtilis ATCC 465 Aspergillus oryzae Streptomyces erumpens MTCC 7317 B. subtilis KIBGE-HAS

B. subtilis C10

Penicillium janthinellum NCIM 4960 Nesterenkonia sp.

Aspergillus flavus Bacillus licheniformis ATCC 9945a Brevibacterium linens DSM 20158 Rhizopus microsporus

Aspergillus oryzae strain S2 Penicillium chrysogenum Bacillus methylotrophicus strain P11-2

Purification Strategy

Optimum Temperature

Optimum pH

Activators

Inhibitors

Molecular Mass

References [75]

Ammonium sulfate fractionation

50 C

5.0

Ultrafiltration and ammonium sulfate precipitation Polyethylene glycol/potassium phosphate aqueous two-phase system Ammonium sulfate fractionation/ anion-exchange chromatography (DEAE) 80% ethanol precipitation, QSepharose anion-exchange chromatography, and Sephacryl S200 gel-filtration chromatography Gel-filtration chromatography Ion-exchange chromatography on a DEAEeSephadex column Ammonium sulfate precipitation, Sephadex G25 desalination, and DEAE-52 cellulose chromatography Acetone precipitation, sizeexclusion and anion-exchange chromatography Ammonium sulfate precipitation and Sephadex G50 filtration 80% ammonium sulfate precipitation, DEAE FF anion exchange, and Superdex 75 10/ 300 gel-filtration chromatography

Mn2þ and Fe2þ

Hg2þ and Cu2þ

Mn2þ, Ca2þ, Mg2þ, Kþ, Co2þ, and Fe3þ

Ba2þ, Cu2þ, Naþ, and Al3þ

66 kDa 54.5 kDa

[76] [77] [78]

[79]

50 C

5.0

45.8 C

7.5

55 C 90 C

5.0 6.5

EDTA

42.7 kDa

[80]

Ca2þ

Fe3þ, Cu2þ, Zn2þ, and Al3þ

100 kDa

[17]

Co2þ, Ni2þ, and Ca2þ Mn2þ and Ca2þ

Hg2þ

55 kDa 31 kDa

[81] [82]

EDTA and Hg2þ

58 kDa

[83]

70 C

5.0

75 kDa

[84]

50 C

5.6

50 and 42 kDa

[85]

60 C

6.0

70 C

7.0

[86] Mg2þ, Ba2þ, Al3þ, and dithiothreitol

44 kDa

[87]


Microorganism


17

was strongly inhibited by Fe3þ, Cu2þ, Zn2þ, and Al3þ. An aqueous two-phase system comprising polyethylene glycol/potassium phosphate was used for the partition and purification of a-amylase from the culture supernatant of B. subtilis C10 [79], which resulted in a 3.56-fold purification of enzyme with a recovery of 59.37%. An a-amylase from Penicillium janthinellum NCIM 4960, purified by ammonium sulfate, showed an almost 20-fold increase in specific activity with a 30.73% yield after anion-exchange chromatography on DEAE cellulose. The purified enzyme had a molecular mass of 42.7 kDa. The optimum pH and temperature were 5.0 and 50 C, respectively. The enzyme showed substrate specificity toward amylose and amylopectin. The chelating agent EDTA inhibited enzyme activity. The enzyme was stable in the presence of commercial detergents and stability increased in the presence of CaCl2 [80]. An a-amylase produced by A. flavus isolated from mangrove soil was partially purified using ammonium sulfate, which resulted in a fivefold increase in enzyme activity. The partially purified enzyme was optimally active at pH 5.0, temperature of 55 C, with a molecular mass of 55 kDa. The extracellular amylase was purified by anionic- and cationic-exchange chromatography and preparative electrophoresis, which resulted in 38-fold purity [81]. Shen et al. [84] purified an acid-stable and thermostable a-amylase from Rhizopus microsporus isolated from distilled liquor. The crude extract was purified using ammonium sulfate precipitation, Sephadex G25 desalination, and DEAE-52 cellulose chromatography. The optimum pH and temperature were 5.0 and 70 C, respectively, with a molecular mass of 75 kDa.

1.8 Applications of a-Amylase a-Amylases find a wide range of biotechnological applications in the textile, food, pharmaceutical, and detergent industries. With the current developments in biotechnology, the applications of a-amylases have been widened to other fields like clinical, medicinal, and analytical chemistry.

1.8.1

Detergent Applications

a-Amylases comprise one of the ingredients of modern compact detergents. One of the main advantages of using enzymes in detergents is that much milder conditions can be used than with enzyme-free detergents [3]. Enzymes help in lowering of the washing temperature. a-Amylases have been used as powerful laundry detergents since 1975. Currently, 90% of all commercially available detergents contain a-amylase and the demand for automatic dishwasher detergents is growing. One of the main limitations of a-amylases in detergents is that it is highly sensitive to calcium and oxidants, which are components of detergents. However, the development of genetically modified strains for the production of a-amylases to improve their bleach stability has been achieved by replacing the oxidation-sensitive amino acids with other amino acids. Replacement of methionine at position 197 by leucine in B. amyloliquefaciens amylase has resulted in improved resistance against oxidative compounds [88e91].


1.8.2

Textile Desizing

a-Amylases find application in the textile industry for textile desizing. To protect yarn from breaking, a removable protective layer is applied to the threads. Starch is an ideal desizing agent because it is cheap, is readily available, and can be removed very easily [3]. An effective desizing of starch-sized textiles is achieved by the application of a-amylases, which selectively remove the sizing and do not attack the fibers. It randomly cleaves the starch into dextrins, which are water soluble and can be removed by washing. Amylase from Bacillus sp. has been widely used in textile industries for a long time. Sarvanan et al. [92] studied the desizing of cotton fabrics using a-amylase from B. licheniformis. The study revealed that pH and amylase concentration exhibited the dominant effect, followed by treatment time, on desizing efficiency.

1.8.3

Medicinal and Clinical Chemistry

There are several processes in the medicinal and clinical areas that require the application of a-amylases [3]. The first enzyme produced industrially was a fungal amylase in 1894 and was used as a pharmaceutical aid for the treatment of digestive disorders. Dumoulin et al. [93] observed that the addition of a-amylase to cross-linked amylose tablets could modulate the release kinetics of the drug.

1.8.4

Paper Industry

a-Amylases find application in the paper and pulp industry for the modification of starches for coating paper. The coating treatment serves to make the surface of the paper smooth and strong to improve the writing quality of the paper. Starch acts as a good sizing agent for the finishing of paper, improving the quality and erasability, in addition to being a good coating for the paper. The sizing enhances the stiffness and strength of the paper [94]. Cold-active a-amylase is used in the paper industry because it reduces the viscosity of starch for the appropriate coating of paper [95].

1.8.5

Starch Liquefaction and Saccharification

One of the most important applications of a-amylase is in starch liquefaction for the production of glucose and fructose syrups. Starch is converted to high-fructose corn syrup and is widely used in the beverage industry as a sweetener for soft drinks because of its high sweetening property. The process requires the usage of highly thermostable a-amylase for starch liquefaction [3]. The enzymatic conversion involves three processes, which include gelatinization, liquefaction, and saccharification. Gelatinization involves dissolution of starch granules, forming a viscous solution; liquefaction involves partial hydrolysis and leads to a loss in viscosity, followed by saccharification involving the production of maltose and fructose [96]. The enzymes from B. licheniformis and B. stearothermophilus are widely used because of their remarkable thermostability.


1.8.6

19

Bread and Baking Industry

a-Amylases have been widely used in the baking industry, especially in bread and rolls to give these products a higher volume, better color, and softer crumb. The enzymes degrade starch into smaller dextrins, which are subsequently fermented by the yeasts. This generates additional sugar in the dough, which improves the crust color, taste, and toasting properties of bread. a-Amylase acts as an antistaling agent and improves the softness retention and shelf life of baked foods. Generally, a-amylases from B. stearothermophilus are used commercially in the baking industry [97].

1.8.7

Alcohol Production

For the production of ethanol from starch, it has to be solubilized and then submitted to two enzymatic steps to obtain fermentable sugars. The conversion process is the liquefaction, which is carried out by a-amylases, followed by the saccharification using glucoamylases, leading to the formation of a hydrolyzate containing sugars, which are then fermented by the yeast to ethanol [98].

1.9 Conclusion and Perspectives a-Amylases are important enzymes for many industrial processes. Though a-amylases are produced by several microbes, bacterial amylases are commonly preferred because of their thermotolerance for several applications. Most of the industrial processes that involve the usage of a-amylase are carried out under extreme conditions of temperature and pH. The major challenges in the commercial production of amylases are the yield, stability, and cost of production. Although considerable successes have been achieved in the production of a-amylases with increased productivity and desired properties for industrial applications, to sustain the economic feasibility and newer technological applications, continued research and technological developments are needed through biotechnological interventions. To meet the industrial demand, it is important to understand the structureefunction relationship. With the development and application of modern techniques such as protein engineering, metabolic engineering, etc., it could be possible to develop tailor-made a-amylases for application in various sectors. Hence, research and technological development efforts must to be directed toward the development and construction of a-amylases with novel and improved properties.

References [1] Sharma A, Satyanarayana T. Microbial acid-stable a-amylases: characteristics, genetic engineering and applications. Process Biochemistry 2013;48:201e11. [2] Pandey A, Nigam P, Soccol VT, Singh D, Mohan R. Advances in microbial amylases. Biotechnology and Applied Biochemistry 2000;31:135e52. [3] Gupta R, Gigras P, Mohapatra H, Goswami VK, Chauhan B. Microbial a-amylases: a biotechnological perspective. Process Biochemistry 2003;38:1599e616.


[4] Lonsane BK, Ramesh MV. Production of bacterial thermostable a-amylase by solid state fermentation: a potential tool for achieving economy in enzyme production and starch hydrolysis. In: Advances in applied microbiology, vol. 35. San Diego: California Academic Press; 1990. p. 1e56. [5] Pradeep NV, Ankitha AK, Pooja J. Categorizing phenomenal features of a-amylase (Bacillus species) using bioinformatic tools. Journal of Advances in Life Science and Technology 2012;4: 27e35. [6] Bordbar AK, Omidiyan K, Hosseinzadeh R. Study on interactionof a-amylase from Bacillus subtilis with cetyl trimethylammonium bromide. Colloids and Surfaces B: Biointerfaces 2005;40: 67e71. [7] Sivaramakrishnan S, Gangadharan D, Nampoothiri KM, Soccol CR, Pandey A. a- amylases from microbial sources e an overview on recent developments. Food Technology and Biotechnology 2006;44:173e84. [8] Stanley D, Farnden KJF, MacRae EA. Plant a-amylases: functions and roles in carbohydrate metabolism. Biologia, Bratislava 2005;60:65e71. [9] Naidu MA, Saranraj P. Bacterial amylase: a review. International Journal of Pharmaceutical and Biological Archives 2013;4:274e87. [10] Setyorini E, Takenaka S, Murakami S, Aoki K. Purification and characterization of two novel halotolerant extracellular proteases from Bacillus subtilis strain. Bioscience, Biotechnology and Biochemistry 2006;70:433e40. [11] Kikani BA, Singh SP. The stability and thermodynamic parameters of a very thermostable andcalcium-independent a-amylase from a newly isolated bacterium, Anoxybacillus beppuensis TSSC-1. Process Biochemistry 2012;47:1791e8. [12] Manojkumar N, Karthikeyan S, Jayaraman G. Thermostable alpha-amylase enzyme production from Bacillus laterosporus: statistical optimization, purification and characterization. Biocatalysis and Agricultural Biotechnology 2013;2:38e44. [13] Sharma A, Satyanarayana T. Optimization of medium components and cultural variables for enhanced production of acidic high maltose-forming and Ca2þ independent a-amylase by Bacillus acidicola. Journal of Bioscience and Bioengineering 2011;111:550e3. [14] Wang S, Liang Y, Liang T. Purification and characterization of a novel alkali-stable a-amylase from Chryseobacterium taeanense TKU001, and application in antioxidant and prebiotic. Process Biochemistry 2011;46:745e50. [15] Rajagopalan G, He J, Yang KL. Production, purification, and characterization of a-amylase from solventogenic Clostridium sp. BOH3. Bioenergy Research 2014;7:132e41. [16] Roohi, Kuddus M. Bio-statistical approach for optimization of cold-active a-amylase production by novel psychrotolerant M. foliorum GA2 in solid state fermentation. Biocatalysis and Agricultural Biotechnology 2014;3:175e81. [17] Shafiei FM, Ziaee A, Amoozegar MA. Purification and biochemical characterization of a novel SDS and surfactant stable, raw starch digesting, and halophilic a-amylase from a moderately halophilic bacterium, Nesterenkonia sp. strain. Process Biochemistry 2010;45:694e9. [18] Jeon E, Jung J, Seo D, Jung D, Holden JF, Park C. Bioinformatic and biochemical analysis of a novel maltose-forming -amylase of the GH57 family in the hyperthermophilic archaeon Thermococcus sp. CL1. Enzyme and Microbial Technology 2014;60:9e15. ¨ zdemir S, Bekler M. Purification and characterization of thermostable lu Fincan S, Enez B, O ¨ log [19] Agu a-amylase from thermophilic Anoxybacillus flavithermus. Carbohydrate Polymers 2014;102:144e50. [20] Ramachandran S, Patel AK, Nampoothiri KM, Chandran S, Szakacs G, Soccol CR, et al. Alpha amylase from a fungal culture grown on oil Cakes and its properties. Brazilian Archives of Biology and Technology 2004;47:309e17.


21

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23

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[80] Sindhu R, Suprabha GN, Shashidhar S. Purification and characterization of a-amylase from Penicillium janthinellum (NCIM 4960) and its application in detergent industry. Biotechnology Bioinformatics and Bioengineering 2010;1:37e45. [81] Bhattacharya S, Bhardwaj S, Das A, Anand S. Utilization of sugarcane bagasse for solid- state fermentation and characterization of a- amylase from Aspergillus flavus isolated from Muthupettai mangrove, Tamil Nadu, India. Australian Journal of Basic and Applied Sciences 2011;5:1012e22. [82] Bozic N, Ruiz J, Santin J, Vujcic Z. Production and properties of the highly efficient raw starch digesting a-amylase from a Bacillus licheniformis ATCC 9945. Biochemical Engineering Journal 2011;53:203e9. [83] Shabbiri K, Adnan A, Noor B, Jamil S. Optimized production, purification and characterization of alpha amylase by Brevibacterium linens DSM 20158, using bio-statistical approach. Annals of Microbiology 2012;62:523e32. [84] Shen H, Mo X, Chen X, Han D, Zhao C. Purification and enzymatic identification of an acid stable and thermostable a-amylase from Rhizopus microsporus. Journal of the Institute of Brewing 2012; 118:309e14. [85] Sahnoun M, Bejar S, Sayari A, Triki MA, Kriaa M, Kammoun R. Production, purification and characterization of two a-amylase isoforms from a newly isolated Aspergillus oryzae strain S2. Process Biochemistry 2012;47:18e25. [86] Doss A, Anand SP. Purification and optimization of fungal amylase from litter samples of Western Ghats, Coimbatore, Tamilnadu (India). Journal of Scientific Research and Reviews 2013;2:001e4. [87] Xie F, Quan S, Liu D, Ma H, Li F, Zhou F, et al. Purification and characterization of a novel a-amylase from a newly isolated Bacillus methylotrophicus strain P11-2. Process Biochemistry 2014;49:47e53. [88] Svendsen A, Bisgaard-Frantzen H. PCT patent publication. WO 94/0 1994. [89] Tierny L, Danko S, Dauberman J, Vaha-Vahe P, Winetzky D. Performance advantages of novel a-amylases in automatic dishwashing. In: Am oil Chem Soc 86th san Antonio Annual meeting; 1995. [90] Bisgaard-Frantzen H, Borchert T, Svendsen A, Thellersen MH, Van Der Zee P. PCT Patent Application. WO 95/10603 1995. [91] Kiran KK, Chandra TS. Production of surfactant and detergent-stable, halophilic, and alkali tolerant alpha-amylase by a moderately halophilic Bacillus sp. Strain TSCVKK. Applied Microbiology and Biotechnology 2008;77:1023e31. [92] Saravana D, Prakash AA, Jagadeeshwaran D. Optimization of thermophilic Bacillus licheniformis a-amylase desizing of cotton fabrics. Indian Journal of Fiber and Textile Research 2011;36:253e8. [93] Dumoulina Y, Cartiliera L, Mateescub M. Cross-linked amylose tablets containing a-amylase: an enzymatically-controlled drug release system. Journal of Controlled Release 1999;60:161e7. [94] Bruinenberg PM, Hulst AC, Faber A, Voogd RH. A process for surface sizing or coating of paper. 1996. European Patent Application, EP 0 690 170 A1. [95] Kuddus M. Microbial cold-active a-amylases: from fundamentals to recent developments. Current Research, Technology and Education Topics in Applied Microbiology and Microbial Biotechnology 2010:1265e76. [96] de Souza PM, de Oliveira Magalha˜es P. Application of microbial a-amylase in industry e a review. Brazilian Journal of Microbiology 2010;41:850e61. [97] van der Maarel MJ, van der Veen B, Uitdehaag JC, Leemhuis H, Dijkhuizen L. Properties and applications of starch-converting enzymes of the alpha-amylase family. Journal of Biotechnology 2002;94:137e55. ¨ ner ET. Optimization of ethanol production from starch by an amylolytic nuclear petite [98] O Saccharomyces cerevisiae strain. Yeast 2006;23:849e56.

2

Amylolytic Enzymes: Glucoamylases S. Negi*, K. Vibha MOTILAL NEHRU NATIONAL INSTITUTE OF TECHNOLOGY , A LLAHABAD , IND IA

2.1 Introduction Starch is one of the most abundant polymers on earth and its industry has a big stake in the market. Starch is composed of unbranched amylose and branched amylopectin, which require three types of amylolytic enzymes for complete hydrolysis into glucose, i.e., a-amylase (4-a-D-glucan glucanohydrolase, EC 3.2.1.1), b-amylase (4-a-D-glucan maltohydrolase, EC 3.2.1.2), and glucoamylase (4-a-D-glucan glucohydrolase, EC 3.2.1.3). a-Amylase cleaves the a-1,4-D-glucosidic linkages between adjacent glucose units in the linear amylose chain; b-amylase cleaves at nonreducing chain ends of amylose, amylopectin, and glycogen molecules; and GA hydrolyzes a-1,4 glycosidic bonds from the nonreducing ends of starch and a-1,6 linkages at the branching points of amylopectin, although at a lower rate than 1,4 linkages, into glucose [1e4]. GA can also catalyze the reverse hydrolysis reaction to produce maltose and isomaltose, which has great significance in industrial processes in which high sugar content is present. GA converts starch and a- and b-limit dextrins into glucose and shows faster reaction on polysaccharides than on oligosaccharides. The rate of hydrolysis depends on the substrate size and the structure, nature, and position of the bond present. GA is ubiquitously present in or produced by all forms of life (plants, animals, bacteria, archaea, and eukaryotes). However it is mainly produced using filamentous fungi, although a host of other microorganisms are also known as good producers of GA. Aspergillus niger, Aspergillus awamori, and Rhizopus oryzae are the commonly used filamentous fungi for industrial production of GA [9,10]. GAs are extensively used in the food and beverage industries. They are used for production of glucose syrup, high-fructose corn syrup, beer, soy sauce, alcoholic beverages, etc. [2,9,11e13]. Most of the GA produced from parent strains catalyzes saccharification efficiently only within a small range of mild temperatures. At high temperatures its catalytic activity reduces sharply because of conformation changes. GA produced from parent fungal sources normally has limited thermostability, catalytic activity, and low pH range, which restrict its application in industrial processes carried out at high temperature and in alkaline medium. At higher temperature the reaction rate is higher; therefore, *



25


processing is faster. It also prevents microbial contamination and reduces the viscosity of the reaction mixture. This leads to reduction in process cost. Production of GA that is stable at higher temperatures would be highly beneficial for starch saccharification. Advances in recombinant DNA technology and site-directed and random mutagenesis and other techniques are being used to improve the thermostability and other functional properties of GA [1]. Over the years a lot of research has been carried out to reduce the cost of production of GA and improve its functional properties to suit industrial requirements. Progress in the fields of molecular biology, protein engineering, and bioinformatics has helped to provide it with improved functional properties, such as enhanced thermostability, better selectivity, wider pH range, improved catalytic activity, etc. [14].

2.2 Sources of Glucoamylase GA occurs in a wide range of organisms. It is present in plants [15], animals [16], fungi, bacteria, and yeast. However, microorganisms are the main source explored for GA production.

2.2.1

Microbial Sources

GA is present in a wide range of bacteria, fungi, and yeasts. Some of the microbial sources exploited for the production of GA are shown in Table 2.1.

2.2.1.1 Fungal Sources Fungal species of Aspergillus, Rhizopus, and Endomyces are the most commonly used sources for production of GA. Aspergillus awamori and A. niger are among the most popular microorganisms used by industry for GA production [2,3,17]. Rhizopus oryzae [18], Rhizopus niveus [19], Mucor [6,20], Penicillium [21], and many other fungal species are capable of producing GA. Enzyme production by molds is generally extracellular, which makes its downstream processing cost-effective and less time consuming.

2.2.1.2 Bacterial Sources There are many bacterial strains capable of producing GA. Flavobacterium sp. [22], Bacillus stearothermophilus [23], Sclerotinia sclerotiorum [24], Sclerotium rolfsii [25], and Thermoanaerobacter tengcongensis [26] are some of the bacteria used for production of GA. The amylolytic bacterial strains Clostridium thermosaccharolyticum [27], Clostridium sp. [28], and Lactobacillus amylovorus [29] have also been used for production of GA [2]. Thermostable GA from thermophilic bacteria can be used at higher temperature for saccharification, thereby reducing the production cost of glucose by saving the cost of cooling. For production of glucose from starch, liquefaction of starch is done by a-amylase first, followed by saccharification by GA. Liquefaction can take place rapidly at 95e105 C by a thermostable bacterial a-amylase. But fungal GAs are stable normally up to a temperature range of 55e60 C. Therefore, most of the fungal GAs are used only

Chapter 2 Amylolytic Enzymes: Glucoamylases

27

Table 2.1 Details of Substrates and Microorganisms Used for Production of Glucoamylase Source

Microorganism

Process

Substrate

Yield

References

Fungal

SSF

Corn flour, wheat bran Wheat bran Wheat bran

5582.4 mmol/m g

[37]

45.21 U/mL 2.0 and 1.99 mmol/mL min

[38] [5]

Fungal

Aspergillus oryzae FK-923 A. oryzae Aspergillus niger and Rhizopus A. oryzae

SSF

4.1 IU

[39]

Fungal Fungal Fungal

A. niger A. niger A. oryzae

SSF SSF SSF

31.214 U/mg 152.85 U/mL 330 mg/mL min

[7] [40] [41]

Fungal

Aspergillus niveus

SmF

600 U/mg of protein

[42]

8.3 U/mL 26.3 U/mL

[9] [43]

Fungal Fungal

Fungal Fungal

SSF SSF

SSF SSF

Aspergillus awamori Thermomucor indicae-seudaticae Fungal A. awamori Fungal Aspergillus flavus and Thermomyces lanuginosus Fungal A. niger Fungal T. lanuginosus Fungal Colletotrichum gloeosporioides Bacterial Thermoanaerobacter tengcongensis Bacterial Bacillus sp. Bacterial Lactobacillus amylovorus Yeast Candida famata Yeast Pichia subpelliculosa Plant

Liquid

Sugar beet (Beta vulgaris L.)

Wheat bran and rice bran Potato starch Potato starch Wheat bran and sugarcane bagasse Starch and yeast extract Potato starch Sucroseeyeast extract

SSF SmF

Wheat bran Cassava and corn starch

9420.6 U/gds

[44] [45]

SSF SSF SSF

Rice bran Starch Starch

695 U/g 60 U/mg of protein 0.45 IU/mL

[46] [47] [4]

SSF

Maltose

80 U/mg

[26]

SSF SSF SSF SmF

Starch Dextrin and starch Starch Commercial washed starch MurashigeeSkoog medium

21.6 U/mL 2587.17 mmol/L m 6500 U/L

[30] [29] [31] [32]

102.2 U/mg

[36]

SmF, submerged fermentation; SSF, solid-state fermentation.

after the reaction mixture is cooled down to this temperature. Thermostable GA from thermophilic bacteria can be used for saccharification without much cooling of the liquefied starch. GA produced from aerobic bacteria, such as B. stearothermophilus, Halobacterium sodomense, and Flavobacterium sp., and anaerobic bacteria, such as Clostridium sp. and C. thermosaccharolyticum, have better thermostability compared to fungal GA [26].


A thermostable GA (TtcGA) from T. tengcongensis was successfully expressed in Escherichia coli by Zheng et al. [26]. Heat treatment and gel-filtration chromatography were used to partially purify the recombinant mature protein, and 30-fold homogeneity was obtained. Maximum activity of the recombinant enzyme was obtained at 75 C and pH 5.0. It was highly thermostable with almost no activity loss at 75 C for 6 h. James and Lee (1995) used L. amylovorus to produce GA in dextrose-free de ManeRogosaeSharpe medium in a 1.5-L fermenter under an optimal dextrin concentration of 1% (w/v), pH 5.5, and 37 C. GA production was maximum at the late logarithmic phase of growth for 16e18 h. Crude enzyme showed maximum activity at pH 6.0 and 60 C. Gill and Kaur (2004) used a bacterial strain of Bacillus for production of TtcGA [30]. Highest production of GA was achieved at 65 C and pH 7.0 after 17e20 h under stationary conditions. Luria broth, supplemented with 0.5% (w/v) soluble starch, yielded highest enzyme activity (21.6 U/mL). GA showed optimum activity at 70 C and pH 5.0. It was highly stable at pH 7.0 with a half-life of 13 h, 8 h, and 3 h 40 min at 60, 65, and 70 C, respectively.

2.2.1.3 Yeast Sources Candida famata [31], Pichia subpelliculosa [32], and Saccharomyces diastaticus [33] are some of the yeast used for production of GA. GA production has also been reported from yeast strains of Saccharomycopsis fibuligera [34] and Lipomyces starkeyi HN-606 [35]. Mohamed et al. isolated C. famata from traditional Moroccan sourdough for production of GA [31]. Starch enhanced GA production, with maximum GA activity at 5 g/L. Yeast extract and (NH4)2HPO4 gave maximum GA and biomass after 72 h of incubation in liquid medium at 30 C, pH 5, at 105 rpm.

2.2.2

Other Sources

There are very few reports of GA production from plant and animal origins. In one such work on GA production from a plant origin, sugar beet (Beta vulgaris L.) was explored for production of GA and a-amylase [36]. They reported the presence of GA and a-amylase in callus and suspension cultures as well as in mature roots of sugar beets (B. vulgaris L.).

2.3 Glucoamylase Production GA is produced by many microorganisms capable of growing in a vast variety of substrates through the fermentation process. Like any other enzyme production, GA also depends on the selection of microbial strain, substrate, medium, and fermentation process and on physicochemical parameters such as incubation time, temperature, pH, relative humidity [in solid-state fermentation (SSF)], inhibitors, oxygen accessibility, etc. Production can be carried out through submerged fermentation (SmF) or SSF depending upon the microbial culture and substrate in use. Until recently, approximately 90% of all industrial enzymes were produced in SmF using a specifically optimized process and genetically manipulated microorganisms. However, scientists have discovered and


29

realized the numerous economical and practical advantages of SSF. Almost all enzymes can be produced in SSF using microorganisms. Some of the sources and GA production details are shown in Table 2.1.

2.3.1

Selection of Fermentation Process

In SmF the substrate is solubilized or suspended as fine particles in a large volume of liquid, whereas in SSF an insoluble substrate is fermented with sufficient moisture but with no free water. SSF is an ideal process when the organism is a filamentous fungus, which is able to withstand the limited water availability. SSF has become popular with scientists as well as industry, particularly with agro-based substrates [2,48,49]. SSF has distinct advantages over SmF in terms of lower production costs, lower energy requirements, higher yield, simple fermentation equipment, and less effluent generation. In SSF, nutrients present in the substrate serve as an anchorage for the microbial cells. In SSF the moisture content varies from 30% to 80% and maximum enzyme production is normally obtained with about 60% moisture content. At lower moisture content microbiological activity ceases.

2.3.2

Composition of Substrate

Composition of the substrate used for production of GA has a big impact on its yield. Selection of substrate is done considering its availability and suitability for the particular strain and process used. Agro-industrial residues like rice husk, wheat bran, rice bran, gram flour, coconut oil cake, sugarcane bagasse, wheat flour, corn flour, tea waste, potato starch, etc., are commonly used as substrates for GA production in SSF [2,17,38,49,50]. Corn starch is a commonly used substrate for production of glucose syrup in the United States and Europe because of its easy availability. The particle size of the substrate has a strong influence on the growth rate and enzyme-producing ability of the organism due to the influence of surface area on growth rates. The optimum particle size is to be decided to ensure maximum mass transfer because bigger particles provide less surface area and smaller particles provide a high degree of solubilization. Pandey reported poor GA activity with a particle size of larger than 1.4 mm and smaller than 180 mm [50]. Wheat bran particles of 425e500 mm were found most suitable for maximizing GA activity. Another vital factor affecting GA production in SSF is water activity, which is an important parameter for mass transfer of the water and solutes across the cell membrane. Pandey et al. reported higher GA yield with higher initial water activity values of substrate [51]. Substrates are normally supplemented with carbon or nitrogen sources to increase the enzyme yield. Supplement can be a simple carbon source like glucose, maltose, or sucrose or a polymeric compound such as starch from some other source, or a nitrogen source such as ammonium or nitrate salts or urea, or a complex source, e.g., corn steep liquor. The requirements for nutrition are normally more complex in SmF compared to SSF. Supplements of fructose, ammonium sulfate, urea, and yeast extract in the substrate,


such as wheat bran, have been reported to enhance GA production in SSF by Aspergillus sp. and A. niger [2,44,52]. Nyamful et al. (2014) reported wheat bran as the best substrate for production of GA by A. niger and Rhizopus strains. They used wheat bran, rice bran, and groundnut pod as different substrates for investigation. Maximum activities of 2.0 and 1.99 U/mL, respectively, for A. niger and Rhizopus were obtained after 48 h on wheat bran, though GA was produced by both strains on all substrates [5]. Ominyi et al. used Aspergillus sp., Mucor, and Rhizopus sp. for production of GA through SmF using soluble starch as a carbon source [6]. Puri et al. (2013) also used four different substrate compositions, i.e., rice bran, wheat bran, rice bran/wheat bran (1:1), and rice bran/paddy husk (1:1) for GA production from Aspergillus oryzae under SSF. A maximum GA activity of 4.11 IU was obtained with rice bran [39].

2.3.3

Optimization of Physical Parameters

Production of any enzyme in itself is not useful unless it meets the requirements of the industry. For industrial application, the yield of enzyme production, the time of production, and the enzyme’s stability are the basic requirements, so that suitable enzymes can be produced in a cost-effective manner. From the very beginning, various techniques have been used to optimize various parameters and supplements that affect the yield and quality of enzymes. Optimization techniques eliminate wasteful expenditure, experimentation, and calculation. Boas (1962) was the first to develop this technique, wherein optimization of the biological system was based on single-factor search [53,62]. Earlier, this “one-at-a time” method was used by workers for optimization, but was incapable of detecting the true optimum conditions, especially because of probable interaction aspects of enzymes among themselves. Thus, the necessity of an optimization technique that would investigate the interaction factor also was felt. This led to techniques such as the evolutionary operation (EVOP) program. Barnett (1960) reported that EVOP provides a system for exploring the relationships between independent and dependent variables [54,55]. He stated that EVOP consists of the systematic introduction of very small changes in the selected independent variables, which affects the process and statistical selection of the best set of conditions. Response surface methodology (RSM), based on factorial experiments, is another statistical approach to study the effects of test variables on measured response [55,56]. The mathematical model for RSM is derived from orthogonal polynomial fitting techniques. Negi and Banerjee (2006) employed the EVOP factorial design technique to achieve optimal pH, temperature, and humidity for production of GA using A. awamori from wheat bran under SSF. A maximum yield of 9420.6 U/gds GA was achieved at 37 C, pH 4, and 85% humidity [44]. Puri et al. (2013) optimized GA production by varying temperature, moisture content, pH, inoculum, and incubation period of culture. Optimization was carried out by varying temperature from 20 to 40 C, moisture content from 10 to 30 mL/ 5 g, pH from 3 to 7, spore suspension from 1 105 to 1 108 spores/mL, and incubation period from 3 to 6 days. A 30 C incubation temperature, 20 mL/5 g moisture content,


31

1 107 spores/mL spore suspension, pH 5.0, and 5-day incubation were found to be optimal conditions for GA production [39]. Keera et al. (2014) used A. oryzae to produce GA by SSF of corn flour. They optimized various parameters, i.e., initial moisture content (50e80% v/w), pH (3e9), and temperature (25e34 C), and used various supplements such as wheat bran (20e50%); carbon supplements (1e3% w/w) of glucose, maltose, sucrose, potato starch, and soluble starch; and inorganic supplements of ammonium sulfate, ammonium oxalate, ammonium, phosphate, diammonium phosphate, triammonium phosphate, ammonium acetate, ammonium nitrate, sodium nitrate, and urea to optimize enzyme yield. Maximum GA production of 5582.4 mmoles of glucose produced per minute per gram of dry fermented substrate was obtained after 72 h at 30 C on corn flour supplemented with 30% (w/w) wheat bran, 1% soluble starch, 0.1% (w/w) urea at pH 5.5 and 60% (v/w) initial moisture [37]. Kumar and Satyanarayana (2001) optimized GA production by the yeast strain P. subpelliculosa and achieved highest yield at 30 C and pH 5.6 in medium containing 0.2% yeast extract, 1% starch, 0.035% NaCl, 0.4% K2HPO4, and 0.1% MgCl2 when agitated at 200 rpm in a shake flask for 11 h. A 15-fold increase in GA secretion was achieved by optimization [32]. Parbat and Singhal optimized GA production under SSF by A. oryzae and maximum yield was obtained at 60 C, pH 5.0, with wheat bran and sugarcane bagasse at a ratio of 1:1 [41]. Gomes et al. (2005) optimized production of GA from starch in SmF by Aspergillus flavus and Thermomyces lanuginosus. Effects of different carbon sources, temperatures, and initial pH of the medium were evaluated using a full factorial design (2 2 3). Cassava starch gave better results than corn starch with A. flavus and exhibited higher activity with starch and maltose mixture. GA produced from A. flavus and T. lanuginosus had high optimum temperature, i.e., 65 and 70 C, respectively, and a wide pH range [45]. Negi and Banerjee (2010) optimized GA production concomitant with protease from A. awamori in a single fermentation and achieved a GA yield of 4528.4 121 U/gds with wheat bran at a Czapek Dox medium ratio of 1:1.5 (w/v), 96 h incubation, 35 C temperature, pH 5.5, and 85% relative humidity through the EVOP factorial design technique. Medium engineered with 1% casein and 1% starch solution further increased the yield by 2.07-fold (9386.5 101 U/gds) [57].

2.4 Purification and Characterization Crude enzyme is needed to be optimally extracted from the fermented mass using a suitable solvent and then purified. The extraction efficiency can be greatly improved by selecting a suitable solvent, concentration of the solvent, soaking time, temperature, and number of washes. The extent of purification depends on the application. Optimization of various extraction conditions such as type of solvent, concentration of the solvent, soaking time, and temperature is done to maximize the extraction of crude extract. Crude enzyme extracted in a suitable solvent normally contains some amount of unfermented mass, microbial cells, cell debris, spores, and other insolubles. Removal of


such insolubles is normally done by centrifugation and microfiltration, etc. Negi and Banerjee (2009) extracted and purified GA, produced concomitant with a protease, by A. awamori in a single fermenter by SSF, by soaking the fermented mass at room temperature (around 30 C) in 10% glycerol for 2 h and then using acetone in a 1:2 ratio for precipitation. GA was purified up to 4.06-fold with 35.3% recovery. To concentrate the desired extracellular protein, sequential and/or parallel precipitation can be used. Salting out with ammonium sulfate or any other suitable organic solvent can be used to precipitate total protein. Precipitation with cold-water-miscible organic solvents, organic polymers, and isoelectric precipitation are some of the methods adopted by scientists. To separate out the enzyme of interest various centrifugation methods, such as ultracentrifugation, density gradient centrifugation, etc., are also used [8].

2.4.1

Purification Techniques

Various techniques have been developed for the purification of proteins prior to their characterization or use in biotechnological and industrial processes. Purification can be achieved through a series of chromatographic techniques such as ion-exchange and gelfiltration chromatography, or in combination, or affinity chromatography and highperformance liquid chromatography. To check the level of purity and molecular weight determination, sodium dodecyl sulfateepolyacrylamide gel electrophoresis is commonly used. In lab scale, purification is generally preferred through ion-exchange and sizeexclusion chromatography; however, one-step purification affinity chromatography is more economical and preferred. For size exclusion Sephadex-G of different grades and silica gel, and in ion exchange DEAEeSephadex and CM-cellulose, are frequently used materials for GA purification. In affinity chromatographic separation of GA the use of b-cyclodextrinechitosan, amylopectin, alginate, acarbose, etc., is common practice. Other methods such as two-step extraction, ionic liquid extraction, liquideliquid extraction, and biphasic separation are also successfully used to purify GA. Details of the methods and materials used for purification in the past few decades are described in Table 2.2.

2.4.2

Characterization

Enzymes are amphoteric molecules containing a large number of acidic and basic groups. Charge on these groups varies with pH according to their acid dissociation constant. Changes in pH affect the activity and structural stability of the enzyme. GA is versatile with respect to pH and has been found active in the pH range 3e11. Negi and Banerjee found GA produced from A. awamori stable at pH 3e9 with optimum pH 4.5 [8]. Amirul et al. reported GA produced by A. niger stable in pH range 3.5e9.0 [90]. A similar pH range for GA from Aspergillus sp. has been reported by others also [91,92]. The thermostability of GA determines its industrial utility. It is directly related to its half-life period (t1/2) at different temperatures. Half-life is defined as the reaction time for the enzyme activity to drop to exactly half the initial activity under optimum conditions. The variation in t1/2 with temperature gives a measure of the temperature

Table 2.2

Details of Extraction and Purification Processes

Extraction and Purification Processes Ultrafiltration, ammonium sulfate precipitation (80%), DEAEeSephacel column chromatography Imarsil, activated charcoal, and Sephadex G-100 chromatography Ammonium sulfate precipitation (80%), Sephadex G-200 gel filtration

DEAEecellulose ion exchange and Sephadex G-100 gel-filtration chromatography Procion Blue H-ERD dye affinity chromatographic separation by elution with a borate solution Ammonium sulfate precipitation (70%), Sephacryl S-200 column gel filtration and S-Sepharose FF cation-exchange and Q Sepharose FF anion-exchange column chromatography Q Sepharose anion-exchange column chromatography, hydrophobic interaction (phenyl Sepharose column) on FPLC Ammonium sulfate precipitation (30%), HiLoad-Q Sepharose column, and monoQ column on FPLC anion-exchange, gel-filtration, hydrophobic interaction (phenyl Superose column) chromatography Sephadex G-75 gel filtration Ammonium sulfate precipitation, Q Sepharose anion-exchange column chromatography, HiLoad 16/60 Sephacryl S-200 gel filtration, and hydrophobic interaction column containing phenyl Sepharose. PM-10 ultrafiltration membrane and an Ultra-Free centrifugal filter device, DEAE Toyopearl 650S column

Optimal Conditions

Yield of GA (%)

References

Aspergillus niger

pH 5, 70 C

54

[58]

Rhizopus oligosporus SK5 mutant A. niger

pH 5, 80 C

57

[59]

pH 8, 40 C pH 6.5, 30 C pH 4.6, 30 C

0.09

[60]

140

[61]

pH 6, 65 C pH 4.5, 75 C

25

[62] [63] [64]

10% glycerol, 40 C pH 4, 60 C

51.9

[8]

9.2 52

[65] [66]

A. niger CCUG 33991 A. niger 2316 Aspergillus flavus HBF34 Rhizopus microsporus var chinensis Aspergillus awamori Paecilomyces variotii A. niger Curvularia lunata

pH 4, 50 C

17.8

[67]

Humicola spp.

pH 4.7, 55 C

33

[68]

Fusarium solani

pH 4.5, 40 C

31.8

[69]

Aureobasidium pullulans N13d Thermomyces lanuginosus

pH 4.5, 60 C

58

[70]

Fomitopsis palustris

pH 5, 70 C

[71]

[72]

33

Continued


Reverse micellar organic phase extraction (by using anionic surfactant, in n-heptane as a solvent) Ni2þ-NTA affinity chromatography Starch affinity chromatography Ammonium sulfate precipitation, aqueous two-phase systems, DEAE-650M chromatography, and Bio-Rad Prep Cell Acetone precipitation, ion exchange, gel filtration

Microbial Source

Details of Extraction and Purification Processesdcont’d

Extraction and Purification Processes Ammonium sulfate precipitation (90%), DEAEeSepharose anion exchange, phenyl Sepharose gel filtration, hydrophobic interaction chromatography ButyleSepharose and Superdex 200 HR gel permeation Macroaffinity ligand-facilitated three-phase partitioning using alginate Lyophilization, acetone precipitation, SP-Sepharose anion-exchange and Sephadex G-50 gel filtration Ultrafiltration and Sephadex G-25, Superdex HiLoad-200 gel filtration Ultrafiltration and Sephadex G-25, Superdex HiLoad-200 gel filtration Ultrafiltration and Sephadex G-25, Superdex HiLoad-200 gel filtration Affinity chromatography Ultrafiltration and Q Sepharose ion exchange Ammonium sulfate precipitation, Sepharose CL-6B, DEAEeSepharose Fast Flow, Q Sepharose Fast Flow, and Superose 12 gel filtration, ultrafiltration Q Sepharose ion exchange, Sephadex G-100 and phenyl Sepharose CL-4B gel filtration Ultrafiltration, DEAEecellulose, CMecellulose, Sepharose-6B column chromatography Affinity precipitation with alginate FPLC using a Bio Sil-SEC-400 filtration column, DEAEecellulose, CMecellulose Ammonium sulfate precipitation (75%), DEAEeSephadex A-50, ion exchange and Sephadex G-25 and G-100 gel filtration Two-step extraction system by using poly(ethylene glycol) and potassium phosphate Mono-Q ion exchanger and Superose-12 gel filtration DEAEecellulose ion-exchange chromatography and concanavalin AeSepharose gel chromatography Protamine sulfate treatment, ammonium sulfate precipitation, gel filtration (Sephadex G-75 sf, Ultrogel AcA 54), DEAEeSephacel chromatography, hydroxyapatite chromatography, and affinity chromatography on acarboseeAHSepharose 4B Acarbose (BAY G-5421) affinity chromatography FPLC, fast protein liquid chromatography; GA, glucoamylase.

Microbial Source Chaetomium thermophilum Sulfolobus solfataricus A. niger Thermomucor indicaeseudaticae Thermoplasma acidophilum Picrophilus torridus Picrophilus oshimae Thermobacterium thermosaccharolyticum A. niger Bo-1 T. lanuginosus

Optimal Conditions

Yield of GA (%)

References

pH 4, 65 C

3.57

[73]

pH 5.5e6, 90 C

4.6 83 1.03

[74] [75] [76]

pH 7, 60 C pH 2, 90 C

[77]

pH 2, 90 C pH 2, 90 C pH 5, 55 C

[77] [77] [78]

pH 4.4e5.6, 70 C 23

[79] [47]

T. lanuginosus F1

pH 6, 70 C

[81]

Scytalidium thermophilum

pH 5.5, 70 C

51

[82] [80] [83] [84]

A. niger, Bacillus amyloliquefaciens S. thermophilum Arthrobotrys amerospora ATCC 34468 A. awamori NRRL 3112

pH 6.5, 60 C pH 5.6, 55 C

81 78 41.6 3.2

pH 4.2, 60 C

100

[85]

Lactobacillus amylovorus Myrothecium strain M1

pH 6, 45 C pH 4, 70 C

21

[86] [87]

Candida antarctica CBS 6678

pH 4.2, 57 C

A. niger, Rhizopus sp.

[88]

80

[89]


Table 2.2


35

stability of the enzyme under actual reaction conditions and allows easy and quantitative comparison of enzyme stability [93]. This provides a way of selecting the most efficient enzyme from a group of enzymes and, hence, gives valuable input for suggesting a suitable enzyme for an industrial process. Negi and Banerjee (2009) observed that GA produced from A. awamori was stable in the temperature range 25e70 C. The highest amylolytic activity was obtained around 70 C; beyond that there was a fall in activity. This happens because, like any other chemical reaction, the rate of catalytic reaction of an enzyme increases with rising temperature, but at higher temperatures denaturation of the enzyme also takes place [8]. Negi et al. reported the half-life period for GA as 210, 120, 60, and 35 min at 50, 60, 70, and 80 C, respectively [17]. Nguyen et al. reported a thermophilic amylase of T. lanuginosus with half-life times longer than 1 day at 60 C [47]. The MichaeliseMenten constant (Km) is another important kinetic parameter and is defined as the concentration of substrate that gives half-maximal velocity (Vmax). Km and Vmax are significant in the biochemical characterization of an enzyme. The more firmly the enzyme binds to its substrate the smaller will be the value of Km. Moreover, Km is independent of enzyme concentration and is a true characteristic of the enzyme under defined conditions of temperature, pH, etc.; and, thus, it can be used as a genetic marker to identify a particular enzyme protein. Normally, higher Vmax and lower Km are the two desirable conditions for efficient enzyme hydrolysis. Negi and Banerjee reported Km and Vmax for GA produced from A. awamori as 9.8 mg/mL and 56.2 mg/mL min, respectively [94]. Nguyen et al. reported Km and Vmax of a-amylase from T. lanuginosus on soluble starch as 0.68 mg/mL and 45.19 U/mg, respectively [47]. For industrial use, the stability of an enzyme in the presence of metal salts is very important. In fact more than 75% of enzymes require metal ions to express their full catalytic activities. The optimum concentration of metal ions can be used to enhance the catalytic activity of an enzyme in that metal ions act as cofactors of many enzymes, but a high concentration of metal ions generally causes denaturation of the enzyme [95]. Negi and Banerjee (2009) reported enhanced activity of GA produced by A. awamori in low amounts of Ca2þ, Co2þ, Cu2þ, Fe3þ, Mg2þ, Zn2þ, and Hg2þ, excepting MnCl2 [94]. Chen et al. [73] also observed that Ca2þ, Mg2þ, Naþ, and Kþ enhanced the GA activity of Chaetomium thermophilum, whereas Fe2þ, Agþ, and Hg2þ inhibited it, and similar results were also observed by Tunga et al. [96], Quang et al. [47], and Spinelli (1996) [97]. Surfactant also influences enzyme activity in several ways. Negi and Banerjee reported an increase in GA activity in the presence of low concentrations (0.03% w/v) of Sodium Lauryl Sulfate and Triton X-100, whereas Tween 40, Tween 60, and Tween 80 inhibited the activity [94]. Negi and Banerjee (2010) [98] studied changes in the patterns or motifs of secondary structures of GA from A. awamori in the presence of denaturants such as urea and guanidine-HCl (Gdn-HCl) and at different pH through circular dichroism (CD) spectroscopy. They reported that 0.5 M concentration of urea increased GA activity and at 6 M concentration of urea GA loses its activity. CD spectra of GA in the presence of GdnHCl also followed a pattern similar to urea. At lower concentration (0.1 M) of Gdn-HCl the negative peak shifted from 208 to 219 nm to a very sharp peak at 198 nm with lower


intensity than the control, and in 3 M Gdn-HCl the spectrum was totally disrupted. Selvakumar et al. reported four different forms of GA produced by A. niger, i.e., GA I, GA I0 , GA II, and GA III, having apparent molecular masses of 112.0, 104.0, 74.0, and 61.0 kDa, respectively [91].

2.5 Enzyme Assay Enzyme assay is carried out to confirm the presence of the targeted enzyme in the source organism and to evaluate the enzyme activity in the reaction mixture. Most of the substances show absorption in a UV range that can be detected by a spectrometer. Spectrophotometric assay is the commonly used technique for observing the enzyme reaction with the mixture. Photometric assays are preferred because they are easy to perform and less susceptible to disturbances [99]. Enzyme assay is carried out at a temperature in the stable zone of the enzyme activityetemperature characteristic curve. Assay of thermophilic enzymes is carried out at higher temperatures, at which their activity is stable [99]. The isolates are screened for starch-hydrolyzing ability. Fungal isolates are normally inoculated on starch potato dextrose agar (PDA) plates. Ominyi et al. [6] inoculated fungal isolates on a 1% starch PDA plate and after 3e4 days of fungal growth the plates were flooded with iodine solution. A dark blue starcheiodine complex covered the entire agar because of the reaction of starch with iodine. Clear zones surrounding streaked lines were seen when the starch was broken down into sugars, indicating starch hydrolysis [100]. Units of enzyme activity: Enzyme activity is the measurement of the amount of desired product converted from substrate per unit time per unit of enzyme. The SI unit of enzyme activity is the “katal,” which is defined as moles (product) per second. One katal is a very large unit for practical purpose, hence, micromoles per minute (mmol/min) is used to express enzyme activity. In fact, the more commonly used unit of enzyme activity is IU (International Unit), given by the Nomenclature Committee of the International Union of Biochemistry (1982). IU is defined as the amount of enzyme required to catalyze conversion of 1 mmol/min of substrate. The parameter observed to determine the rate of substrate conversion is the change in concentration of the assay; hence, assay volume is also measured to determine the enzyme unit from the rate of change in concentration. Lakshmi and Jyothi (2014) used A. oryzae to produce GA using wheat bran as substrate. GA activity was determined by incubating the reaction mixture consisting of enzyme with 1% soluble starch solution in 50 mM citrate buffer (pH 5.5) at 50 C for 20 min [38]. The liberated glucose was measured with 3,5-dinitrosalicyclic acid reagent using glucose as a standard [101]. The GA activity unit (U) was expressed as the amount of enzyme releasing 1 mol of glucose equivalent per minute per milliliter. Keera et al. used A. oryzae to produce GA by SSF on corn flour [37]. The dinitrosalicylic acid method was used to determine the GA activity by measuring released reducing sugars using glucose as a standard. Enzyme assay was performed at 50 C in 1.0% (w/v) soluble starch


37

in 50 mM phosphate buffer at pH 5.0. An enzyme unit (U) was defined as the amount of enzyme that released 1 mmol of reducing sugars per minute, and enzyme activity was given in terms of units per gram dry original substrate (U/g). Nguyen et al. (2002) used T. lanuginosus for production of GA on starch-based medium. For GA assay, 1 mL reaction mixture containing 0.25 mL 0.1 M sodium acetate buffer, pH 4.6, and 0.25 mL 1% (w/v) soluble starch solution were preincubated at 50 C for 10 min. Incubation was continued for a further 15 min after addition of the appropriately diluted culture filtrate (0.5 mL) [47]. The reaction was terminated by placing the tubes in a boiling bath for 30 min and then allowed to cool. Glucose concentration was estimated by the glucose oxidase/peroxidase method using a standard glucose curve prepared under the same conditions [102]. A TtcGA from T. tengcongensis MB4 was successfully expressed in E. coli by Zheng et al. [26]. They determined GA activity in 50 mM HOAceNaOAc buffer (pH 5.0) containing 2% (w/v) maltose. The reaction mixture was incubated at 75 C and samples were removed at various time durations and the reaction was stopped by boiling at 100 C for 3 min. Removal of denatured proteins was done by centrifugation at 12,000 g for 5 min and measurement of liberated glucose was done using a D-glucose kit. One unit of enzyme activity was defined as the amount of enzyme that liberates 1 mmol of glucose per minute under assay conditions. Maximum enzyme activity was found at 75 C and pH 5.0.

2.6 Strain Improvement Selection of a particular strain out of many microbial cultures is a very vital but tedious task, especially when a commercially competent enzyme yield is to be achieved. For example, a strain of A. niger is capable of producing a large number of different enzymes. Selection of a suitable strain for the required purpose depends upon a number of factors, such as type of fermentation process, nature of the substrate, nutrients, functional characteristics of the desired enzyme, environmental conditions, etc. Microbial strains isolated from the living organism are required to be screened, grown, and maintained in a suitable environment before they are utilized for fermentation. Selective media prepared by supplementing the base medium with other compounds help in the growth of the targeted microorganism while inhibiting growth of other organisms. GA produced from conventional methods of using the parent strain of the microorganism normally exhibits normal catalytic activity in a temperature range of 50e60 C and pH range of 4e7. Many industrial process conditions are much more stringent, and in them the GA produced from parent strains is ineffective. Moreover, the cost of the process increases when bringing the temperature, the pH, etc., of the process into the range of the GA. With new techniques of protein engineering, microbiology, and molecular biology, novel strains capable of producing GA with better thermostability, better pH range, and better catalytic activity can be synthesized from the parent strain. In


industrial microbiology, induced mutagenesis using physical or chemical mutagens followed by selection is one of the effective strain improvement techniques. Pavezzi et al. produced A. awamori GA expressed in Saccharomyces cerevisiae by SmF in starches from various sources [1]. A mutant GA with improved thermostability was produced by a mutagenic polymerase chain reaction. Gene mutation fashioned three amino acid alterations in the protein structure (Ser54 / Pro, Thr314 / Ala, and His415 / Tyr) leading to an increase in the thermostability of mutant GA. A 7 C increase in the optimum temperature and 3.6 kJ/mol increase in the free energy of thermoinactivation (DG) at 65 C, and 1.8 kJ/mol at 80 C, was exhibited by the mutant GA compared to parent GA. The mutant GA showed better activity on potato starch, better yield, and a half-life twice that of the parent GA at 65 C. Kumar and Satyanarayana used nitrous acid and g-irradiation (60Co) to obtain a mutant Thermomucor indicae-seudaticae for production of GA [14]. Nitrous acid treatment was carried out by mixing the spore suspension of T. indicae-seudaticae in a 0.07 M solution of NaNO2 prepared in acetate buffer (0.2 M, pH 4.5). The samples were diluted with phosphate buffer (0.1 M, pH 7.0) to stop the reaction. In the next phase of mutagenesis, spore suspensions were exposed to 60Co g-irradiation in a dose range of 0e150 KR. A mutant strain of T. indicae-seudaticae produced 1.8-fold higher GA, retaining all the functional properties of the parent strain. Riaz et al. investigated g-ray-mediated mutagenesis of A. niger for enhancing the production of GA [7]. Mutant GA was more efficient and more stable at temperatures higher than 60 C than the parent GA. A comparison of the properties of mutant and wild-type GAs after strain improvement is listed in Table 2.3.

2.7 Commercially Available Glucoamylases GA has a wide variety of applications in various food industries. It is used in saccharification processes of starch or dextrin to produce glucose, which is further utilized by many industries like the beverage and baking industries as a substrate for various products. Because of its high demand and applications in various fields, many industries are also taking interest in industrial production of GA. For commercial purposes GA has been traditionally produced by using filamentous fungi such as Aspergillus and Rhizopus spp. Both SSF and SmF techniques are involved in industrial production of GA. One of the commercial forms of GA is available on the market by the name of Enzeco Glucoamylase Powder RO. The source of this commercial enzyme is R. oryzae. It shows GA activity and also possesses a considerable amount of protease and amylase activity. Another example of a commercial GA is Enzeco Glucoamylase-L, a liquid GA derived from A. niger. The manufacturer of these enzymes is Enzyme Development Corporation, which is a New York-based company. Dextrozyme is among one of the commercially available GAs used in various fermentation processes and is produced by A. niger. This enzyme is produced by


Table 2.3

39

Comparison of Activities of Mutant and Wild-Type Glucoamylase

Source

Mutation

Effect on Mutant Type

Reference

Aspergillus niger

g-Ray-mediated mutagen

[7]

Aspergillus awamori

Mutagenic polymerase chain reaction

Thermomucor indicae-seudaticae

Using nitrous acid and g-irradiation

Sulfolobus solfataricus A. awamori

Gene cloning and expression in Escherichia coli Cys Ala246

A. awamori

Cys, chemical Gln400 modification to cysteine sulfonic acid Ala Ser411

Improved kinetic properties, Kcat ¼ 343 and 727/s, Km ¼ 0.25 and 0.16 mg/mL, Kcat/Km ¼ 1374 and 4510 mg/mL s, for mutant and parent enzyme, respectively Better yield and thermal stability at 65 C with increased enzyme activity Improved productivity, i.e., >23 U/mL, versus parent strain (18 U/mL) and higher specific growth rate than parent strain Extremely thermostable GA having optimum temperature of 90 C Thermal stability increased (stable at 66 C) Activity increased to 160% of wild type Specific activity increased by 32% compared to wild type Mutant GA showed increase in specific activity from 45 to 67.5 C. Yield increased (by 100-fold)

A. awamori A. awamori Pichia pastoris

P. pastoris A. awamori

60

Co

Disulfide bonds introduced through protein engineering Cloning and expression of catalytic domain of GA from A. awamori Heterologous expression, vector used was pHIL-D2 Gly137 Ala

Aspergillus candidus Link var aureus

Chemical mutagenesis using MNNG

A. awamori var kawachi

Protease-less mutant

Higher thermal stability and yield increases 10% higher activity than wild type, i.e., 24.5 IU/mg Mutant produces 111 U/mL GA compared to 50 U/mL GA produced from parent strain Higher GA activity

[1]

[14]

[74] [103] [103] [104] [105] [106]

[107] [108] [109]

[110]

GA, glucoamylase; MNNG, N-methyl-N0 -nitro-N-nitrosoguanidine.

Novozymes, located in Denmark. There are many industries present in China that are extensively involved in the production of the GAs, for example, DuPont Genencor Science, producing an enzyme by the name of Glucoamylase GA-L New from A. niger; Wuxi Syder Bioproducts Co., Ltd., which provides GA by the name of Syder Brand


Table 2.4

Some Commercially Available Glucoamylases [14]

Commercial Name

Source of Enzyme

Manufacturer

Glucoamylase GA-L New Amyloglucosidase A107823 Enzeco Glucoamylase-L Liquid/Solid Glucoamylase Syder Brand Glucoamylase Glucoamylase Sunson GA-L, GA KDN-GE01TM Dextrozyme DEXTRO 300L Glucozyme Af6

DuPont Genencor Science, Wuxi, China Aladdin Industrial Co., Ltd., Shanghai, China Enzyme development Corporation Sichuan Shan Ye Bio-Tech Co., Ltd. Wuxi Syder Bioproducts Co., Ltd. Sunson Industry Group Co., Ltd. Qingdao Continent Industry Co., Ltd. Novozymes (Denmark) Advanced Enzyme Technologies Ltd. Amano Enzymes USA Co., Ltd.

Glucoamylase YY0515

Aspergillus niger A. niger A. niger e e A. niger A. niger A. niger e A. niger Rhizopus niveus Rhizopus delemar A. niger

Amyloglucosidase A7420 Glucoamylase Liquid 25 KG/Drum

A. niger e

Shanghai Yuanye Biological Technology Co., Ltd., Shanghai, China Sigma Aldrich, St. Louis, MO, USA Jinzhu Tibet Co., Ltd.

e, information about the source of enzyme is not available.

Glucoamylase; and Glucoamylase YY0515 from A. niger by Shanghai Yuanye Biological Technology Co., Ltd., Shanghai, China. Some other examples of commercially available GAs and their manufacturers are listed in Table 2.4.

2.8 Conclusion and Perspective The huge commercial demand for GA makes even a small improvement in production and catalytic efficiency lucrative. There are continuous efforts to synthesize improved strains with a wide pH range and thermostability and better catalytic efficiency to suit commercial starch processing. Advances in molecular biology and protein and genetic engineering are being used to synthesize improved GA-producing strains to enhance the yield and functional properties of the enzyme. To reduce the cost of starch saccharification, GAs with better thermostability, better yield, and wider range of pH stability are being produced using advanced techniques such as site-directed mutagenesis, recombinant DNA technology, and cloning. Cheap biomass requiring minimal additional nutrients is being explored to reduce the production cost. There is a lot of room to further improve the strains, design functional properties of GA to suit particular industrial processes, and optimize production and purification of GA using new techniques and developments in the areas of microbiology, molecular biology, protein engineering, fermentation technology, bioreactors, etc., so that industrial applications using GA can become more economical.


41

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45

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Pectinolytic Enzymes

3

Héctor A. Ruiz*, Rosa M. Rodríguez-Jasso, Ayerim Hernandez-Almanza, Juan C. Contreras-Esquivel, Cristóbal N. Aguilar FOOD RESEARCH DE PARTMENT, SCHOOL OF CHEMISTRY, AUTONOMO US UNIVERSITY OF COAHUILA, SALTILLO, COAHUILA, MÉ XICO

3.1 Introduction Nowadays, the bioprocesses based on the utilization of renewable raw materials probably make a major contribution to sustainable development. However, it is not enough just to develop more eco-friendly processes; sustainability also requires economic and social justification [1,2]. According to Yang [3], a bioprocess consists mainly of raw material, pretreatment, fermentation, downstream processing, and purification. Furthermore, the actual bioprocess is basically dependent on the substrate and organisms used and the nature and applications of the final product [3]. In a specific case, the development of a bioprocess for traditional industrial enzymes production has progressed significantly. Additionally, a large number of industrial processes in the areas of food processing, beverage production, animal feed, leather, textiles, detergents, biofuel, pulp paper, fats/oils, and organic synthesis utilize enzymes [4,5]. Moreover, developments in biotechnology are yielding new applications for enzymes. Deswal et al. [5] mentioned that enzymes play an important role in improving productivity and the cost of product formation in the bioprocess. Pectinase is an important enzyme that finds application in many food-processing industries and the microbial pectinases have been reported to account for 25% of the global food enzymes sales, and majority of them are from fungal sources [6]. The pectinase enzymes are a complex system of proteins, which include hydrolases, lyases, and oxidases, that play important roles in the degradation or modification of pectin substances and are favorable for particular processes, for example, in the extraction and clarification of fruit juices and in wine production [7,8]. On the other hand, the production of pectinases is based on two bioprocesses, solid-state fermentation and submerged fermentation, and originates from fungi, especially Aspergillus niger. These days, the production of pectinase is done industrially and thus can be considered a consolidated bioprocess. This chapter focuses on the development of a consolidated bioprocess for pectinase production, and updated *



47


information is presented on pectic substances, pectinase classification, pectinase assays, pectinase production processes (raw materials, carbon sources, microorganisms, systems of production by solid-state and submerged fermentation, types of bioreactors), downstream processes, purification methods, and technoeconomic analysis of pectinase production.

3.2 Pectic Substances Pectic substances are galacturonic acid-containing polysaccharides assembled in the plant cell wall and middle lamella together with other structural biopolymers such as cellulose, hemicelluloses, proteins, and lignin. These acidic substances may be associated with the plant cell-wall components through physical, covalent, or ionic bonding and/or hydrogen bonds [9]. Water-soluble pectic substances are extracted with water, whereas covalent pectic substances are released into the aqueous medium by the use of cleaving agents such as chemical, enzymatic, or physical treatments. Chelating agents are used to obtain ionically bonded pectic substances. The main components of pectic substances are D-galacturonic acid with varying extent of methyl-esterified carboxyl groups. The other minor components are a variety of neutral sugars such as rhamnose, galactose, arabinose, apiose, fucose, glucose, mannose, and xylose and ulosonic acid derivatives. Ferulic acids are common residues, which are linked to the neutral sugars of side chains of sugar beet pectin [10]. The chemical structure of pectins mainly consists of galacturonans (homogalacturonan, rhamnogalacturonan II, and xylogalacturonan) and rhamnogalacturonan I [11]. Most pectic substances comprise straight “smooth” and ramified “hairy” regions [12]. The smooth regions consist of a linear homogalacturonan backbone, whereas the hairy regions consist of a rhamnogalacturonan backbone with side branches of arabinogalactan or galactan of varying length. Detailed information on the elaborate model of pectin structure can be found in several reviews [9,13,14].

3.3 Pectinase Classification Pectinases can be divided into depolymerizing and deesterifying enzymes following criteria such as cleavage type of glycoside linkages, reaction mechanisms (exo- or endotypes), and esterification degree. Furthermore, deesterifying enzymes are classified according to type of ester group (methyl, acetyl, or feruloyl ester). Deesterifying enzymes comprise pectin methylesterase, pectin acetylesterase, ferulic acid esterase, and rhamnogalacturonan acetylesterase. Other pectinases are the oxidases (i.e., galacturonic acid oxidase) and those responsible for the metabolism of degradation products of pectin. Such classification refers to the mode of action of pectinases on water-soluble substrates. Table 3.1 shows a general classification of pectinases according to the International Commission of Enzymes. Traditionally breaking down of pectic substances was done through the use of well-known pectic enzymes (pectin methylesterase, polygalacturonase, pectin lyase, etc.) able to degrade only the smooth regions [11]. Pectic enzymes occur in higher plants and are produced by microorganisms. Several new enzymes have now been reported,

Chapter 3 Pectinolytic Enzymes

Table 3.1

Enzyme

49

Modern Classification of Pectic Enzymes

Common Name

EC Code

Glycosyl Hydrolase Family

Main Products or Action Mode

Hydrolases Esterases

Glucosidases

Pectin methylesterase Pectin acetylesterase Rhamnogalacturonan acetylesterase Feruloyl esterase Endopolygalacturonase Exopolygalacturonase Rhamnogalacturonan hydrolase Rhamnogalacturonan galacturonic hydrolase Rhamnogalacturonan rhamnohydrolase Xylogalacturonan hydrolase Endogalactanase a-Galactosidase b-Galactosidase Endoarabinase a-L-arabinofuranosidase

3.1.1.11 3.1.1.6

Pectin lyase Pectate lyase Rhamnogalacturonan lyase

4.2.2.10 4.2.2.2

8 12

RG þ ferulic acid Oligogalacturonates þ galacturonic acid Galacturonic acid, digalacturonate Random hydrolysis of linkages between galacturonic acid and rhamnose on RG Random hydrolysis of linkages between galacturonic acid and rhamnose on RG Release end rhamnose linked to galacturonic acid on RG (exohydrolysis) Release xylose from XG

3.2.1.15 3.2.1.67

3.2.1.22 3.2.1.23

Low methoxyl HG þ methanol HG deacetylated þ acetic acid RG deacetylated þ acetic acid

53 27, 36 35 51, 54

Remove residues of galactose Remove residues of galactose Random hydrolysis of arabans Remove residues of arabinose

Lyases Unsaturated methyloligogalacturonates Unsaturated oligogalacturonates Random hydrolysis of RG

Oxidases Galacturonic oxidase

Oxidation of galacturonic acid

HG, homogalacturonan; RG, rhamnogalacturonan; XG, xylogalacturonan.

which degrade parts of the hairy region, such as rhamnogalacturonan hydrolase, rhamnogalacturonan lyase, etc. [12,15]. Based on the chemical structure of pectic substances, pectinases are classified by the structural region that cleaves along the pectin molecule. Nomenclature, properties, and characteristics of classical and new pectic enzymes are discussed in several reviews [14]. In the past pectinases were described only as those enzymes that acted on homogalacturonan chains. However, several enzymes that act on rhamnogalacturonan I chains have now been described.

3.4 Pectinase Assays Depolymerizing pectinases release reducing sugars. Several colorimetric methods have been used to evaluate the reducing sugars generated by cleavage of pectic


polysaccharides based on the reduction of an oxidation agent by the reducing sugars [16]. The most frequently used methods to estimate the generated reducing sugars from polysaccharides are the colorimetric NelsoneSomogyi and dinitrosalicylic acid procedures [16]. The reducing-sugar colorimetric assays have been adapted to microassays using microtiter plates [16]. A new colorimetric assay for quantifying endopectinase utilizes ruthenium red as a detection reagent in a liquid medium reaction [17]. Rapid screening methods to assay pectinolytic depolymerizing activities in a cupplate or microplate reader are available by using chromogenic pectic substances. Table 3.2 shows the insoluble and insoluble chromogenic pectic polysaccharides used to detect specific enzymatic activities such as pectinases, rhamnogalacturonan-degrading enzymes, and/or accessory pectinolytic enzymes (arabinases and/or galactanases). Early studies of chromogenic pectic polysaccharides were developed with pectins for assaying total endopectinase activity [18,19]. Dalboge et al. [20] used chromogenicspecific pectic polysaccharides for screening and expression cloning of fungal enzyme genes of industrial relevance. Visualization of pectic enzymes with specific soluble chromogenic substrates in gel electrophoresis or isoelectrofocusing has been described [15]. Table 3.3 shows a range of water-soluble polysaccharides for pectinolytic assay with specific pectic polysaccharides. The pectin methylesterase assay comprises evaluation of released methanol from high-methoxylated pectin by colorimetric or chromatographic methods [21]. Another common practice is evaluating the generated carboxylic acid from methoxylated pectin by manual or automatic alkaline titration. Continuous spectrophotometric assays for pectin methylesterase activity are also available [22,23]. Pectin acetylesterase [24] and rhamnogalacturonan acetylesterase activities are indirectly evaluated after analysis of released acetic acid by colorimetric or chromatographic methods [25]. A wide variety of cup-plate and zymogram methods are offered for assay pectin methylesterase [26,27].

Table 3.2 Soluble and Insoluble Chromogenic Pectic Polysaccharides for Assay of Pectinolytic Enzymes Name

Water Solubility

Enzyme

Manufacturer/References

Azo-galactan (potato) Azo-galactan (potato) Azo-rhamnogalacturonan (soy) DISANED pectina Red arabinan (sugar beet) Azurine arabinan Azurine galactan Azurine rhamnogalacturonan

Soluble Soluble Soluble Soluble Soluble Insoluble Insoluble Insoluble

Endogalactanase Endogalactanase Endorhamnogalacturonase Endopectinase Endoarabinase Endoarabinase Endogalactase Endorhamnogalacturonase

Megazyme, Ireland SigmaeAldrich, USA Megazyme, Ireland Friend and Chang [18] Megazyme, Ireland Megazyme, Ireland Megazyme, Ireland Megazyme, Ireland

a

Azo dye N-[1-[4-[(3,6-disulfo-l-naphthyl)azo]- naphthyl]]ethylenediamine.


Table 3.3

51

Water-Soluble Nonchromogenic Pectic Substrates

Substrate

Characteristics

Suitable for Pectinase

Seller

Pectin (citrus/apple)

High methoxylated

SigmaeAldrich

Pectin (citrus/apple)

Low methoxylated

Pectin (sugar beet) Polygalacturonic acid (citrus)

Acetylated Nonmethoxylated

Rhamnogalacturonan I (soy) Rhamnogalacturonan I (potato)

Free arabinogalactan Free arabinogalactan

Pectin methylesterase Polygalacturonase Pectin lyase Polygalacturonase Pectate lyase Pectin acetylesterase Polygalacturonase Pectate lyase Rhamnogalacturonan-degrading enzymes Rhamnogalacturonan-degrading enzymes

Megazyme

Megazyme Megazyme Megazyme

3.5 Pectinase Production Processes The production of pectic enzymes is in high demand at the industrial level to satisfy a lot of determined requirements. Great amounts of agro-industrial waste rich in polysaccharides, such as pectic substances, are produced worldwide. Some of these wastes are used for the production of pectin. Purified pectic enzymes have been proposed for extracting and modifying pectin. Currently, pectin is extracted at the industrial scale by physicochemical methods, but new biotechnological (microbial and enzymatic) alternatives are being developed. Pectinases are produced by several microorganisms; particularly, fungal strains are known as good producers. Fungi such as Aspergillus and Penicillium are also used for pectinase production at the industrial scale, which is carried out mainly by submerged fermentation (SmF), although solid-state fermentation (SSF) is also used. A list of several bacteria and fungi used to produce pectic enzymes is shown in Table 3.4. For most microbial bioprocesses used for pectinase enzyme production, the use of pectin-rich materials is a common practice because such materials act as inducers of the pectinolytic complex. However, it is well known that in the presence of high sugar concentration the expression of associated genes is catabolically repressed; and the fedbatch system is an excellent alternative to avoid this biochemical phenomenon [51]. Table 3.5 summarizes recent advances in pectic enzyme production, showing the new microorganisms used as sources of these kinds of biocatalysts, the fermentation system evaluated, the most popular bioreactors, and the most common substrates.

3.5.1

Raw Materials for Pectinase Production

Common pectinase enzymes have been produced for more than 3 decades and can be obtained by either SmF or SSF [87e89]. Fungal pectic enzymes are possibly the most used commercially, and for their production, pectin-rich materials are used as the source of needed inducers of these biocatalysts. These materials can be agricultural by-products


Table 3.4

Microorganisms Producing Pectic Enzymes

Microorganism

PME

PGase

e-PPLase

PLase

References

Bacteria Bacillus subtilis Bacteroides thetaiotaomicron Erwinia chrysanthemi Lachnospira multipara Thermoanaerobacter italicus

Nasser et al. [28] McCarthy et al. [29] Shevchik et al. [30] Silley [31] Kozianowski et al. [32]

Fungi Aspergillus aculeatus Aspergillus alliaceus Aspergillus kawachii Aspergillus niger Aspergillus oryzae Colletotrichum lindemuthianum Corticium rolfsii Fusarium oxysporum Fusarium solani Kluyveromyces marxianus Penicillium capsulatum Penicillium frequentans Rhizoctonia solani Rhizopus stolonifer Saccharomyces cerevisiae Saccharomyces fragilis Sclerotinia sclerotiorum Fusarium oxysporum

Foda et al. [33] Sreenath et al. [34] Contreras-Esquivel [35] Kester and Visser [36] Kitamoto et al. [37] Wijesundera et al. [38] Tagawa and Kaji [39] Huertas-González et al. [40] Guo et al. [41] Sakai et al. [42] Gillespie et al. [43] De Fatima-Borin et al. [44] Bugbee [45] Manachini et al. [46] Gainvors et al. [47] Lim et al. [48] Oliva et al. [49] di Pietro and Roncero [50]

e-PPLase, endo-pectate lyase; PGase, polygalacturonase; PLase, pectate lyase; PME, pectin methylesterase.

such as cassava fibrous waste, wheat bran, apple pomace, corn barn, citrus waste, coffee pulp, sugarcane bagasse, and raw starch from cassava tuber [7,90e99].

3.5.2

Submerged Fermentation

At the industrial level, SmF is carried out at volumes from 20 to several hundred cubic meters. Because most of the microorganisms used for pectinase production are aerobic, air must be supplied at rates from 0.1 to 2.0 vvm. Oxygen transfer from the gas phase to the liquid phase is enhanced by mechanical or air-lift agitation of the culture medium, and agitation helps to maintain homogeneous conditions of pH, temperature, and dissolved oxygen in the broth. Zetelaki-Horvath and Vas [100] reported that pectic enzyme production by a mutant strain of A. niger growing in a culture medium with sugar beet slices supplemented with malt extract and mineral salts was strongly influenced by the oxygen uptake rate. Whereas growth was stimulated at an oxygen uptake rate of 100 mmol/L/h, pectin

Table 3.5 Summary of Advances in Pectinase Enzyme Production by Solid-State Fermentation and Submerged Fermentation Culture System

Microorganism

PGase and PME

Aspergillus niger

PGase

Trichoderma reesei Rut C-30

SmF and SSF SSF

Thermostable alkaline pectinase Endo-PGase

Bacillus sp.

SSF

Paecilomyces clavisporus

SmF and SSF

Endo-PGase PGase

A. niger Sporotrichum thermophile

SSF SmF

NR

Exo- and endo-PGase

A. niger

SSF

Alkaline pectinase

Streptomyces

PME PME

Aspergillus flavus Cryptococcus cylindricus and Mrakia frigida Bacillus gibsonii S-2

SmF and SSF SmF SmF

Erlenmeyer flask Erlenmeyer flask NR NR

SSF

Penicillium viridicatum RFC3

SmF

PGase Endo- and exo-PGase and PLase

Bioreactor

Substrate

References

Flask

Apple pectin and rice bran

Fawole and Odunfa [52]

Bioreactors manufactured at home Erlenmeyer flask Erlenmeyer flask

Beetroot

Olsson et al. [53]

Wheat bran, rice bran, and pomace Manachini solution (0.9% K2HPO4, 0.1% (NH4)2SO4, 0.01% MgSO4 7H2O, 0.09% and 0.1% yeast extract) and wheat bran with citrus pectin Citrus peel Yeast extract and citrus pectin Sugar beet pulp

Raj-Kashyap et al. [54]

Erlenmeyer flask Erlenmeyer flask and polypropylene boxes 20 30 cm

Souza et al. [55]

Dhillon et al. [56] Kaur et al. [57]


Pectic Enzyme

Bai et al. [58]

Horokoshi medium and wheat bran Agarose with pectin Pectin

Kuhad et al. [59] Mellon et al. [60] Nakagawa et al. [61]

Sugar beet pulp

Li et al. [62]

Orange bagasse and wheat bran

Silva et al. [63]

53

Continued

Culture System

Pectic Enzyme

Microorganism

PGase

Paenibacillus sp. BP-23 and Bacillus sp. BP-7

SmF

Endo-PGase

SSF

Exo- and endo-PGase

Kluyveromyces marxianus CCT3171 Kluyveromyces wickerhamii and Stephanoascus smithiae A. niger DMF27

Exo- and endo-PGase

PGase

Bioreactor

Packed-bed reactor

SmF SmF and SSF

Erlenmeyer flask and glass bottles

A. niger

SmF and SSF

Erlenmeyer flask and glass containers

PGase

A. niger

SSF

Petri dishes

PGase

A. niger

SSF

Xylanase and PGase Xylanase and thermostable alkalophilic pectinase PGase

Aspergillus awamori Bacillus subtilis and Bacillus pumilus

SSF SSF

Erlenmeyer flask Petri dishes Erlenmeyer flask

Aspergillus sojae ATCC 20235

SSF

PGase

A. niger

SSF

Erlenmeyer flask Erlenmeyer flask

Substrate

References

Nutritive broth, 1% glucose, polygalacturonic acid or citrus pectin Porous silicate glass (glucose) and cellulosic support Glucose

Soriano et al. [64]

Almeida et al. [65] Silva et al. [66] Patil et al. [67]

(NH4)2SO4, 0.1%; MgSO4 7H2O, 0.5%; KH2PO4, 0.5%; FeSO4 7H2O, 0.0005%, and 8 g sunflower head powder (NH4)2SO4, 0.1%; MgSO4 7H2O, 0.5%; KH2PO4, 0.5%; FeSO4 7H2O, 0.0005%, and lemon peel, sorghum, and sunflower powder Wheat bran and rice dextrose Orange peel

Nighojkar et al. [70]

Grape marc Wheat bran

Botella et al. [71] Ahlawat et al. [72]

Corn (crushed, flour corn and cob corn) Wheat bran, beet bagasse, and minerals

Ustok et al. [73]

Patil and Dayanand [68]

Debing et al. [69]

Dinu et al. [74]


Table 3.5 Summary of Advances in Pectinase Enzyme Production by Solid-State Fermentation and Submerged Fermentationdcont’d

Fusarium oxysporum, A. niger, Neurospora crassa, Penicillium decumbens B. subtilis RCK

SSF

Erlenmeyer flask

Orange peel

Mamma et al. [75]

SSF

Wheat bran

Gupta et al. [76]

Aspergillus foetidus

SSF

Erlenmeyer flask NR

Taskin and Eltem [77]

Debaryomyces nepalensis Aspergillus oryzae

SmF SmF

Beetroot pulp and wheat bran Galactose and lemon peel Wheat bran, citrus pectin

Exo-PGase

Thermomucor indicaeseudaticae N31

SmF and SSF

PGase

A. niger

SSF

PGase

Pseudozyma sp. SPJ

SSF

Polymethylgalacturonase

Aspergillus spp.

SSF

PGase

A. niger

SSF

PLase

SSF

PGase

Oidiodendron echinulatum MTCC 1356 A. niger LFP-1

SSF

PGase

A. niger Aa-20

SSF

Endoglucanase, xylanase, PGase Exo-PGase PGase and polymethylgalacturonase PLase and e-PPLase PGase

Batch Stirred tank and airlift Erlenmeyer flask Horizontal drum Erlenmeyer flask Erlenmeyer flask NR

Martin et al. [80]

Citrus pectin, wheat bran, sugarcane bagasse, and orange bagasse Citrus peel

Rodríguez-Fernández et al. [81]

Citrus peel

Sharma et al. [82]

Wheat bran and orange peel

Heerd et al. [83]

Wheat bran and mosambi peel Wheat bran, sugarcane bagasse Pomelo peel

Khan et al. [84]

Darah et al. [86]

Peel pomace

Ruiz et al. [7]

Yadav et al. [85]

e-PPLase, endopecatelyan pectate lyase; PGase, polygalacturonase; PLase, pectate lyase; PME, pectin methyl esterase; SSF, solid-state fermentation; SmF, submerged fermentation.


Erlenmeyer flask Erlenmeyer flask Column-tray bioreactor

Gummandi and Kumar [78] Fontana et al. [79]

55


esterase, endopolygalacturonase, and pectin lyase production was stimulated at oxygen uptake rates of 13, 49, and 60 mmol/L/h, respectively. However, the macerating activity, due to pectin esterase and endopolygalacturonase activities, gave two maximal values at 12 and 14 mmol/L/h. On the other hand, some yeast strains such as Kluyveromyces marxianus produced nine isoenzyme forms of endopolygalacturonase when growing under anaerobic conditions with glucose as the sole carbon source [101]. Most of the pectinases are inducible enzymes that require the presence of an inducer to be synthesized. Although pectin is the natural inducer for pectinase production, its elevated cost makes it difficult to use at the industrial level. A number of agricultural products containing pectin and other polysaccharides have been used for pectinase production [7,102,103]. Depending on the raw material used as the source of inducer and carbon, the culture medium needs to be supplemented with minerals to improve the microbial growth and enzyme production. Ammonium sulfate is widely used as a nitrogen source. Macroelements such as P, K, and Mg are generally supplemented as KH2PO4 and MgSO4 and microelements such as Mo, Zn, Fe, Mn, and Co are added as mineral salts at low concentrations (below 0.05%, w/w). A comparative study of polygalacturonase production by A. niger and Penicillium dierckxii from different pectin sources showed that sugar beet pectin (at 10 g/L) was the most active inducer and ammonium sulfate the best source of nitrogen for polygalacturonase production by both strains [104]. The use of a mixture of sugar beet pulp and alkaline-extracted sugar beet pulp instead of sugar beet pulp alone slightly increased the polygalacturonase production by Trichoderma reesei [53]. As stated above, although pectinase production by fungi has been mostly inducible, a constitutive exopectinase was produced by Aspergillus sp. CH-Y-1043 grown on glucose, sucrose, fructose, glycerol, and galacturonic acid [51]. Pectinase production by Aspergillus sp. and Neurospora crassa was induced by pectin and repressed by glucose and by the degradation products of pectin [105,106]. In contrast, Erwinia carotovora produced a pectic acid lyase that was induced by the breakdown products of pectic acid [107]. The complexities of the regulatory mechanisms involved in pectinase production require permanent programs for the selection of catabolic-resistant strains. However, technological aspects such as fed-batch cultures or SSF can be used to minimize the catabolic repression by glucose or by the breakdown products of pectin [105,108].

3.5.3

Solid-State Fermentation

SSF used for the production of polygalacturonases by A. niger with sugarcane bagasse as solid support showed that endopolygalacturonase and exopolygalacturonase productivities were 18.8 and 4.5 times higher in SSF than in SmF [105]. Apparently, regulatory phenomena such as inductionerepression related to pectinase synthesis by A. niger are different in the two types of fermentation [105]. The use of sugarcane bagasse as a sole carbon source allowed higher production of pectin esterase and polygalacturonase by A. niger in SSF compared with that obtained in SmF. Moreover, glucose


57

addition improved pectinase production in SSF but it was decreased in SmF [96]. Similar results were obtained with polyurethane (as inert support) and a culture medium containing pectin as the carbon source, showing that protease production is lower in SSF than in SmF [108]. However, Morita and Fujio [109] compared the specific polygalacturonase activities from Rhizopus sp. MKU 18 in a metal-ion-regulated liquid medium and a solid wheat bran medium. Their work suggested that some advantages could be found in producing polygalacturonase in the metal-ion-regulated liquid medium. The use of sugar from cane bagasse as support, impregnated with a defined culture medium, showed that pectinases produced by SSF were more stable at higher pH and temperature than those produced by SmF [98]. Although pectinase production at low water activity (aw) values was lower than that obtained at high aw values, the specific activity increased up to 4.5 times in SSF [89]. Polygalacturonase production by thermophilic Thermoascus aurantiacus by SSF was optimized using a mixture of sugar from cane bagasse and orange bagasse at a ratio of 1:1 at pH 5, with substrate moisture of 70% and temperature of 50 C [110].

3.5.4

Bioreactors for Pectinase Production

Microbial production of pectic enzymes can be achieved in several kinds of bioreactors for SmF and SSF in which free and immobilized microbial cells can be used. However, for each culture system there are technological advantages and disadvantages associated with control of the bioprocess, environmental conditions, culture medium and microbial strain, etc. Similar to other types of biotechnological production, cultures for pectic enzymes can be carried out by using various cultivation modes (batch, fed-batch, repeated-batch, or continuous with or without biomass recycling) in various bioreactor types (stirred-tank bioreactor, packed-bed, fluid-bed, or various types of tubular and drum bioreactors). Most reports on pectic enzyme production using SmF include the use of shake-flasks, stirred tanks, or internal- and external-loop air-lift bioreactors; however, for this kind of cultivation few innovations have been reported in past years. On the other hand, there has been a renewed interest in SSF because of the high productivity of metabolites produced through this bioprocess [7,111]. The use of traditional trays, columns, bags, and horizontal tanks has been widely employed for the production of enzymes and other metabolites. Since 2000, very few, but innovative and promising, bioreactors have been reported, including the Growtek bioreactor, the column tray, the gas double-dynamic bioreactor, and a continuously mixed bioreactor [7,111e114]. Several modern alternatives for the design of novel bioprocesses can be applied to the production of pectic enzymes; among these are the use of cell immobilization, which offers numerous advantages over normal suspended cultures, such as cell stability, higher cell densities, enhanced fermentation productivity, and feasibility of continuous processing. The use of surface adhesion culture is another biotechnological strategy for the production of fungal enzymes.


In this chapter we describe the last contribution to the topic of pectinase production by Ruiz et al. [7]. They reported the design of an SSF process for the production of pectinase by A. niger Aa-20, using lemon peel pomace as the support and carbon source in a solid-state bioreactor. The SSF process was operated in a new column-tray bioreactor producing high levels of fungal pectinase activity, being described as a very promising process. Casciatori et al. [115] reported the use of orange pulp and peel for pectinase and phytase production by SSF in a fixed-bed bioreactor, demonstrating its hygroscopic properties as a support and that it can be used as a substrate in SSF for the biosynthesis of pectinase, controlling the microbial metabolism, which is strongly affected by the moisture content. This work therefore provides significant new information, particularly interesting for engineers dealing with design and scaling up of SSF bioreactors. Maciel et al. [116] reported polygalacturonase production in a fixed-bed reactor operated under various operational conditions: immobilized cells without aeration, immobilized cells with aeration, immobilized cells with aeration and added pectin, and free cells with aeration. They reported that the highest exo- and endopolygalacturonase activities were obtained with immobilized cells of A. niger URM 5162 without aeration, and the process allowed enzyme production in a fixed-bed bioreactor.

3.6 Downstream and Purification Methods The downstream stage of enzyme production is responsible for about 70% of the final cost of the product in industrial processes and is often identified as the process bottleneck; therefore, continuous scientific research has been focused on optimizing the unit operations to minimize the number of steps necessary to maximize the product recovery at a specified concentration and purity. The most relevant phases in downstream design that are needed to follow the upstream stage are recovery, isolation, and purification. However, enzyme isolation and purification require complex, multistep operations, using significant amounts of chemical and auxiliary material [117e119]. A brief description of recent studies and advances in downstream processing for pectinase purification is provided next.

3.6.1

Pectinase Enzyme Recovery

Earlier it was described that pectinases are principally produced by plants and by microbial sources, mainly from filamentous fungi, yeasts, and filamentous and nonfilamentous bacteria. For that reason the first step in the downstream process is the recovery of the upstream fraction containing the target enzyme. Actually, pectinases produced by microbial sources are principally expressed as extracellular biomolecules; for that reason the culture broth is separated from the biomass to eliminate the majority of the insoluble contaminants. For SmF the most common preliminary step for biomass elimination is


59

filtration. In contrast, because of the low water content of the SSF process, it is necessary to add distilled water or buffer over the solid support to obtain the enzymatic extract before the elimination of insoluble contaminants (biomass and support) by filtration.

3.6.2

Isolation and Concentration Methods

The pectinase crude extract solution obtained after the preliminary separation is commonly concentrated by the addition of saline solutions and dialysis and ultrafiltration procedures. These techniques have the advantages of being simple, rapid, and economical; however, it is necessary to optimize the isolation conditions to avoid protein denaturation and loss of enzymatic activity. The use of ultrafiltration techniques as the only concentration step has been demonstrated to be inefficient at eliminating contaminants able to cause possible enzymatic inhibition. Duque Jaramillo et al. [119] evaluated the production of Aspergillus oryzae pectinase by the fermentation of passion fruit peel. The obtained crude extract was concentrated by ultrafiltration with a 10-kDa cutoff-point membrane. The results showed that, although the concentrated product showed a high enzymatic activity, the final extract contained colorful pigments associated with passion fruit phenolic compounds that can affect the pectinase viability, especially in concentrated broth. Alkaline pectinase produced by Bacillus subtilis 7576 has been selectively isolated using centrifugation, ultrafiltration, and ammonium sulfate precipitation [120]. The recovery rate of crude enzymatic extract by ultrafiltration membrane (cutoff 10 kDa) was 73.3%, and the specific activity was 259.7 U/mg; after fractional salting out, the recovery rate was up to 62.9%, and the specific activity was 1084 U/mg. Moreover, ammonium sulfate fractionation was also used to concentrate an extracellular pectin lyase (PL) secreted by Fusarium decemcellulare MTCC 2079 under SSF and an exp-polygalacturonases (PG) from a soil isolate produced by SmF of Paecilomyces variotii. The procedure was followed by an overnight dialysis against water or buffer [121,122]. Li et al. [123] showed the relevant effect of applied ammonium sulfate saturation as the first step in the isolation of a Bacillus clausii PL; the total enzyme recovery was over than 57.6% and the specific activity increased from 6.6 to 15.2 U/mg. Silva et al. [124] reported the concentration of pectinolytic enzyme obtained from Penicillium viridicatum RFC by SSF using kaolin (hydrated aluminum silicate, Caulin) to remove the pigments and some of the protein with a short incubation time (10 min). The supernatant phase obtained after centrifugation was dialyzed and then concentrated by ultrafiltration (10-kDa cutoff). The kaolin treatment showed a successful effect on crude enzyme solution decolorization; 71.0% of the total protein was removed, with a loss of only 16% of the enzyme activity.

3.6.3

Pectinase Purification

Pectinases from various microorganisms have been purified to homogeneity, after the enzyme concentration step, with the purpose of characterizing and studying the


properties of these microbial enzymes. Purification of pectinases has been realized by combinations of chromatography procedures (i.e., ion exchange and gel filtration). The efficiency of the final pectinase yield and specific activity increase depends on the selection of the proper column characteristics and chromatographic conditions [125e127]. Yadav et al. [121] applied two-step sequential chromatographic techniques in combination with dialysis to purify an F. decemcellulare MTCC PL. The enzymatic extract was loaded on a CM-cellulose (CMC) support and a Sephadex G-100 column. The purification procedure resulted in 6.8-fold purification and 2.5% yield. A similar procedure of sequential CMC size-exclusion and Sephadex ion-exchange chromatography was followed to purify an exo-PG from P. variotii with a 41.91-fold increase and recovery yield of 26.90% [122], a PG from Penicillium sp. with a final isolation up to 12-fold [128], and a Streptomyces lydicus PG with a yield of 57.1% and a purification fold of 54.9 [129]. Moreover, a DEAEeSepharose CL-4B anion chromatography exchange and gel filtration by Sephadex-G75 column was also tested in the purification of alkaline pectinase from B. subtilis, with results up to 2.5-fold of the crude enzyme [123]. There are also reports of the use of a single chromatographic technique with high recovery yields of pectinase activity, such as the research carried out by Silva et al. [124], with the utilization of Sephadex G50 column to elute an exo-PG from P. viridicatum RFC3. The purification was efficient, with a 31-fold increase in specific activity and a final yield of 6.5%. Pedrolli and Carmona [130] also described the DEAEeSephadex A-50 anion-exchange column as the only chromatographic method to purify an Aspergillus giganteus PG. Castruita-Domıńguez et al. [131] evaluated the utilization of a nickelcharged agarose affinity chromatography column to purify a Pichia pastoris recombinant polygalacturonase. The eluted recombinant PG preserved more than 60% of the maximum enzymatic activity. PL purification has been widely studied by the evaluation of various combinations of chromatographic columns. Li et al. [123] isolated to apparent homogeneity a PL from a B. clausii strain and estimated that in the DEAEeSepharose Fast Flow column, most of the enzyme activity was present in the first protein peak, reaching a specific activity of 49.5 U/mg and a recovery yield of 12.1%; this behavior was confirmed with gel-filtration chromatography by the presence of the majority of the activity in the first protein peak. The final purified enzyme showed a specific activity of 297.1 U/mg and a yield of 5.5%. Yadav et al. [132] reported that the extracellular PL from Aspergillus terricola was purified by a DEAE cellulose column. The active fractions obtained after ion-exchange chromatography were concentrated by ultrafiltration and dialysis and then loaded on a Sephadex G-100 column. After the final purification step, most of the activity was present in a single protein peak, indicating that the enzyme preparation was relatively pure. The specific activity went up to 4.08 IU/mg, and the recovery was 45.79%. The use of DEAE cellulose and Sephadex G-100 columns was also reported by Fawzi [133] in the SSF production of PL from Geobacillus stearothermophilus Ah22 with a 40.77-fold increase in specific activity. In contrast, PL produced by G. stearothermophilus Ah22 was purified to


61

40.8-fold by a DEAE cellulose anion-exchange column followed by gel-filtration Sephadex G-100 chromatography [134]. Pectin methylesterase (PME) isolation from fruit extract has been studied to determine the enzyme’s stability and kinetic characterization. A PME extracted from guava fruit (Psidium guajava L.) was partially purified by gel filtration on Sephadex G-100 and the highest specific activity peak showed a 104-fold purification of PME compared to the values obtained for the crude extract [135]. Rodrigo et al. [136] studied the isolation of crude tomato PME extract by the use of a cation-exchange column. The elution profile for the enzyme extract showed the PME activity in the first eluted phase. An evaluation of emerging nonchromatographic separation techniques for pectinase purification has been provided by Dogan and Tari [137], with the development of a three-phase partitioned (TPP) bioseparation system. An exo-PG produced by Aspergillus sojae ATCC 20235 was purified by this one-step technique. A TTP separation system was based on the combination of various concentrations of ammonium sulfate saturation with various enzyme extract/tert-butanol ratios. The best results were obtained at 30% saturation of ammonium sulfate and 1:1 (v/v) ratio of crude extract to tert-butanol, with 6.7-fold purification and 25.5% recovery. As of this writing, the most recent advances in this area were carried out by Duque Jaramillo et al. [119]. They reported an aqueous two-phase micellar system for extraction/prepurification of the pectinase produced by A. oryzae, without the presence of activity loss. The developed system was based on the coexistence curve of the Triton X-114/buffer system; the micellar concentrations were separated by a gradual temperature increase to reach a cloudy solution indicating the onset of phase separation, defined as cloud temperature, at which the micelle-rich (larger) cells remained in the bottom phase and the micelle-poor cells migrated to the top phase. Therefore, pectinase enzyme was found in the top phase, and the contaminants partitioned to the bottom phase, probably because of the hydrophobic character of the phase. Moreover, Mehrnoush et al. [138] studied another novel method for pectinase purification denominated as an aqueous organic-phase system (AOPS). The aqueous-based solution is based on the polarity differences between organic solvents and salts; the manipulation of concentrations and volumes between these solutions allows the formation of the two-phase separation. Thus, the hydrophobic interaction between the enzyme and the organic solvent leads to the desirable partition to the top phase and the salting-out effect in the bottom phase. Based on this system, the purification factor of pectinase enzyme was 11.7, with a yield of 97.1%, achieved in an AOPS of 19% (w/w) ethanol and 22% (w/w) potassium.

3.7 Technoeconomic Analysis of Pectinase Production In the scale-up of bioprocesses, the cost estimation is an important factor that should be taken into account for determining whether the process is economically viable and


industrially applicable. With respect to the technoeconomic analysis of pectinase production via SSF or SmF, only a few analyses have been reported. In a work by Nakkeeran et al. [139], they developed a technoeconomic study on the production of polygalacturonases by Aspergillus carbonarius using SmF and SSF. They studied the downstream processing, including an integrated membrane process and other types of alginate affinity precipitation. Fig. 3.1 shows a diagram of polygalacturonase production using an integrated membrane process and Fig. 3.2 shows a diagram of alginate affinity precipitation, in both cases for SmF. It is important to mention that the diagram of alginate affinity precipitation was also applied to SSF. As mentioned earlier the downstream stage for enzyme production is responsible for about 70% of the final cost, therefore it is important to study various configurations. Nakkeeran et al. [139] reported that for a production of 30 kL purified polygalacturonase, the downstream processing cost was 47% lower using an integrated membrane process compared to the alginate affinity precipitation process. Moreover, the total capital investment was lower for SmF using an integrated membrane process compared with SmF and SSF using an alginate affinity precipitation process. Another important aspect,

FIGURE 3.1 Diagram of polygalacturonase production using submerged fermentation with an integrated membrane system. Reprinted from Nakkeeran E, Gowthaman MK, Umesh-Kumar S, Subramanian R. Techno-economic analysis of processes for Aspergillus carbonarius polygalacturonase production. Journal of Bioscience and Bioengineering 113(5):643e40. Copyright (2012), with permission from Elsevier.


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FIGURE 3.2 Diagram of polygalacturonase production using submerged fermentation with an alginate affinity precipitation system. Reprinted from Nakkeeran E, Gowthaman MK, Umesh-Kumar S, Subramanian R. Technoeconomic analysis of processes for Aspergillus carbonarius polygalacturonase production. Journal of Bioscience and Bioengineering 113(5):643e40. Copyright (2012), with permission from Elsevier.

mentioned by Nakkeeran et al. [139], is that it is more appropriate to evaluate the enzyme yield and productivity in terms of carbon source. Additionally, they concluded that the downstream process costs are far more than the upstream process costs.

3.8 Conclusions and Perspectives In this chapter, the development and progress in the production of pectinases shows innovations in all aspects of the process as raw material, technologies of fermentation, downstream processes, and technoeconomic analysis. In general and from the operating point of view, the production of pectinase could be considered a consolidated bioprocess, because this enzyme is produced industrially and has wide applications, mainly in the food industry. Depending on the operational conditions (carbon source or inducer, microorganism, SmF, SSF, downstream process) different types of pectinases


can be produced. In addition, the technoeconomic studies are important to analyze the industrial feasibility of the bioprocess. Moreover, the engineering processes (fermentation technologies/downstream) and the genetics will play important roles in improving the consolidated bioprocess of pectinase production.

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[113] Chen H, Li Y, Xu F. Impact of operating conditions on performance of a novel gas double-dynamic solid-state fermentation bioreactor (GDSFB). Bioprocess and Biosystems Engineering 2013;36: 1753e8. [114] Nagel FJJ, Tramper J, Bakker MS, Rinzema A. Temperature control in a continuously mixed bioreactor for solid-state fermentation. Biotechnology and Bioengineering 2001;72: 219e30. [115] Casciatori FP, Laurentino CL, Costa KKL, Casciatori PA, Thomeó JC. Hygroscopic properties of orange pulp and peel. Journal of Food Process Engineering 2013;36:803e10. [116] Maciel M, Ottoni C, Santos C, Lima N, Moreira K, Souza-Motta C. Production of polygalacturonases by Aspergillus section Nigri strains in a fixed bed reactor. Molecules 2013;18: 1660e71. [117] Flickinger MC. Downstream industrial biotechnology: recovery and purification. Wiley; 2013, ISBN 978-1-118-13124-4. p. 872. [118] Grote F, Ditz R, Strube J. Downstream of downstream processing: development of recycling strategies for biopharmaceutical processes. Journal of Chemical Technology and Biotechnology 2012;87:481e97. [119] Duque Jaramillo PM, Rocha Gomes HA, de Siqueira FG, Homem-de-Mello M, Magalha˜es PO. Liquideliquid extraction of pectinase produced by Aspergillus oryzae using aqueous two-phase micellar system. Separation and Purification Technology 2013;120:452e7. [120] Li M, Sun X, Zhou L, Wang H, Li D, Wang S, et al. Purification of alkaline pectinase in engineering Bacillus subtilis. In: Proceedings of the 2012 international conference on applied biotechnology (ICAB 2012). Berlin, Heidelberg: Springer; 2012. p. 1419e30. [121] Yadav S, Dubey AK, Anand G, Kumar R, Yadav D. Purification and biochemical characterization of an alkaline pectin lyase from Fusarium decemcellulare MTCC 2079 suitable for Crotolaria juncea fiber retting. Journal of Basic Microbiology 2014;54:161e9. [122] Patil NP, Patil KP, Chaudhari BL, Chincholkar SB. Production, purification of exopolygalacturonase from soil isolate Paecilomyces variotii NFCCI 1769 and its application. Indian Journal of Microbiology 2012;52:240e6. [123] Li Z, Bai Z, Zhang B, Li B, Jin B, Zhang M, et al. Purification and characterization of alkaline pectin lyase from a newly isolated Bacillus clausii and its application in elicitation of plant disease resistance. Applied Biochemistry and Biotechnology 2012;167:2241e56. [124] Silva D, Martins ES, Leite RSR, Da Silva R, Ferreira V, Gomes E. Purification and characterization of an exo-polygalacturonase produced by Penicillium viridicatum RFC3 in solid-state fermentation. Process Biochemistry 2007;42:1237e43. [125] Gummadi SN, Panda T. Purification and biochemical properties of microbial pectinases - a review. Process Biochemistry 2003;38:987e96. [126] Jacob N. Pectinolytic enzymes. In: Singh-Nee Nigam P, Ashok, editors. Biotechnology for agroindustrial residues utilisation. Netherlands: Springer; 2009. p. 383e96 [Chapter 21]. [127] Heftmann E. Chromatography: Fundamentals and Applications of Chromatography and Related Differential Migration Methods-Part B: Applications. 6th ed. The Netherlands: Elsevier; 2004. [128] Patil NP, Chaudhari BL. Production and purification of pectinase by soil isolate Penicillium sp. and search for better agro-residue for its SSF. Recent Research in Science and Technology 2010;2: 36e42. [129] Jacob N, Asha Poorna C, Prema P. Purification and partial characterization of polygalacturonase from Streptomyces lydicus. Bioresource Technology 2008;99:6697e701.


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Cellulases

4

Reeta R. Singhania1, M. Adsul1, A. Pandey2, A.K. Patel1, * DBT- IOC C ENTRE FOR ADV ANCED BIOE N ER GY RE SEARCH, INDIANOIL C ORPORATION LIMITED; 2 CE NTER OF INNOVATIVE AND APPLIED BIOPROCESSING, (A NATIONAL INSTITUTE UNDER DEPT OF BIOTECHNOL OGY, MINISTRY OF S&T, GOVT OF INDIA), MOHALI, PUNJAB , INDIA 1

4.1 Introduction Cellulases are the second largest industrial enzyme by dollar volume and have experienced increased demand since 1995 in several industrial applications in the detergent industry, textile industry, animal feed and food industry, paper industry, and biofuel industry. The commercial potential of cellulases lies in their efficiency at converting cellulose into its monomers. Since 2005 cellulases have received tremendous popularity because of the resurgence in lignocellulosic ethanol production via the enzymatic route. Bioethanol is seen as an alternative to fossil fuel. Even though there are various routes of converting cellulosic biomass to ethanol, the enzymatic route is most popular because of its sustainability and environmental friendliness. Cellulases are multienzyme complexes consisting of mainly three different components, endo-1,4-b-D-glucanase (EC 3.2.1.4), exoglucanase/exo-cellobiohydrolase (EC 3.2.1.91), and b-glucosidase (EC 3.2.1.21). All these three components act synergistically to hydrolyze the cellulose polymer completely into its glucose monomers (Fig. 4.1). Endo1,4-b-D-glucanase acts first, randomly, in between cellulose fibers to generate reducing FIGURE 4.1 Enzymatic hydrolysis of cellulose. A schematic diagram showing cellulase synergy is presented.

*


Current Developments in Biotechnology and Bioengineering: Production, Isolation and Purification of Industrial Products http://dx.doi.org/10.1016/B978-0-444-63662-1.00004-X Copyright © 2017 Elsevier B.V. All rights reserved.

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and nonreducing ends, which are further attacked by exo-cellobiohydrolase, releasing cellobiose, which is glucose dimers linked by b-1,4 glycosidic bonds. Cellobiose is finally hydrolyzed by b-glucosidase (BGL) into glucose monomers, the final end product. Most of the cellulases produced by filamentous fungi, including Trichoderma reesei, are limited by the amount of BGL. BGL plays an important role in overall cellulase performance and is the limiting factor because of its glucose-sensitivity property, in that it is inhibited by its own end product (glucose), and more importantly, it is also known to be inhibited by its own substrate, cellobiose. So, in view of these facts, a BGL that was insensitive to glucose and cellobiose, or at least tolerant to some extent, would be highly desirable for the hydrolysis performance of cellulase on cellulosic biomass for conversion to glucose. Research based in this direction has opened up several new avenues for overall improved cellulase performance. Potent glucose-tolerant BGL has been reported from Aspergillus niger and Aspergillus oryzae as well as Candida peltata. One of the major approaches taken toward improving cellulase for biomass hydrolysis includes increasing the copy number of the BGL gene so as to improve the amount of BGL in the cellulase mixture produced by a fungal strain, and the other is to introduce a glucose-tolerant BGL-producing gene into the fungal strain producing other components of cellulases in optimal amounts. Modification of cellulase properties to enhance efficiency or impart desired features is another area of research. Proteinengineering approaches adopted for cellulase modification provide basic information about cellulase enzymes at the molecular level, which could be crucial for designing strategies toward the genetic improvement of fungal strains for enhanced cellulase production. The success of the lignocellulosic ethanol project will depend on the ability to generate an efficient cellulase system. Preparations of cellulase cocktails by mixing various components from heterogeneous sources at varied ratios have become attractive, as the overall cellulase activity could increase. Hydrolytic efficiency of multienzyme complexes for lignocellulosic biomass saccharification depends on both the properties of the individual enzyme components and their ratios in the cocktail. Filamentous fungi are the major source of cellulase and hemicellulase and various mutant strains of Trichoderma are the best-known producers; however, their amount of BGL is comparatively lower, resulting in inefficient hydrolysis of biomass. The ideal cellulase cocktail must possess high activity on the intended biomass feedstock, should be able to hydrolyze the biomass completely, and should be able to operate at mild acidic pH and withstand process stress and be cost-effective, too. Thus, identifying a potent cellulase producer is the key to the success of any technology involving cellulases. Microbial sources have been extensively explored for cellulase production.

4.2 Sources All living beings are carbohydrate degraders, but are not necessarily cellulose degraders. They are highly dependent on their microflora for cellulose digestion. Only microorganisms have the capacity to produce cellulases in nature; hence they have the capacity

Chapter 4 Cellulases

75

to hydrolyze or utilize cellulose. However, there are a few reports that show that several mollusks, including snails, a sea slug, a periwinkle, and some bivalves, have the capacity to produce cellulase. Some researchers have reported possible endogenous enzyme sources such as the hepatopancreas, gastric teeth, and crystalline styles [1]. Cellulolytic microbes are primarily carbohydrate degraders and are generally unable to utilize lipids or proteins as a source of energy for growth [2]. The order Actinomycetales comprises two genera that produce cellulases, which are Microbispora and Thermomonospora. Fractionation of Microbispora bispora cellulases has identified six different enzymes, whereas fractionation of Thermomonospora fusca cellulases has identified five different enzymes, all of which were purified to homogeneity and partially characterized [3]. Both aerobic and anaerobic forms of bacteria are involved in cellulose degradation. The anaerobic bacteria are found in the soil, on decaying plant materials, in rumens, in sewage sludge, in termite gut, in wood-chip piles, in compost piles, and at paper mills and wood-processing plants. Examples of some anaerobic bacteria with active cellulolytic systems are Acetivibrio cellulolyticus, Bacteroides cellulosolvens, Butyrivibrio fibrisolvens, Clostridium acetobutylicum, Clostridium aldrichii, Clostridium cellobioparum, Clostridium cellulofermentans, Clostridium cellulolyticum, Clostridium cellulovorans, Clostridium herbivorans, Clostridium hungatei, Clostridium josui, Clostridium papyrosolvens, Clostridium thermosuccinogenes, Ruminococcus albus, Ruminococcus flavefaciens, and Clostridium thermocellum [4]. The aerobic bacteria are usually found in soil, in water, on plant materials, in humus, in animal feces, in sugarcane fields, and in leaf litter. These are Bacillus megaterium, Bacillus pumilus, Cellulomonas fimi, Cellulomonas flavigena, Cellulomonas gelida, Cellulomonas iranensis, Cellulomonas persica, Cellulomonas uda, Cellvibrio gilvus, Cellvibrio mixtus, Pseudomonas fluorescens, Streptomyces antibioticus, Streptomyces cellulolyticus, Streptomyces lividans, and Streptomyces reticuli [5]. Fungi are the most studied microorganisms for cellulase production because of their higher enzyme yield and their ability to secrete complete cellulase complex extracellularly. These are Trichoderma reesei, Penicillium pinophilum, Penicillium funiculosum, Fusarium oxysporum, Aspergillus niger, Sclerotium rolfsii, and Humicola sp. However, other cellulolytic systems of Phanerochaete chrysosporium, Talaromyces emersonii, and Melanocarpus albomyces and other anaerobic fungi belonging to the genera Neocallimastix, Caecomyces, and Oprinomyces have also been well characterized. Thermophilic fungi such as Sporotrichum thermophile, Thermoascus aurantiacus, Chaetomium thermophilum, Humicola grisea, and Myceliopthora thermophila are also known to produce cellulases. These fungi are of more interest because of their capacity to produce thermostable cellulases. These enzymes have shown stability at highly acidic or alkaline pH as well as temperatures up to 90 C. To overcome the complexity of cellulosic substrates, fungi often produce a variety of cellulase components that differ in their molecular characteristics such as molecular weight, amino acid composition and sequence, isoelectric point, carbohydrate content, etc. They also differ in their capacity to adsorb onto cellulose, their substrate specificity, and even their catalytic activities [6].


Most of the cellulolytic fungi and bacteria such as Cellulomonas are capable of utilizing a variety of other carbohydrates in addition to cellulose for their growth, whereas anaerobic cellulolytic species such as Clostridium have a strict carbohydrate range, which lies between cellulose and its hydrolytic products [7]. However, large numbers of microorganisms are known to degrade cellulose, such as white rot fungi, soft rot fungi, aerobic and anaerobic bacteria, actinomycetes, etc., but only a few of them are able to produce it extracellularly, being capable of degrading crystalline cellulose. Most of the bacterial cellulases are incapable of hydrolyzing cellulose, being incomplete. Certain filamentous fungi have the characteristic property of secreting large amounts of extracellular protein, being most suited to higher production of cellulases. These cellulases contain all three components in different proportions and hence are capable of hydrolyzing native cellulose completely. The most commonly studied microorganisms include fungi (Trichoderma, Penicillium, Humicola, Aspergillus), bacteria (Cellulomonas, Pseudomonas, Clostridium), and actinomycetes (Streptomyces). Most of the industrial cellulases available are produced from filamentous fungi such as Trichoderma, Penicillium, Phanerochaete, Fusarium, Humicola, and Aspergillus. Trichoderma reesei is as of this writing known to be the best cellulase producer, capable of hydrolyzing native as well as derived cellulose into glucose. Table 4.1 gives brief information about microbial sources of cellulose. Fungal cellulases are different from bacterial cellulases and are preferred because of their high production titers. Both fungal and bacterial cellulases have been discussed. Though in the environment, more cellulolytic organisms are present, the majority of organisms cannot be cultured by using available protocols. Consequently, the isolation and characterization of cellulases from uncultured organisms from nature have been limited. Advances in molecular techniques like metagenomics made it possible to screen a large number of cellulolytic genes. Metagenomics is a cultivation-independent analysis of the metagenome of a habitat and involves direct isolation of DNA from the environment followed by cloning and expression of the metagenome in a heterologous host [8]. Although characterization of cellulases from uncultured microorganisms could still reveal much information about cellulolytic diversity and industrial potential, there are very few reports on metagenome-derived cellulases [9]. In 2009, a metagenomic approach was employed to produce ionic-liquid-stable endoglucanases. However, they were found to be moderately active and stable in 1-butyl-1-methyl-pyrrolidinium trifluoromethanesulfonate [10].

4.3 The Cellulase System 4.3.1

Noncomplex System

The most studied fungus for cellulase production is T. reesei. It produces two cellobiohydrolases (CBHs) as CBHI and CBHII and two endoglucanases (EGs) as EGI and EGII in a rough proportion of 60:20:10:10, which together constitute up to 90% of the enzyme


Table 4.1

List of Potent Cellulase Producers

Major Group

Genus

Species

Fungi

Aspergillus

A. niger A. fumigatus A. terreus A. nidulans A. oryzae F. solani F. oxysporum H. griesa H. insolens T. aurantiacus C. thermophilum M. albomyces N. crassa P. fusisporus P. chrysosporium P. brasilianum P. occitanis P. decumbens P. janthinellum P. funiculosum P. pinophilum T. reesei T. longibrachiatum T. harzianum T. viride A. cellulolyticus B. subtilis B. megaterium B. pumilus C. acetobutylicum C. thermocellum C. hungatei C. cellulolyticum P. cellulosa R. marinus C. fimi C. bioazotea C. uda S. drozdowiczii S. cellulolyticus S. antibioticus S. lividans T. fusca T. curvata

Fusarium Humicola Thermoascus Chaetomium Melanocarpus Neurospora Paecilomyces Phanerochaete Penicillium

Trichoderma

Bacteria

Acidothermus Bacillus

Clostridium

Actinomycetes

Pseudomonas Rhodothermus Cellulomonas

Streptomyces

Thermomonospora

Modified from Singhania RR. Cellulolytic enzymes. In: Nigam P, Pandey, A, editors. Biotechnology for agro-industrial residues utilization. USA: Springer; 2009. p. 371e82.

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cocktail, whereas seven BGLs (BGLI to BGLVII) secreted typically by this fungus constitute up to 1% of the secreted protein. All these components act synergistically to convert crystalline cellulose into glucose. The majority of cellulases have a characteristic two-domain structure, a catalytic domain and a cellulose-binding domain (CBD), which is also termed as a carbohydrate-binding module, connected through a linker peptide. As per the name, the catalytic domain contains the catalytic site and the CBD facilitates binding of the enzyme to the substrate/cellulose. Broadly there are two kinds of cellulase systems, complex and noncomplex. Noncomplex systems are more common and most exploited for industrial applications. These are characteristic of aerobic fungi and are largely described based on Trichoderma, Penicillium, Aspergillus, Phanerochaete, Fusarium, Humicola, etc., in which a large number of cellulases are found. All these fungal cellulases have been studied in detail. The Humicola insolens cellulolytic system is analogous to T. reesei, with two CBHs and five EGs, but it lacks CBD in both exo- and endoglucanases. H. grisea produces a thermostable EG (Cel12A) enzyme with sequence similarity to T. reesei Cel12A25. Aspergillus niger is also known to produce all three components of the cellulase complex and exhibits a strong hydrolytic effect against cellulose, but is mostly known for its high BGL activity, comparatively higher than that of T. reesei. Some of these BGLs are known to be glucose tolerant [12]. Normally T. reesei’s BGL is subject to product inhibition; however, it is sufficient to support its growth on cellulosic material. But for biomass hydrolysis at the industrial level it is often supplemented with Aspergillus BGLs for efficient saccharification. Phanerochaete chrysosporium, a white rot fungus, produces a complex array of cellulases, hemicellulases, and ligninases capable of lignocellulose degradation. High titers of cellulase production are also reported in Penicillium. Noncomplex cellulase systems have also been reported in some of the aerobic bacteria like Thermobifida, which also produces all the major components of cellulases.

4.3.2

Complex Cellulases/Cellulosomes

Complex cellulase systems are characteristic of anaerobic bacteria (however, a few anaerobic fungi have also been reported), which produce high-molecular-weight complexes called “cellulosomes.” Cellulosomes protuberate from the cell wall of the bacteria and harbor stable enzyme complexes and are capable of bringing about degradation of cellulose by effectively binding to it. By far the most comprehensive information on cellulosomes is available from the system of Clostridium as C. thermocellum has been studied in detail. The cellulaseehemicellulase complex of C. thermocellum contains up to 26 polypeptides with at least 12 endo- and exocellulases, 3 xylanases, a lichenase, and a noncatalytic cellulosome-integrating protein (CipA), or scaffoldin. Enzymes bind through dockerin moieties onto complementary receptors on scaffoldin called cohesions. The basic architectural structure is conserved in cellulosomes of all anaerobic systems; however, the kinds of activities and the number of catalytic domains may vary in other anaerobic bacteria.


79

Cellulosomes in anaerobic fungi have a catalytic subunit linked by a serine/ threonine-rich linker to two or three copies of a 40-amino-acid cysteine-rich, noncatalytic docking domain (NCDD), which is conserved. The NCDD bears no homology to bacterial dockerins, but in contrast does have similar size and number of polypeptides. Enzymes associated with fungal cellulosomes are modular and are from the genera Neocallimastix, Oprinomyces, and Piromyces. The molecular arrangement of fungal cellulosomes is still unknown.

4.4 Regulation of Cellulase Expression The synthesis and action of cellulases are intricately controlled by the organisms producing this enzyme. Cellulases are inducible enzymes and their secretion/production is meticulously controlled by activation and repression mechanisms of coordinately regulated genes. Significant information is available on cellulase gene expression in filamentous fungi, especially on T. reesei, which could be very well taken as a model organism for explanation. Lactose is a known inducer of cellulases that has been employed commercially for economic reasons. The mechanism of lactose induction is not fully understood; however, it is known that a lack of galactose mutarotase activity is crucial for fungal cellulase induction [13]. Sophorose is a proposed inducer of cellulase, at least for T. reesei, which could be generated by trans-glycosylation activity of basally expressed BGL. Cellobiose, 1,5-lactone, and other oxidized products of cellulose hydrolysis can also act as inducers of cellulases. The T. reesei cellulase gene was also induced by several other disaccharides, namely, laminaribiose, gentiobiose, and xylobiose [14,15]. The monosaccharide L-sorbose was also found to induce cellulase [16]. Inducers need to be transported inside cells to induce cellulase-producing genes. Because cellulose is too large to be transported into cells, an inducer capable of passing through the cell wall needs to be formed when cellulose is used as the inducer for cellulase production. The expression of a very low level of cellulase is considered to be related to the formation of an inducer from cellulose. The conidia of T. reesei have been shown to contain cellulases, mainly CBHII, on their surface. These conidia-bound cellulases are supposed to be responsible for the initial release of an inducer from cellulose, resulting in cell growth on cellulose [17,18]. However, cellulase transcripts are not detected from glucose-, sorbitol-, or glycerol-grown mycelia in Northern blot analysis and no hybridization signals of cbh1, cbh2, or eg1 were detected in a slot-blot analysis [19]. The presence of glucose, which is easy to metabolize and energetically favorable to the microbe, leads to the repression of other genes needed for the use of other carbon sources. The controlling mechanism is called glucose (carbon catabolite) repression. The cellulase production by T. reesei is under glucose repression. In filamentous fungi, glucose repression is manipulated by the transcription factors CREA and CREI, which are the proteins responsible for glucose repression [20e22]. The expression of cre1 in T. reesei is regulated by the carbon source, and the expression is higher in the presence of


an inducing carbon source than in the presence of glucose [22,23]. This indicates that the CREI protein function is regulated at the posttranscriptional level or by the presence of other factors. Trichoderma reesei Rut-C30 is a glucose-derepressed mutant strain, which contains a truncated cre1 gene. Isolation of catabolite repressor mutants is a better choice for high-titer cellulase production. An analogue of glucose such as 2-deoxy Dglucose is generally used for isolation of catabolite repressor mutants. Though this regulation can be overcome by strain improvement, attempts to increase cellulase titers are usually tried by employing improved bioprocess technologies for production.

4.5 Research on Bioprocesses for Improved Cellulase Production Production of low titers of cellulase has always been a major concern, and thus several workers are trying to improve production titers by adopting multifaceted approaches such as the use of better bioprocess technologies, using cheaper or crude raw materials as substrates for enzyme production, bioengineering the microorganisms, etc. [24]. Bioprocess improvement strategies for enhancing the yield and specific activities of cellulases have also been well addressed by researchers worldwide. The majority of reports on microbial production of cellulases utilize the submerged fermentation (SmF) technology and the widely studied organism used in cellulase production, T. reesei, has also been tested mostly in liquid media. The SmF method involves the cultivation of microorganisms in a nutrient-rich aqueous medium. However, considerable expense is involved in concentrating and extracting enzymes from this large aqueous environment. An alternative to the SmF method is solid-state fermentation (SSF), which involves the growth of microorganisms on solid materials in the absence of free liquid. In nature, the growth and cellulose utilization of aerobic microorganisms elaborating cellulases resembles SSF rather than a liquid culture [25]. Since 1995, SSF has regained interest because of the high titers of enzyme production employing fungal cultures. The lignocellulosic substrate type had the greatest impact on cellulase secretion. Some of the substrates significantly stimulated lignocellulolytic enzyme synthesis without supplementation of the culture medium with specific inducers [26]. Nevertheless, the advantages of better monitoring and handling are still associated with the submerged cultures. Cellulase production in cultures is growth associated and is influenced by various parameters including the nature of the cellulosic substrate, pH of the medium, and nutrient availability; and a large-scale production of cellulases requires understanding and proper control of the growth and enzyme production capabilities of the producer. This is, however, extremely complicated because many factors and their interactions can affect cellulase productivity. Microbial cellulases are subject to induction and repression mechanisms and the process design and media formulation for cellulase production have to take care of these aspects. The media formulation for fermentation is of significant concern because no


81

general composition can give the optimum growth and cellulase production. Also, the medium used is mostly specific for the organism concerned. In T. reesei, a basal medium after Mandel and Reese [27] has been most frequently used with or without modifications. The carbon sources in the majority of the commercial cellulase fermentations are cellulosic biomass including straw, spent hulls of cereals and pulses, rice or wheat bran, bagasse, paper industry waste, and various other lignocellulosic residues that induce cellulase production. The majority of the cellulase production processes are batch processes, but fed-batch or continuous mode helps to override the repression caused by the accumulation of reducing sugar. The major technical limitation in fermentative production of cellulases remains the increased fermentation times with low productivity. Information on the types of bioprocesses employed for cellulase production, microorganisms employed, as well as magnitude of production, is available in reviews by Sukumaran et al. [28] and Singhania et al. [29].

4.5.1

Solid-State Fermentation

SSF is defined as fermentation in the absence or near absence of free water [30]. SSF for the production of industrial enzymes is rapidly gaining interest as a cost-effective technology, as the microorganisms, especially fungal cultures, produce comparatively high titers of metabolites in response to the conditions of fermentation, which shows similarity to the natural environment [31]. Filamentous fungi such as T. reesei, A. niger, Penicillium sp., etc., have been employed for cellulase production using SSF in which a basal mineral salts medium was used for moistening the substrate. A koji chamber can be used for large-scale production for economic reasons, though maintenance of sterile conditions is difficult. Any cellulosic biomass could be employed as substrate. For cellulase production, inocula can be prepared in a stirred-tank reactor and can be sprayed onto the sterile medium in the shallow tray. Either spores or mycelia can be used as inoculum in the case of filamentous fungi. In this case, temperature and humidity are controlled inside the chamber, and incubation is allowed for 7 days or as specified. A suitable buffer or distilled water with appropriate Tween percentage is used as the extraction liquid. The medium is homogenized with extraction liquid and centrifuged to remove the biomass and cell debris. The supernatant contains the extracellular cellulase, which could be concentrated by acetone precipitation or salting out. For biomass hydrolysis, it could be used as is, and for other applications it depends on the degree of purity of cellulase required. A well-designed solid-state fermenter should (1) have perfect control systems for temperature, airflow rate, and humidity; (2) have a well-designed system for preventing contamination; (3) be homogeneous in water activity, temperature, and composition so that microbes can grow uniformly; (4) be able to remove harmful metabolites, such as CO2, quickly; and (5) be labor-saving and easy to scale up for handling solid medium. As of this writing, none of the available SSF bioreactors can satisfy all these points. Several bioreactors that were engineered for cellulase production to satisfy the discussed points


and enable continuous monitoring are the shallow-tray fermenter, column fermenter, deep-trough fermenter, rotating-drum fermenter, stirred-tank fermenter, rotating-disk reactor, rocking-drum reactor, and fluidized-bed fermenter, though each of them has its own limitations [32]. Fermenters are engineered to maintain growth and production conditions. There are several key factors that play important roles in cellulase production via SSF, such as pH, temperature, moisture content and water activity, aeration, and substrate composition. These operating conditions may differ with the organism and substrate used. For example, fungi prefer to grow at acidic pH and low moisture content (35e70%) compared to bacteria (70e90%) and usually grow well at 25e30 C, whereas bacteria prefer neutral pH and high moisture content and grow well at 37 C. In the case of SSF, a lot of heat is generated by the vigorous metabolic activities of organisms. There is always a limitation on heat transfer as it is relatively poor in the solid layer and overheating occurs in the substrate particle. This causes unfavorable conditions for spore germination, mycelium growth, and enzyme accumulation and secretion. Temperature control in the environment of the solid-state fermenter is relatively convenient to achieve, but temperature regulation within the solid substrate layer is relatively difficult. A few measures can be taken, such as a proper thickness of the solid phase, which could facilitate heat transfer, and also the aeration rate has to be controlled to supply oxygen and mass transfer, for which convection could be employed. Controlling the moisture content in the medium is the key factor for cellulase production and is an essential component for the growth of microorganisms. If the water content is high, the void space, as well as the gas-phase volume within the solid substrate, is reduced, which increases the mass transfer resistance of oxygen and carbon dioxide, as well as the possibility of contamination, whereas low water content is unfavorable to spore germination and substrate swelling. Substrate swelling is essential for fungi to attack and digest the solid substrate. Water activity is even more important than the moisture content and is closely related to moisture content, but is not exactly equal to it. It gives the amount of unbounded water in the immediate surroundings. It is necessary to maintain the optimal value, but this tends to vary because of metabolism and evaporation. To a certain extent, it can be controlled by a humidifier, which could be incorporated into the solid-state fermenter/bioreactor. Another important factor is pH, which affects the growth of microorganisms and hence the cellulase production. It is difficult to monitor the pH in a solid substrate, but the pH of the basal medium, which usually contains nitrogen sources having buffering capacity, can be adjusted. Solid cellulosic biomass has a buffering capacity, which rules out the necessity to adjust the pH during SSF. Though there are several indirect methods for biomass measurement, such as total protein estimation, fungal cell wall component measurement (N-acetylglucosamine), etc., as well as direct methods such as CO2 evolved and O2 intake, in the case of SSF, measurement of the biomass is difficult. It is not feasible to monitor the growth pattern of microorganisms, which makes it difficult to develop suitable models for SSF. Nevertheless, SSF can be proposed as a better technology for commercial


83

production of cellulases considering the low-cost input and ability to utilize naturally available sources of cellulose as substrate.

4.5.2

Submerged Fermentation

SmF has been defined as fermentation in the presence of excess water. Almost all the large-scale enzyme-producing facilities are using the proven technology of SmF because of better monitoring and ease of handling. Though bacteria and actinomycetes are also available for cellulase production, the titers are too low to make the technology economically feasible. Most of the commercial cellulases are produced by the filamentous fungi T. reesei and A. niger under SmF. Cellulase production in culture is highly influenced by various parameters including the nature of the cellulosic substrate, pH of the medium, nutrient availability, inducer supplementation, fermentation temperature, etc. Mostly, pure cellulose preparations like Solka-Floc and Avicel have been used in liquid cultures of cellulolytic microbes for production of the enzymes, but while using soluble substrates, the breakdown products may hamper cellulase synthesis by promoting catabolite repression due to the accumulation of free sugars. Increased production in a fermenter may be achieved by a gradient feed of a suitable cellulose and maintenance of process conditions at their optimum. Large-scale production of cellulases requires understanding and proper control of the growth and enzyme production capabilities of the producer. Cellulases produced by compost organisms such as the filamentous fungi Trichoderma, Penicillium, Aspergillus, Humicola, etc., can perform at diverse ranges of pH and temperature. Microbial cellulases are subject to induction and repression mechanisms and the process design and medium formulation for cellulase production has to take care of these aspects. A two-stage continuous process for cellulase production could be employed in which the growth phase and production phase could be separated by different pH and temperature optima. This could help in overcoming the technical limitations of low productivity and long fermentation time for cellulase production. Repression by glucose and cellobiose is a known feature of cellulase systems, and several attempts have been directed toward the development of mutants resistant to catabolite repression. For SmF, huge bioreactors are available and they also provide ease of control of various operating factors such as pH, temperature, aeration, etc. As of this writing, SmF is the most accepted technology for industrial production of primary and secondary metabolites. In SmF, all the parameters required for modeling can be monitored, and hence most of the modeling studies have been done for metabolites production via SmF. A list of fermentation technologies adapted for cellulase production, with the magnitude, microorganism employed, as well as the amount of cellulase produced, has been given by Sukumaran et al. [28] as well as Singhania et al. [29]. Types of bioprocesses and their magnitude for cellulase production by various microorganisms are listed in Table 4.2.


Table 4.2 Type of Bioprocess and Its Magnitude for Cellulase Production by Various Microorganisms Microorganism

Substrate

Method

Enzyme and Activity

References

Aspergillus niger NII-08121 A. niger NRRL3

Wheat bran

SmF

BGL, 1400 U/mL

[12]

Wheat bran/ corncob Untreated oil palm trunk

SSF

Cellobioase, 215 IU/g cellulose

[33]

SSF

[34]

CMC cellulose/ glycerol Carboxymethyl cellulose Soybean industry residue Banana waste

SmF SmF

CMCase, 54.27 U/g substrate FPase, 3.36 U/g substrate BGL, 4.54 U/g substrate CMCase, 1.9 U/mL, cellobiase, 1.2 U/mL CMCase, 0.17 U/mg protein

SSF

FPase, 1.08 U/mg protein

[37]

SSF

[38]

Chaetomium thermophilum CT2 Melanocarpus albomyces Mixed culture Trichoderma reesei and A. niger Mucor circinelloides Neurospora crassa

Cellulose

SmF

FPase, 2.8 IU/gds, CMCase, 9.6 U/gds, cellobiase, 4.5 U/gds CMCase, 2.7 IU/mL

Solka-Floc

SmF

[40]

Rice chaff/wheat bran (9:1)

SSF

Cellulase, 1160 ECU/mL, endoglucanase, 3290 ECU/mL FPase, 5.64 IU/g

Lactose Wheat straw

SmF SmF

[42] [43]

Penicillium decumbens

Wheat straw/ bran (8:2) Paper pulp

SSF

EGL, 0.25 U/mL FPase, 1.33 U/mL, CMCase, 19.7 U/mL, BGL, 0.58 U/mL FPase, 20.4 IU/g FPase, 23 U/mL, CMCase, 21 U/mL FPase, 0.55 U/mL, CMCase, 21.5 U/mL, BGL, 2.31 U/mL Cellulase, 3.2 IU/mL

[45]

FPase, 8.3 U/mL, endoglucanase, 37.3 U/mL 3.7 FPU/mL 134 FPU/g delignified sugarcane bagasse Endoglucanase, 97.7 U/mL CMCase, 148 IU/mL, Avicelase, 45 IU/mL, BGL, 137 IU/mL Endoglucanase, 987 U/gds, BGL, 48.8 U/gds

[48]

Aspergillus fumigatus SK1 Bacillus pumilus Bacillus sp. KSM N252 Bacillus subtilis B. subtilis

Penicillium occitanis

SmF, fedbatch SmF

Penicillium janthinellum

Sugarcane bagasse

P. janthinellum EMS UV-8 Penicillium echinulatum

Wheat bran/ Avicel Cellulose

P. echinulatum

Pretreated sugarcane bagasse CMC CMC

SmF SmF

Wheat straw

SSF

Rhodothermus marinus Streptomyces sp. T3-1

Thermoascus aurantiacus

SmF SmF, batch and fed-batch SmF fedbatch

[35] [36]

[39]

[41]

[44]

[46]

[47]

[49]

[50] [51]

[52]


85

Table 4.2 Type of Bioprocess and Its Magnitude for Cellulase Production by Various Microorganismsdcont’d Microorganism

Substrate

Method

Enzyme and Activity

References

Trichoderma reesei RUT-C30 T. reesei

Wheat bran

FPase, 3.8 U/gds

[53]

FPase, 0.69 U/mL/min

[54]

T. reesei

Steam-treated willow Corncob residue Corn stover residue Sugarcane bagasse

SSF, shake flask SmF, continuous SmF

FPase, 108 U/g cellulose

[55]

FPase, 158 U/gds Cellulase, 5.48 IU/mL, FPase, 0.25 U/mL FPase, 0.88 U/mL, CMCase, 33.8 U/mL, BGL, 0.33 U/mL

[56] [57]

T. reesei ZU 02 T. reesei ZU 02 Trichoderma viride

Xylose/sorbose

SSF SmF SmF

[46]

BGL, b-Glucosidase; CMC, carboxymethylcellulose; CMCase, carboxymethylcellulase; EGL, endoglucanase; FPase, filter paper assay units; SmF, submerged fermentation; SSF, solid-state fermentation. Modified from Singhania RR, Sukumaran RK, Patel AK, Larroche C, Pandey A. Advancement and comparative profiles in the production technologies using solid-state and submerged fermentation for microbial cellulases. Enzyme and Microbial Technology 2010;46:541e9.

The key to developing robust cellulases is to artificially construct them, either by enzyme assembly to form cocktails or by engineering cellulase producers to express the desired combination of cellulase enzymes.

4.6 Strain Improvement There are three main methods of strain improvement available to increase the extracellular production of cellulases, i.e., (1) mutagenesis and selection, (2) genome shuffling, and (3) gene cloning.

4.6.1

Mutagenesis and Selection

Random mutagenesis and selection is also called classical strain improvement or nonrecombinant strain improvement. In this method there is a permanent alteration of one or more nucleotides at a specific site along the DNA strand. Mutation may be associated with a change of a single nucleotide (point mutation), through substitution, deletion, or rearrangement of one or more base pairs in the chromosome. In many cases mutations are harmful but sometimes they create a strain that is more adaptable to its environment and improve its biocatalytic performance. This method involves three basic steps: (1) mutagenesis of a population to induce genetic variability, (2) random selection and primary screening of the surviving population to find an improved strain, and (3) actual checking for the desired improvement in fermentation or by assay. The improved mutant may again act as a parent strain for further mutagenesis. This is continued until we get a superior mutant. An improvement is achieved by subjecting the cells or spores


Table 4.3 Chemical and Physical Mutagens Used for Strain Improvement Mutagen

Role

Physical X-rays, g-rays UV rays

Single- or double-strand breakage of DNA Pyrimidine dimerization

Chemical (Base Analogues) 5-Chlolouracil 5-Bromouracil 2-Aminopurines deamination agent Hydroxylamine

Results in incorrect pairing Results in incorrect pairing DNA replication error Deamination of C

Nitrous Acid

Deamination of C, A, and G

Alkylating Agents N-nitrosoguanidine Ethyl methanesulfonate

Alkylation of C and A Alkylation of C and A

Intercalating Agents Ethidium bromide, acridine dyes

Intercalation between two base pairs

of a microbial strain to a variety of chemical or physical agents called mutagens. Table 4.3 shows some examples of mutagens. The efficiency of random selection also depends on several other factors such as type of culture used (spores or conidia), mutagen dose and time of exposure, type of damage to the DNA (deletion, addition, transversion, substitution of bases, or breakage of DNA), frequency of mutagen treatment, etc. [58]. In the case of cellulase improvement, after mutagen treatment of the spores or conidia, the desired mutants are screened on solid agar plates or they are enriched in a liquid medium. On solid agar plates, the zone of hydrolysis of cellulosic materials around the colony is the measure of an improved mutant for cellulase production. The cellulosic materials used for selection are soluble carboxymethylcellulose, insoluble Solka-Floc, Avicel, and acid-swollen cellulose. The fungal strain improvement for cellulase production was first carried out by Mandel and Weber [59]. They screened more than 100 wild-type strains of Trichoderma species to identify the best cellulolytic strain, and the best strain, called Trichoderma viride QM6a, was selected. The mutant, QM6a, was further mutated with UV light and chemicals, which resulted in the isolation of mutant QM 9414, with higher filter paper activity [60]. The work was continued by other people to produce more improved strains such as strains M7, NG14 [61], and RUT-C30 [62]. A mutant of P. funiculosum was isolated using UV irradiation. This mutant showed an ability to metabolize inorganic nitrogen sources like urea and sodium nitrate both for growth and for enzyme production [63]. A number of mutant strains overproducing cellulase, BGL, and xylanase were isolated from the cellulolytic fungus P. pinophilum


87

87160iii by UV mutagenesis. Selection was carried out using either an agar plate or an enrichment technique. Cellulase [filter paper-hydrolyzing activity (FPA)] production by some of the mutants in shake-flask culture was approximately fourfold higher than that of the wild-type strain. Improvements in BGL production were on the order of eight- to ninefold. The morphology of the mycelium of the mutants was quite different from that of the wild type. The mutants, for example, produced mycelia that were highly branched and thicker in cross section. In several of the mutants, synthesis of xylanase and BGL was completely derepressed in the presence of glycerol, which is a known repressor of the synthesis of these enzymes. Several mutants produced BGL enzyme that showed altered kinetics of hydrolysis in the presence of inhibitors [64]. A mutant of P. pinophilum produced cellulase (FPA) activity of 9.8 units/mL at productivities of 137 units/L/h in submerged culture in a fermenter. BGL levels were on the order of 35 units/mL. The productivity of cellulase by mutant NTG III/6 was comparable to that produced by the best mutant (C-30) isolated from the extensively studied T. reesei. Especially, the yield of BGL by P. pinophilum is much higher than that by T. reesei [65]. Mutant Pol6 of Penicillium occitanis is an interesting strain that produces both cellulases and hemicellulases. When nitrate was used in Mandels and Weber’s basal growth medium with a C/N ratio below 20.2, it resulted in more cellulase production than when urea or ammonium sulfate was used. Crude substrates such as wheat bran and wheat flour residues were used in combination with a local cellulose, esparto grass paper pulp, as an alternative nitrogen source with cellulose substrates. These combinations gave high cellulase yields. The greatest cellulase yields and productivity were obtained by fedbatch cultivation (23 filter-paper activity units (FPU)/mL and 168 FPU/L/h) [66]. Mutants of Penicillium echinulatum were isolated by treating conidia with hydrogen peroxide or 1,2,7,8-diepoxyoctane followed by incubating the conidia for 48 h in broth containing microcrystalline cellulose and 0.5% (w/v) aqueous to enrich the population grown in the presence of 2-deoxyglucose. These germinated conidia were washed and then plated onto cellulose agar containing 1.5% (w/v) glucose. The colonies showed the fastest production of halos on cellulose hydrolysis. This approach resulted in the isolation of two new cellulase-secreting P. echinulatum mutants: strain 9A02S1, showing increased cellulase secretion (2 IU/mL FPA) in submerged culture in agitated flasks containing a mineral salts medium and 1% cellulose, and strain 9A02D1, which proved more suitable for the production of cellulases in semisolid bran culture, in which it produced 23 IU of BGL per gram of wheat bran [67].

4.6.2

Genome Shuffling

Protoplast fusion is an important approach and has been widely used in fungal genetic modification since 1976, and it can induce DNA recombination between two strains. Genome shuffling is established on the basis of protoplast fusion, but it is actually the recursive fusion of multiple parents with the combination of a suitable screening method. Genome shuffling offers great potential for the improvement of industrially important microorganisms through protoplast fusion. Genome shuffling is a process that can efficiently combine the advantage of multiparental crossings with the recombination of entire


genomes, normally done with conventional breeding or through recursive protoplast fusion that greatly increases recombination compared to general protoplast fusion. Additionally, genome shuffling can accelerate directed evolution by facilitating recombination between members of a diversely selected population. This technique was first successfully used in a bacterial system [68] especially to improve acid tolerance in Lactobacillus sp. [69]. It has been widely applied in improving some important phenotypes of microorganisms, such as lipase production in Penicillium expansum, improvement of tylosin production in Streptomyces fradiae, acceleration of screening and breeding of high taxol-producing Nodulisporium sylviforme, pentachlorophenol degradation in Sphingobium chlorophenolicum, etc. Genome shuffling accelerates combination of the advantages distributed in multiparents. Thus, it is more efficient and saves lots of energy and time. Genome shuffling provides a new tool for cell and metabolic engineering and requires no sequence information or sophisticated genetic tools. Genome shuffling improves production of cellulase by Penicillium decumbens JU-A10. After two rounds of genome shuffling, three fusants, GS2-15, GS2-21, and GS2-22, were obtained, showing 100%, 109%, and 94% increase in cellulase activity compared to JU-A10, respectively. The cellulase production of the fusants on various substrates, such as corn stover, wheat straw, bagasse, and corncob residue, was studied. The maximum productivities of GS2-15, GS2-21, and GS2-22 were 92.15, 102.63, and 92.35 FPU/L/h on the corncob residue at 44 h, respectively, which were 117%, 142%, and 118% higher than that of JU-A10 (42.44 FPU/L/h, at 90 h). The improvements in cellulase production in the fusants could be due to their enhanced growth rates, earlier cellulase synthesis, and higher secretion of extracellular proteins [70].

4.6.3

Recombinant DNA/Gene Cloning

Using recombinant DNA technology, cloning the genes encoding the enzymes and heterologously expressing them in commonly used industrial strains has become a common practice. Such heterologous expression has become a powerful tool to improve yields and titers of enzymes. To develop a robust fungal strain producing enhanced levels of cellulases, many fungal cellulases have been cloned and expressed that produce higher titers of cellulase activities. Cre1 sites in the T. reesei genome and other fungal strains have been silent through several strategies such as RNA interference that resulted in increased cellulase activity. BGL from T. emersonii was expressed in T. reesei RUT-C30 using the strong cbh1 promoter, which resulted in expression of a highly thermostable BGL with high specific activity [71]. Zhang et al. [72] have developed a T. reesei strain by overexpression of BGL under the control of the cbh1 promoter. The resultant recombinants produce high levels of BGL and filter paper activity. Similarly, CBHI and CBHII were overexpressed using additional copies of the genes cloned under the cbh1 promoter. This resulted in a 1.5-fold increase in CBHI activity and 4-fold increase in CBHII expression [40]. The cbh1 strong promoter was employed to overexpress the cbh2 gene for enhancing CBH production in T. reesei because the CBHII component has higher


89

specific activity than CBHI and is an important component in cellulase. The recombinant plasmid pCAMBIA1300-hph-PsCT, containing a strong expression cassette, was constructed and transformed into T. reesei via optimized Agrobacterium-mediated transformation. Ten fast-growing T. reesei transformants were selected, among which C10 was found to have the highest filter paper activity, 28.92 2.45 IU/mL, 4.3-fold higher than that of ZU-02, 6.71 0.79 IU/mL. C10 also has the highest CBH activity, 122.44 7.42 U/mL, 5.4 times higher than that of ZU-02, 22.49 2.27 U/mL. The cellulase from C10 performed better (93.06 2.83%) than the one from ZU-02 in enzymatic hydrolysis because the exoeexo synergism played a role [73]. In addition, chimeric proteins have also been developed, for example, the EG from Acidothermus cellulolyticus was fused with T. reesei CBH and expressed in T. reesei. This bifunctional cellulase (endo and exo acting) has been demonstrated to improve saccharification yields [74]. Penicillium echinulatum is effective in bioconversion processes. However, nothing is known about the molecular biology of its cellulolytic system. Rubini et al. [75] described, for the first time, the isolation, cloning, and expression of a P. echinulatum cellulase cDNA (Pe-egl1) encoding a putative EG [75]. Degradation of cellulosic substrates requires enzymes that hydrolyze completely these substrates to their respective monomers. This will be possible only when new strains with high cellulase activity profiles are developed using system biology, recombinant DNA technology, synthetic biology, and metabolic engineering approaches. These strains/enzymes must be robust enough to tolerate extreme conditions employed during cellulose hydrolysis, which may reduce further the downstream cost. Ninety-nine percent of microbes are uncultivable and hence remain untapped for their potential applications [76]. These untapped sources could be exploited for isolating efficient cellulase producers with desirable properties. This could be possible using a metagenomics approach, which is considered to be the most viable method to search for desirable enzymes such as cellulases.

4.7 Cellulase Global Market Cellulases are currently the third largest industrial enzyme worldwide, by dollar volume, because of their use in cotton processing, in paper recycling, as detergent enzymes, in juice extraction, and as animal feed additives. However, cellulases will become the largest volume industrial enzyme if ethanol from lignocellulosic biomass through the enzymatic route becomes a major transportation fuel. The demand for cellulases is consistently on the rise because of its diverse applications. There are several companies involved in cellulase production for the textile detergent, paper, and other industries. Globally, there are two major players known for cellulase production for biomass conversiondGenencor and Novozymes. Both companies have played significant roles in bringing down the cost of cellulase several fold by their active research and are continuing to bring down the cost by adopting novel technologies. Now DuPont has


taken over Genencor. For the conversion of biomass feedstocks, DuPont has developed a market-leading enzyme solution. Accellerase TRIO is the first single enzyme product to deliver all major enzyme activities required for efficient biomass hydrolysis into both C5 and C6 sugarsdoften at half the dose of previous enzyme innovations [77]. Novozymes also has a diverse range of cellulase preparations available based on application, such as Cellusoft AP and Cellusoft CR for bioblasting in textile mills, Carezyme and Celluclean for laundry in detergent, Denimax for modifying the surface of the denim at low temperature, as well as many others specific for particular applications. Cellic CTec/Cellic HTec are among the enzyme families developed by Novozymes that are undergoing testing for various pretreated biomass saccharification [78]. Though no information is available on the source of production as well as availability in the market, these all contain different enzyme components required for efficient hydrolysis of different feedstocks. Amano Enzyme, Inc., in Japan and Advanced Enzymes in India are other enzyme industries actively involved in cellulase production. Table 4.4 shows the major players marketing cellulases with various trademarks and their source of origin, most of which may be genetically modified strains. Commercially, Table 4.4

Commercial Cellulases Produced by Companies and Their Sources

Enzyme Samples

Supplier

Source

Cellubrix (Celluclast) Novozymes 188 Cellulase 2000L Rohament CL Viscostar 150L Multifect CL Bio-Feed Beta L Energex L Ultraflo L Viscozyme L Cellulyve 50L GC 440 GC 880 Spezyme CP GC 220 Accelerase 1500 Cellulase AP30K Cellulase TRL Econase CE Cellulase TAP106 Biocellulase TRI Biocellulase A Ultra-Low Microbial (ULM) Accellerase TRIO

Novozymes (Denmark) Novozymes RhodiaeDanisco (Denmark) RohmeAB Enzymes (Rajamaki, Finland) Dyadic (Jupiter, FL) Genencor International (South San Francisco, CA) Novozymes Novozymes Novozymes Novozymes Lyven (Columbelles, France) GenencoreDanisco (Rochester, NY) Genencor Genencor Genencor Genencor Amano Enzyme (Troy, VA) Solvay Enzymes (Elkhart, IN) Alko-EDC (New York, NY) Amano Enzyme Quest International (Sarasota, FL) Quest International Iogen (Ottawa, ON, Canada) DuPonteGenencor

Trichoderma longibrachiatum and Aspergillus niger A. niger T. longibrachiatum/Trichoderma reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. longibrachiatum/T. reesei T. reesei A. niger T. reesei/T. longibrachiatum T. reesei/T. longibrachiatum Trichoderma viride T. reesei/T. longibrachiatum A. niger T. reesei/T. longibrachiatum T. reesei

Modified from Singhania RR, Sukumaran RK, Patel AK, Larroche C, Pandey A. Advancement and comparative profiles in the production technologies using solid-state and submerged fermentation for microbial cellulases. Enzyme and Microbial Technology 2010;46:541e9.


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the enzyme majors Genencor and Novozymes have launched a series of cocktails of cellulolytic enzymes for biomass hydrolysis, such as the Accellerase series of enzymes and the Cellic series of enzymes. The advanced enzyme preparations from both companies contain BGL supplements, indicating the importance of this enzyme in biomass hydrolysis. The cellulases available commercially and their sources have been listed in Table 4.4.

4.8 Protocol for Assay of Filter Paper Activity This protocol was adapted from the IUPAC protocol (Ghosh [79]). The following reagents and materials are needed. 1. Glucose stock: 10 mg/mL in 50 mM citrate buffer, pH 4.8 2. Sodium citrate buffer: 50 mM, pH 4.8 (may be prepared as 1 M, pH 4.5, and diluted) 3. 3,5-Dinitrosalicylic acid (DNS) reagent: prepare exactly as in the IUPAC protocol [79]. (DNS reagent needs DNS (Sigma Cat. No. D0550), phenol, sodium metabisulfite, and sodium potassium tartrate; we use all of these GR grade from Merck.) 4. Whatman No. 1 filter paper, 50-mg strips (1 6-cm strips normally give 50 mg)

4.8.1

Glucose Standard

Standards with varying concentrations of glucose are prepared. Standard: 0.5e3.0 mg S1

S2

S3

Glucose, 10 mg/mL stock (mL) 50 100 150 Sodium citrate buffer (mL) 1950 1900 1850 Glucose, absolute (mg) 0.5 1.0 1.5 Incubate for 60 min at 50 C. DNS reagent (mL) 3 3 3 Boil for 5 min, allow the reaction mix to cool rapidly in water Reaction mix (from above) 0.5 0.5 0.5 Distilled water 4.5 4.5 4.5 Take Reading at 540 nm. Absorbance (OD540)

S4

S5

S6

Blanks

200 1800 2.0

250 1750 2.5

300 1700 3.0

d 2000 0.00

3

3

3

3

0.5 4.5

0.5 4.5

0.5 4.5

0.5 4.5

Standard graphs are prepared by plotting the absolute glucose content in milligrams against optical densities using any spreadsheet program (we use MS Excel). Find the regression equation using the built-in function in Excel (manual plotting may not be accurate because it does not fit a regression line and rather tries only to join the points).


The equation for a line is obtained in the format y ¼ mx þ c, where m is the slope and c is the y intercept. So x (here the amount of glucose) can be calculated from the equation when y (OD540) is known.

4.8.2

Enzyme Assay

Appropriately diluted enzyme (0.5 mL) is incubated with 50 mg (w1 6 cm strip) of Whatman No. 1 filter paper for 1 h in a total volume of 2 mL. At the end of the reaction, 3 mL of DNS reagent is added to terminate the reaction mix. The reaction mix is boiled for 5 min for color development and allowed to cool in water. The unreacted paper is allowed to settle and 0.5 ml of the reaction mix is diluted 10 in distilled water and the absorbance is measured. Substrate Blank (SB) Whatman No. 1 filter paper (mg) 50 Sodium citrate buffer (mL) 2.0 Enzyme (appropriately diluted) (mL) ee Incubate for 60 min at 50 C. DNS reagent (mL) 3 Boil for 5 min, allow the reaction mix to cool in water Reaction mix (from above) 0.5 Distilled water 4.5 Absorbance (OD540)

Enzyme Blank (EB)

Test (T0 )

ee 1.5 0.5

50 1.5 0.5

3

3

0.5 4.5

0.5 4.5

The actual OD of the test sample is calculated as T ¼ T0 (SB þ EB). The absolute amount of glucose released is calculated from the equation as mentioned earlier. Enzyme concentrations (1/X, where X is the times of dilution, e.g., for a 10X; diluted sample, the enzyme concentration is 0.1) are plotted against the amount of glucose released by 0.5 mL of enzyme on a semilog plot. Enzyme concentration is plotted on a log axis and the glucose released on the normal axis. Here an exponential regression is fitted and the enzyme concentration at which 2.0 mg glucose is released is found by solving the regression equation. Once this dilution is obtained the FPU can be calculated as 0.37/concentration at which 2.0 mg glucose is released. An example is given in the following: Dilution (X)

Enzyme Conc. (1/X)

Glucose Released (mg)

5 10 20 40

0.200 0.100 0.050 0.025

2.187 1.765 1.234 0.876


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An example for the FPU calculation follows: 0.300

1.000

0

0.5

1

1.5

2.5

1

3

0.250 0.200

Enzyme Conc.

Enzyme Conc.

y = 0.0068e1.545x

0.200 0.150 0.100

0.100

0.050

R2 = 0.995

0.200 0.100 0.050

0.050

0.025

0.025 0.000

0

0.5

1

1.5

0.100

2

2.5

3

glucose released by 0.5ml

0.010

glucose released by 0.5ml

Plotted on normal Cartesian plane. Please note that when plotted on normal axes the linearity can be seen only around a glucose amount of 2 mg.

Plotted on semilog plot. Linearity is obtained over the whole range. A semilog plot is therefore used for calculations.

Here the enzyme concentration at which 2 mg glucose is released can be calculated based on the equation y ¼ 0:0068E1:545X

hence, in this case, y(enzyme conc.) ¼ 0.0068E1.545 2 ¼ 0.14944. So the FPU can be calculated as 0.37/0.14944 ¼ 2.48. This follows from the fact that 2 mg of glucose is released by 0.37 FPU of cellulase (please refer to Ghosh [79] for a detailed explanation). It may be noted that 1 mg of glucose is released by 0.185 FPU of cellulase, and in cases in which even undiluted enzyme fails to release 2 mg of glucose, FPU may be calculated by multiplying the amount of glucose released by 0.185. For example, if a sample releases only 1.5 mg of glucose under the assay conditions, the FPU are 1.5 0.185 ¼ 0.278. The validity of this calculation may be checked by the fact that if we multiply 2 by 0.185, the value obtained is 0.37, which is the FPU needed to release 2 mg of glucose.

4.9 Challenges for Enzymatic Biomass Conversion There are several challenges that have yet to be overcome, for example, the recalcitrance of lignocellulosic biomass, which necessitates the pretreatment step to open up the fibers and decrease the crystallinity of cellulose, which again adds to the cost of lignocellulosiceethanol technology. Pretreatment methods also need to vary from biomass to biomass based on their compositional characteristic. Second, the overall cost


of the technology of ethanol production from biomass is far from being economically feasible. For developing an economically feasible technology, the use of cheaper raw material as a substrate for cellulase production could bring down the production costs. SSF could be preferred over SmF for cellulase production for this particular application, as it is a cheaper technology that gives several fold higher production [24,31]. SSF imitates the natural habitat of the filamentous fungi, which could be a probable reason for better adaptability and higher production compared to SmF. Also, eliminating the steps in downstream processing of the enzyme for bioconversion might help to bring down the cost of cellulases, as would other approaches like improving the specific activity and temperature and low pH tolerance, as well as engineering the organism for improved production [80]. Most of the commercial cellulases available are produced from T. reesei and A. niger, but T. reesei lacks sufficient amounts of BGL to perform a proper and complete hydrolysis. Thus, the cellobiose accumulated from incomplete conversion caused by the limiting amounts of BGL inhibits exo- and endoglucanases. BGLs are also subject to product inhibition by the glucose beyond certain levels that vary between the different preparations and sources of the enzyme. One way to solve this issue is to add a glucose-tolerant BGL to the reaction mixture containing other cellulase components and to employ this cocktail for biomass hydrolysis, which would increase the efficiency of hydrolysis. There are several reports available in which an enzyme cocktail has been employed successfully for biomass conversion [81].

4.10 Future Perspectives The major goals for future cellulase research would be a reduction in the cost of cellulase production, which could be attained by (1) increasing the production level, (2) using cheaper raw materials as a substrate for production, (3) using alternative cheaper production technologies such as SSF, and (4) improving the efficiency of cellulases. The first task may include such measures as optimizing growth conditions or processes, whereas the improvement in cellulase efficiency requires directed efforts in protein engineering and microbial genetics to improve the properties of the enzymes. Optimization of growth conditions and processes has been attempted to a large extent to improve cellulase production. The section on fermentation production of cellulases describes many of these works basically dealing with empirical optimization of process variables to improve productivity. Many of the current commercial production technologies utilize SmF technology and employ hyperproducing mutants. Despite several efforts directed at generating hyperproducers by directed evolution, the cost of enzymes has remained high. Alternative strategies thought of in cellulase production include mainly SSF on lignocellulosic biomass, particularly by using host/substratespecific microorganisms. Filamentous fungi have been well exploited for the production of optimal enzyme complexes for the degradation of host lignocellulose, as SSF imitates their natural survival conditions rather than generating an artificial habitat. It is


95

also reported that the performance of enzyme complexes on lignocellulosic material is best when these complexes are prepared with the same lignocellulosic material as the host/substrate in fermentation. Another strategy is to use a mixed culture in the production of enzyme. Mixed culture gives improved production and enzyme complexes with better hydrolytic activity. Thus, among the other strategies tried in production optimization and process developments for cellulase enzyme production, SSF may be considered as a cost-effective means for large-scale production of cellulases that probably would be several fold cheaper compared to the current commercial preparations [80]. But SSF has its own limitations, as it is still not feasible to monitor regularly or to provide controlled conditions for the fermentation. Several large-scale SSF bioreactors have been engineered for cellulase production, which has been discussed in detail in a review [32] focusing on the direction of overcoming these limitations. Over several decades, the basic studies on cellulase have moved in the direction of understanding enzymatic diversity. There is now a vast and diverse understanding of the regulation of enzyme production, but still we lack comprehensive and specific knowledge on the mechanism of induction of cellulase by any of the known inducers. No information is available on the nature of intracellular inducers, the possible signaling pathways, and the cofactors and transducers involved in the induction of cellulase. Reports have shown that cellulases are subject to regulation by various factors and some of the cis-acting promoter elements have been characterized. Active research in this field has led to genetic improvement of cellulase production by various methods, including overexpressing cellulases from the cbh1 promoter of T. reesei and generation of desired variations in the cellulase production profile of organisms. The cbh1 and cbh2 promoters of T. reesei have also been exploited for expression of foreign proteins in Trichoderma. Feedback inhibition of cellulase biosynthesis by the end products, glucose and cellobiose generated by endogenous cellulolytic activity on the substrate, is another major problem encountered in cellulase production. Cellobiose is an extremely potent inhibitor of CBH and EG biosynthesis. Trichoderma and the other cellulase-producing microbes make very little BGL compared to other cellulolytic enzymes. The low amount of BGL results in a shortage of capacity to hydrolyze the cellobiose to glucose, resulting in a feedback inhibition of enzyme production and, in the case of biomass conversion applications, in the inhibition of cellulases. This issue has been addressed by various means like addition of exogenous BGLs to remove the cellobiose and engineering BGL genes into the organism so that it is overproduced. More and more research is oriented toward genetic manipulations of the cellulase producers for improving productivity. The developments in process design and medium formulations may be considered to have come to an age, and the future definitely requires controlled genetic interventions into the physiology of cellulase producers to improve production and thereby make the cellulase production process more costeffective. The major tasks ahead include overriding the feedback control by glucose and development of integrated bioprocesses for the production of cellulases.


Improvements in cellulase activities or imparting of desired features to enzymes by protein engineering are probably other areas in which cellulase research has to advance. Active-site modifications can be imparted through site-directed mutagenesis, and the mutant proteins can be used for understanding the mechanisms of action as well as for altering the substrate specificities or improving the activities. There are several reports of developments made in this direction. A mutant enzyme with EG-like features and activity improved by deleting the C-terminal loop of C. fimi CELB has been successfully generated [82]. Protein engineering has been successfully employed to improve the stability of a Humicola cellulase in the presence of detergents; to improve the thermostability of an alkaline, mesophilic endo-1,4-b-D-glucanase from an alkalophilic Bacillus sp.; and for altering the pH profile of CBH and EG from T. reesei. Such modifications affecting the enzyme properties may be beneficial in improving the overall performance of the cellulases and a better understanding of their mode of action, which will enable better utilization of the enzymes in biomass conversion. More basic research is needed in this direction to be able to make designer enzymes suited to specific applications in the future.

4.11 Conclusion After decades of research on lignocellulose utilization, it is now a consensus opinion that enzyme-based technologies for biomass conversion are the most efficient, cost-effective, and environmentally friendly. As of this writing, the cost of enzymes needed for biomass saccharification is the major hindrance to development of biomass conversion technologies. The leading enzyme companies have brought down the price of cellulases significantly. They have succeeded partly through developments in production technologies adopting multifaceted approaches such as using cheaper bioprocess technology, employing cheaper substrate, and employing engineered organisms and partly by development of artificial/engineered cellulases and cocktails of enzymes. Although the commercial lignocellulosic ethanol production has just begun in some parts of the world, continuous research is still needed to improve varied aspects of cellulase production (such as cost, specific activity, and substrate specificity) to achieve better technoeconomic feasibility. Artificial/engineered cellulases and enzyme cocktails rich in glucose-tolerant BGL have proved successful for increasing the rate or efficiency of hydrolysis of biomass so as to prove the technology economically feasible. Understanding of the microbial physiology and genetics of cellulase producers is still required. Whole-genome sequencing of the T. reesei genome is a major step in this direction. Similar efforts will be needed in the cases of other major cellulase producers also, so that more information is built up on the molecular biology of cellulase-producing fungi and their gene regulation. This information will be critical for future development of strains for cellulase production. With the current pace of research on cellulases, it can be asserted that more knowledge will be generated in the near future that will aid our progress toward a greener and sustainable carbohydrate-based economy.


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CURRENT DEVELOPMENTS IN BIOTECHNOLOGY AND BIOENGINEERING

[59] Mandels M, Weber J. The production of cellulases. In: Cellulases and their applications. Adv Chem Ser, 95; 1969. p. 391e414. [60] Mandels M, Weber J, Parizek R. Enhanced cellulase production by a mutant of Trichoderma viride. Applied Microbiology 1971;21:152e4. [61] Montenecourt BS, Eveleigh DE. Preparation of mutants of Trichoderma reesei with enhanced cellulase production. Applied and Environmental Microbiology 1977;34:777e82. [62] Montenecourt BS, Eveleigh DE. Selective screening methods for the isolation of high yielding cellulase mutants of Trichoderma reesei. In: Adv Chem Ser, vol. 81; 1979. p. 289e301. [63] Joglekar AV, Karanth NG. Studies on cellulase production by a mutant e Penicillium funiculosum uv-49. Biotechnology and Bioengineering 1984;26(9):1079e84. [64] Brown JA, Falconer DJ, Wood TM. Isolation and properties of mutants of the fungus Penicillium pinophilum with enhanced cellulase and b-glucosidase production. Enzyme and Microbial Technology 1987a;9(3):169e75. [65] Brown JA, Collin SA, Wood TM. Development of a medium for high cellulase, xylanase and b-glucosidase production by a mutant strain (NTG III/6) of the cellulolytic fungus Penicillium pinophilum. Enzyme and Microbial Technology 1987b;9(6):355e60. [66] Chaabouni SE, Belguith H, Hassairi I, M’Rad K, Ellouz R. Optimization of cellulase production by Penicillium occitanis. Applied Microbiology and Biotechnology 1995;43(2):267e9. [67] Dillon-Aldo JP, Zorgi C, Camassola M, Henriques JAP. Use of 2-deoxyglucose in liquid media for the selection of mutant strains of Penicillium echinulatum producing increased cellulase and b-glucosidase activities. Applied Microbiology and Biotechnology 2006;70(6):740e6. [68] Zhang YX, Perry K, Vinci VA, Powell K, Stemmer WP, del Cardayre SB. Genome shuffling leads to rapid phenotypic improvement in bacteria. Nature 2002;415:644e6. [69] Patnaik R, Louie S, Gavrilovic V, Perry K, Stemmer WP, Ryan CM, et al. Genome shuffling of Lactobacillus for improved acid tolerance. Nature Biotechnology 2002;20:707e12. [70] Cheng Y, Song X, Qin Y, Qu Y. Genome shuffling improves production of cellulase by Penicillium decumbens JU-A10. Journal of Applied Microbiology 2009;107:1837e46. [71] Murray P, Aro N, Collins C, Grassick A, Penttila M, Saloheimo M, et al. Expression in Trichoderma reesei and characterization of thermostable family 3 b-glucosidase from the moderately thermophilic fungus, Taleromyces emersonii. Protein Expression and Purification 2004;38:248e57. [72] Zhang J, Zhong Y, Zhao X, Wang T. Development of cellulolytic fungus T. reesei strain with enhanced b-glucosidase and filter paper activity using strong artificial cellobiohydrolase 1 promotor. Bioresource Technology 2010;101:9815e8. [73] Fang H, Xia L. High activity cellulase production by recombinant Trichoderma reesei ZU-02 with the enhanced cellobiohydrolase production. Bioresource Technology 2013;144:693e7. [74] Bower B, Larenas E, Mitchinson C. Exo-endocellulase fusion protein. 2005. Patent WO2005093073. [75] Rubini MR, Dillon AJP, Kyaw CM, Faria FP, Pocas-Fonseca MJ, Silva-Pereira I. Cloning, characterization and heterologous expression of the first Penicillium echinulatum cellulase gene. Journal of Applied Microbiology 2010;108:1187e98. [76] Singh BK. Exploring microbial diversity for biotechnology: the way forward. Trends in Biotechnology 2010;28:111e6. [77] http://biosciences.dupont.com/global-challenges/energy/dated 09/06/2015. [78] http://www.novozymes.com/en/about-us/brochures/Documents/Enzymes_at_work.pdf dated 09/ 06/2015. [79] Ghosh TK. Measurement of cellulase activities. Pure and Applied Chemistry 1987;59:257e68.


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[80] Singhania RR, Saini R, Adsul M, Saini JK, Mathur A, Tuli DK. An integrative process for bio-ethanol production employing SSF produced cellulase without extraction. Biochemical Engineering Journal 2015;102:45e8. [81] Adsul M, Sharma B, Singhania RR, Saini JK, Sharma A, Mathur A, et al. Blending of cellulolytic enzyme preparations from different fungal sources for improved cellulose hydrolysis by increasing synergism. RSC Advances 2014;4:44726e32. [82] Meinke A, Damude HG, Tomme P, Kwan E, Kilburn DG, Miller Jr RC, et al. Enhancement of the endobeta-1,4-glucanase activity of an exocellobiohydrolase by deletion of a surface loop. Journal of Biological Chemistry 1995;270:4383e6.


5

Industrial Enzymes: b-Glucosidases Reeta R. Singhania1, *, A.K. Patel1, R. Saini1, A. Pandey2 1 DBT- IOC C ENTRE FOR ADV ANCED BIOE N ER GY RE SEARCH, INDIANOIL C ORPORATION LIMITED; 2 CENTER OF INNOVATIVE AND APPLIED B IOPROCESSI NG, (A NATIONAL INSTITUTE UNDER DEPT OF BIOTECHNOLOGY, M INISTRY OF S &T , GOVT OF INDIA), MOHALI, PUNJAB , INDIA

5.1 Introduction b-Glucosidase (b-D-glucoside glucohydrolase; EC 3.2.1.21) (BGL) is ubiquitous in nature and can be found in bacteria, fungi, plants, and animals. BGL plays pivotal roles in many biological processes. The physiological roles associated with this enzyme are diverse and depend on the biological system in which they occur as well as the location of the enzyme. In cellulolytic microorganisms, BGL is involved in cellulase induction and cellulose hydrolysis [1]. In plants, the enzyme is involved in b-glucan synthesis during cell wall development, pigment metabolism, fruit ripening, cleavage of glycosylated flavonoids, and defense mechanisms [2], whereas in humans and other mammals, BGL is involved in the hydrolysis of glucosyl ceramides [3]. Defects in BGL activity in humans are associated with Gaucher disease, a nonneuropathic lysosomal storage disorder. These enzymes have broad substrate specificity and are used in a range of biotechnological processes, from liberating flavors, aromas, and isoflavone aglycones to the synthesis of oligosaccharides and alkylglycosides. The most intensively studied area of their application is the saccharification of cellulosic biomass for fuel ethanol production. Their synthetic activity in the production of oligosaccharides and arylglycosides is also a subject of intensive research. BGLs catalyze the hydrolysis of alkyl and aryl-b-glucosides as well as disaccharides and short-chain oligosaccharides. BGLs are well-characterized biologically important enzymes that catalyze the transfer of glycosyl groups between oxygen nucleophiles. The transfer reaction results in the hydrolysis of b-glucosidic linkages present between carbohydrate residues such as arylamino- or alkyl-b-D-glucosides, cyanogenic glucosides, short-chain oligosaccharides, and disaccharides under physiological conditions, whereas, under defined conditions, the synthesis of glycosyl bonds between two molecules can occur. Many BGLs also show synthetic activity via reverse hydrolysis, or transglycosylation [4]. Their synthetic activity in the production of oligosaccharides and arylglycosides is a subject of intensive research. They have been classified based on sequence as well as substrate specificity. *



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5.2 Classification of b-Glucosidases BGLs are a heterogeneous group of hydrolytic enzymes and have been classified according to various criteria. There is no single well-defined method for the classification of these versatile enzymes. In general, two methods for their classification appear in the literature, on the basis of (1) substrate specificity and (2) nucleotide sequence identity [5]. Based on substrate specificity, these enzymes have been classified as (1) aryl-BGLs, which act on arylglucosides; (2) true cellobiases, which hydrolyze cellobiose to release glucose; and (3) broad substrate specificity enzymes, which act on a wide spectrum of substrates. Most of the BGLs characterized so far are placed in the last category with various abilities for the cleavage of b-1,4; b-1,6; b-1,2; a-1,3; a-1,4; and a-1,6 glycosidic bonds. The most accepted method of classification of BGLs is by nucleotide sequence identity, proposed by Henrissat and Bairoch [5], which is based on the sequence and folding similarities (hydrophobic cluster analysis, HCA) of these enzymes. HCA of a variety of such enzymes suggests that the a-helices and b-strands are localized in similar positions in the folded conformation. Moreover, a number of highly conserved amino acids are also clustered near the active site. Such a classification is expected to reflect the structural features, evolutionary relationships, and catalytic mechanisms of these enzymes. Also, the identification of the nucleophile and the putative acidebase catalyst in one member of a family in effect identifies them in all the members of the family. It is also expected that as the size of the family increases, the residues conserved in all the members of the family usually will be important, structurally or catalytically. More sequence data and threedimensional structure of enzymes belonging to these families are required to confirm this scheme. The sequence-based classification is useful in characterizing the enzymes from the structural point of view but the substrate specificity with respect to the aglycone moiety still serves a primary or, in some cases, the only lead in isolating and characterizing the unknown or structurally undefined glucosidases. BGLs are mostly placed in either family 1 or family 3 of the glycosyl hydrolases, though these enzymes are also found in families 5, 9, and 30 [6,7]. Family 1 comprises nearly 62 BGLs from archaebacteria, plants, and mammals and also includes 6-phosphoglycosidases and thioglucosidases. Most family 1 enzymes also show significant b-galactosidase activity, whereas family 3 consists of b-glucosidases from bacteria, fungi, and plants. Families 1 and 3 include retaining enzymes that hydrolyze substrates with retention of an anomeric carbon via a double-displacement method. The family 1 BGLs are also classified as members of the 4/7 superfamily, with a common eightfold b/a-barrel motif. Here, the active site is placed in a wide cavity defined along the axis of the barrel, with a putative acidebase catalyst located at the end of b-strand 4 and a catalytic nucleophile near the end of b-strand 7 [8]. The 4/7 superfamily also includes other enzymes like family 2 b-galactosidases, family 5 cellulases, family 10 xylanases, and family 17 barley glucanases [9]. Family 3 of the glycosyl hydrolases (GH-3) consists of nearly 44 BGLs and hexosaminidases of bacterial, mold, and yeast origin. The structural data on the representatives

Chapter 5 Industrial Enzymes: b-Glucosidases 105

of GH-3 are still scarce, because only three of their structures are known, and only one of them has been thoroughly characterizeddthat of a b-D-glucan (exo-1 / 3, 1 / 4) glucanase from Hordeum vulgare, which catalyzes the hydrolysis of cell-wall polysaccharides. The enzyme consists of an N-terminal (a/b) eight-TIM barrel domain and a C-terminal domain of a six-stranded b-sandwich. The nonhomologous region, a helixlike strand of 16 amino acid residues, connects the two domains [4]. The catalytic center is located in the pocket at the interface of the two domains. Asp285 in the N-terminal domain acts as a catalytic nucleophile, whereas Glu491 in the C-terminal domain acts as a proton donor [10]. In general, BGLs cleave the b-1,4 glucosidic bonds in a variety of glucosides. Two carboxylic acids are involved in the catalysis at the active site. Protein engineering can be applied to increase the stability of BGLs. A classical example includes the work done by Nam et al., [11], in which they modified the N- and C-termini of BGL and provided an insight for increasing the stability of the protein.

5.3 Mechanism of Action BGLs are retaining enzymes because their products retain the same anomeric configuration as the substrate. Their reaction follows a double-displacement mechanism.

5.3.1

Hydrolysis

BGLs normally catalyze the hydrolysis of b-1,4 glucosidic bonds in aryl- and alkylb-D-glucosides from the nonreducing terminus. In the first step, the enzyme’s nucleophile in the active center attacks the substrate and an a-glycosyl enzyme intermediate is formed; in the second step, the intermediate is hydrolyzed by H2O and b-glucose is released as the product. BGL catalyzes the hydrolysis of glycosidic linkages formed between the hemiacetaleOH group of a cyclic aldose or between glucose and the eOH group of another compound, viz., sugar, amino-alcohol, aryl-alcohol, or primary, secondary, or tertiary alcohols. This reaction proceeds via the following steps: 1. During glycosylation, an enzymatic nucleophile attacks the anomeric (C1) center of the substrate glycoside, resulting in the formation of a covalently linked a-glycosyl enzyme intermediate, through an oxocarbenium ion-like transition state. Thus, the anomeric configuration at C1 is reversed. 2. Another active residue of the enzyme serves as the acidebase catalyst and donates a Hþ to the glycosidic oxygen, thereby assisting in the departure of the aglycone (or other glycone, as in disaccharides) group. 3. The glycosyleenzyme intermediate (2) is hydrolyzed via general base-catalyzed attack by the water at the anomeric center to release b-glucose as the product (3). The trans-addition of an eOH group results in the net retention of the b-anomeric configuration. The nucleophilic residue also acts as the leaving group in the deglycosylation step.

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The formation and hydrolysis of the enzymeeglycosyl intermediate occur via an oxocarbenium ion-like transition state.

5.3.2

Reverse Hydrolysis or Transglycosylation

The reactions for the biosynthesis of glycoconjugates occur either by reverse hydrolysis or by transglycosylation. In reverse hydrolysis, modification of reaction conditions such as lowering of water activity (aw), trapping of product, or high substrate concentration leads to a shift in the equilibrium of reaction toward synthesis. The two-step mechanism employed by retaining the enzymes such as BGLs allows these enzymes to transglycosylate. In reverse hydrolysis, the substrate (1) has an H in place of R. The enzymeeglycosyl intermediate is intercepted by R0 OH, where R0 is another sugar, yielding a disaccharide product (4). The reaction is under thermodynamic control. In the transglycosylation method, the substrate (1) has an R in place of H and is an “activated” donor, or a preformed donor glycoside (e.g., a disaccharide or aryl-linked glucoside) is first hydrolyzed by the enzyme, with the formation of an enzymee glycosyl intermediate. The enzymeeglycosyl intermediate may be trapped by a nucleophile other than water, viz., aryl- or alkyl-alcohol (as R0 OH), to yield a new glycoside. Here, the efficiency of the formation of the product is determined by the competition between the water and the acceptor R0 OH for the enzymeeglycosyl intermediate. This is then trapped by a nucleophile other than water (such as a monosaccharide; disaccharide; aryl-, amino-, or alkyl-alcohol; or monoterpene alcohol) to yield a new elongated product. The reaction is under the kinetic control. In the synthetic reactions, the reactive molecule in the second step is an R0 OH instead of H2O, yielding oligosaccharides or other glycosides. In reverse hydrolysis, the substrate is a sugar, mainly glucose, yielding a disaccharide product. In the transglycosylation process, formation of the product is the result of competition between water and the acceptor molecule. In many cases, lowering the water activity or a high substrate concentration would shift hydrolysis to transglycosylation. Little is known about the interaction of BGLs with their various substrates, especially with respect to the aglycone moiety, which forms the basis of tremendous diversity in terms of substrate range and is responsible for the subtle differences in substrate specificity. In most of the cases, several BGLs show high catalytic activity and high Km with artificial substrates such as para-nitrophenyl-b-D-glucopyranoside (pNPG) and methylumbelliferyl b-D-glucoside (MUG) but not with cellobiose. This is because cellobiose requires a conformational change for the catalysis as BGL has a very rigid structure in the S1 substrate binding site. The second glucose of cellobiose changes the conformation using the rotation of the s-bond of the glucoside so as to fit in the substrate binding site [11]. Thus, the glucose in the S10 pocket may determine the kinetic activity of the enzyme using steric hindrance, thereby affecting the substrate specificity. This could help in understanding the kinetics of BGL.


5.4 Sources of b-Glucosidases BGLs are widely distributed in the living world and play pivotal roles in many biological processes. They are ubiquitous in nature and can be found in bacteria, fungi, plants, and animals. The focus of this chapter, however, is limited to microbial sources only. BGLs, which are important for the terminal hydrolysis of complex polymers, are produced by a wide array of microorganisms and can be intracellular, extracellular, or membrane-bound. Intracellular BGLs are generally synthesized after exhaustion of the carbon source in the medium. Membrane-bound BGLs are, however, common in yeast. BGL from Aspergillus niger, which is generally used to complement the cellulolytic cocktail of Trichoderma reesei, is produced extracellularly, leading to its easy separation and purification, thus reducing the cost of downstream processing to a great extent. Microbial sources have been widely exploited for BGL production by both solid-state fermentation (SSF) and submerged fermentation (SmF). Filamentous fungi, among the microorganisms, are generally considered the most important sources of BGL. There are several reports of BGL production involving them, such as A. niger [12], Aspergillus oryzae [13], Penicillium brasilianum [14], Penicillium decumbens [15], Phanerochaete chrysosporium [16], Paecilomyces sp., [17], etc. There are also various reports of BGL production from yeasts (majority of them from Candida sp.) and a few bacteria. Even anaerobic bacteria such as Clostridium sp. are known to produce BGL along with other cellulolytic components. Table 5.1 shows the fungal strains involved in BGL production and the bioprocess for their production. Commercial production of BGL is often achieved by the use of species of Aspergillus. Aspergillus are known to produce higher titers of the enzyme [18]. Nevertheless, reports on the large-scale production of BGLs are limited in the public domain.

5.5 Production of b-Glucosidases As mentioned in Section 5.4, both the bioprocess technologies, i.e., SSF and SmF, are widely exploited for BGL production by microorganisms. Normally, all the cellulolytic microorganisms produce BGLs. Filamentous fungi, among the microorganisms, are generally considered the most important sources of BGL secreted outside the cell, and Aspergillus sp. is considered the best among these. The world market of enzymatic preparation represents a broad spectrum of cellulases, most of which are produced from T. reesei (Trichoderma longibrachiatum); however, there are also preparations from Penicillium and Aspergillus [19]. There are studies based on the comparison of cellulase activities but there are rare reports to compare BGL activities and properties with these fungal traits. Korotkova et al. [20] studied the properties of BGLs produced from Penicillium verruculosum, Aspergillus japonicus, and T. reesei for their possible employment in biomass saccharification. They presented the specific activity of BGL from these three fungi against different substrates at pH 5.0 and temperature 40 C. BGL from A. japonicus was a true cellobiase,

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Table 5.1

Production of b-Glucosidase and Its Titers, Properties, and Production Process BGL Units and Properties

Carbon Source

Production Process

Application

References

Debaryomyces pseudopolymorphus Trichoderma atroviride Penicillium pinophilum

Glucose tolerant, acidic, exocellular

Cellobiose

Shake-flask fermentation

[44]

2.5 IU/g

Pretreated willow Cellulose

Shake-flask fermentation

Enhancing wine aroma Biomass hydrolysis Cellobiose, cellotriose hydrolysis Cellulosic bioconversion

[23]

Rice bran

SmF, small fermenter with working volume 200 mL Solid-state fermentation

Cellulose

SmF

[46]

Wheat bran

SmF

Microcrystalline cellulose Cellulose

SmF

Biomass conversion Biomass (corncob) conversion Biomass hydrolysis

Penicillium citrinum

Periconia sp. Penicillium decumbens Penicillium echinulatum Stachybotrys sp. Humicola insolens Fomitopsis palustris

Fomitopsis sp.

Aspergillus niger Aspergillus unguis

83 U/mg protein, Vmax ¼ 1120 U/mg protein, Km ¼ 5.5 mM, Ki ¼ 26.6 mM, extracellular, acidic 159.1 U/g solid, Vmax ¼ 85.93 U/mg, Km ¼ 1.2 mM, Ki ¼ 17.9 mM with pNPG, thermoacidophilic Thermotolerant, active at wide pH range Thermotolerant, acidic BGL, Km ¼ 0.006 mM toward pNPG 0.85 U/mg of protein with pNPG

Fed-batch SmF

Km ¼ 0.22 mM toward pNPG and 2.22 mM for cellobiose, active at 50 C and pH 5 BGL active at 50 C and at pH 6.0, BGL activity was stimulated at 400 mM conc. Acidic BGL active at 55 C, Km ¼ 0.706 and 0.971 mM for pNPG of BGL1 and BGL2, respectively 53 U/g

Microcrystalline cellulose Cellobiose

Shake flask

Wheat bran

Solid-state fermentation

Thermostable, acidic, glucose resistant showing 92% activity retention at 250 mM glucose Glucose tolerant with 60% activity at 1.5 M glucose

Lactose, wheat bran Lactose, cellulose

BGL, b-glucosidase; pNPG, para-nitrophenyl-b-D-glucopyranoside; SmF, submerged fermentation.

Shake flask

Cellobiose hydrolysis Cellulosic biomass hydrolysis Cellobiose hydrolysis

[22]

[45]

[15] [47] [48] [49] [50]

[51]

SmF, shake flask

Rice straw and wheat straw hydrolysis Biomass hydrolysis

SmF, shake flask

Biomass hydrolysis

[18]

[21]


Microorganism


as the other two exhibited exo-1,3/1,4-BGL activity along with cellobiase activity. BGL from A. japonicus was superior based on its temperature stability and glucose inhibition constant. Relatively pure forms of cellulose and native as well as crude and pretreated biomass have been used successfully as carbon sources for production of BGL under both SmF and SSF (Table 5.1). It is also possible to obtain a BGL with desired properties by exploiting the conditions of fermentation as well as the nutrient source [21]. SSF has been often referred to as advantageous for the production of industrial enzymes [22,23]. In view of the findings that most BGLs expressed by Aspergillus in SmF remain adhered to the cell surface [24], SSF appears a better choice for BGL production. Process optimization is essential in improving productivity and to understand the effects of parameters on fermentation. In the conventional method for the optimization of enzyme production, the “one variable at a time” approach is used, which involves changing one parameter at a time while keeping all the other parameters constant [25,26]. The optimized concentration of the previous experiment is then incorporated into the next experiment. The same procedure is followed for all the parameters to complete the optimization [27]. But this process is cost, labor, and time intensive and also does not consider the interaction between variables. An alternative and more efficient approach is the use of statistical methods. Several statistical methods ranging from two-factorial to multifactorial designs are available [28]. Plackett and Burman designs [29] are fractional factorial designs used when one needs to screen a large number of factors to identify those that may be important (i.e., those that are related to the dependent variable of interest). In such situations a design that allows one to test the largest number of factor main effects with the smallest number of observations is desired. To enable this, the Plackett and Burman design has the interaction effects of variables confounded with new main effects. Because the added factors are created by equating (aliasing) the “new” factors with the interactions of a full factorial design, these designs will always have 2k runs, e.g., 4, 8, 16, 32, and so on. Full factorial design is fractionalized in a different manner, to yield saturated designs in which the number of runs is a multiple of 4, rather than a power of 2. In an experimental procedure for studying the effects of the process parameters (independent variables) under question, the selection of high (1) and low (1) values of the variable is very critical [25]. The difference between the levels of each variable must be large enough to ensure that the optimum response will be included. After the experiments are performed, the responses obtained are analyzed statistically to determine the effect of that variable on the response, the experimental errors, and the significance of the influence of each variable on the response [30]. The effect of a variable is the difference between the average responses of that variable at higher and lower levels. Probability tests are run to determine the level of significance of the effects of each variable. The design of experiments and analysis of responses are now routinely done using software made for the purpose, e.g., Statistica (Statsoft, Inc., USA), Design-Expert (Stat-Ease, USA), etc.

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Metagenomic approaches have proven successful in exploring the diversity of indicator genes, which are employed to test hypotheses about the structural composition and structure of soil microbial flora [31]. Molecular analysis of BGL diversity and its function in the soil revealed that only a narrow range of bacteria are capable of processing cellobiose in the soil [32]. The significant role of BGL produced by A. niger in degrading soil organic matter and the effects of soil minerals in enzymatic activity have been reported [33]. Several filamentous fungi exhibit the property of expressing different isoforms of BGL depending on the culture conditions or carbon source [21]. Various isoforms of BGL are expressed in response to carbon sources in Aspergillus terreus [34]. The sequential induction of isoforms has been associated with the presence of distinct metabolites [35]. As an accepted model, the induction of the cellulases is mediated either by low-molecular-weight soluble oligosaccharides that are released from the complex substrates as a result of hydrolysis by the constitutive enzymes or by the products (positional isomers) of transglycosylation reactions mediated by the constituent BGL [36]. These metabolites enter the cell and signal the presence of extracellular substrates and provide the stimulus for the accelerated synthesis of constituent enzymes of the cellulase complex. However, this process is complex in view of the fact that many fungi and bacteria are known to express functionally multiple cellulases/hemicellulases in the presence of different carbon sources. This multiplicity may be the result of genetic redundancy, differential mRNA processing, or posttranslational modification such as glycosylation, autoaggregation, and/or proteolytic digestion [37]. However, the regulation of expression of these multiple isoforms is still not clear, which necessitates further research regarding the sequential and differential expression of the isoforms. It must be emphasized that although the regulation of cellulases is apparently mediated through induction and repression as the two major mechanisms of controlling the expression of these enzymes, the highly specialized and complex nature of regulating the expression of cellulases in diverse microorganisms has also been reported [38,39]. There may be a relationship between the metabolites present in the culture extracts and the induction of various isoforms. Understanding the regulation would be important in designing the culture conditions for overproducing the desired kinds of isoforms or secondary metabolites. The structure and nature of the carbon source can also play an important role in differential induction of the enzyme system. Culturing under SmF or SSF also influences the expression of distinct isoforms. The multiplicity of BGL is well known in the case of filamentous fungi [21,34,40,41], and probably this multiplicity is essential, considering the vast and diverse roles these enzymes play in fungal metabolism and survival. BGL multiplicity can be attributed to the presence of multiple genes or to differential posttranscriptional modifications [42,43]. Differential expression of the various BGL proteins is reported in response to the carbon source supplied in the medium or the conditions of culture [21,34,40] and could be a probable adaptation of the fungi to the


changing immediate environment. This property, however, could be exploited for the selective expression of a desired isoform from a fungus by manipulating the culture conditions/carbon source carefully.

5.6 Assay of b-Glucosidases Little is known about the interactions of BGLs with their substrates, especially with respect to the aglycone moiety, which forms the basis of tremendous diversity in terms of substrate range and is responsible for subtle differences in substrate specificity. BGL activities are measured using artificial substrates such as pNPG or MUG. Few instances show BGL activities based on cellobiose substrate. In most of the cases, BGL shows high catalytic activity and high Km with artificial substrates such as pNPG and MUG but not with cellobiose. The kinetics of BGL depends on the configuration of its substrate, and cellobiose requires a conformational change for its catalysis. BGL has a very rigid structure in the S1 substrate binding site, which accommodates the glucose of cellobiose, but the second glucose of cellobiose changes the conformation, using rotation of the r-bond of the glucoside so as to fit in the substrate binding site [11]. It is not needed in the case of pNPG, in that nitrophenol follows the same binding patterns as glucose. This is the reason behind the low Kcat/Km of BGL toward cellobiose compared to the substrate pNPG. Hence, it is necessary to investigate the potentialities of BGL based on a natural substrate, as it needs to deal with cellobiose rather than pNPG or MUG in the natural process, which is the probable reason for not seeing a significant effect on the hydrolysis even after supplementation with heterologous BGL to the cellulase employed for hydrolysis of a biomass. It is necessary to mention again that the final evaluation of BGL should be made based on an assay in which cellobiose has been employed as substrate rather than pNPG derivatives or MUG.

5.6.1

b-Glucosidase Assay Using pNPG as Substrate

The BGL assay is done using 10 mM pNPG as substrate (in 0.05 M citrate buffer, pH 4.8). A standard graph is prepared using varying concentrations of p-nitrophenol (5e50 mM) in the same buffer. An appropriately diluted enzyme sample (0.5 mL) is incubated with 0.5 mL of substrate solution at 40 C for15 min. The reaction is stopped by adding 2.0 mL of 0.2 M Na2CO3. The activity of BGL is estimated spectrophotometrically by reading the absorbance of the liberated p-nitrophenol at 400 nm. One unit of BGL activity is defined as the micromoles of p-nitrophenol released per milliliter of enzyme per minute under standard assay conditions [21]. Alternatively, it is also done using 5 mM pNPG as substrate (in 0.05 M citrate buffer, pH 4.8). A standard graph is prepared using varying concentrations of p-nitrophenol (5e50 mM) in the same buffer. An aliquot of appropriately diluted enzyme (100 mL) is incubated with 1.0 mL of substrate solution at 50 C for 30 min. The reaction is stopped

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by adding 2.0 mL of 1% Na2CO3. The activity of BGL is estimated spectrophotometrically by reading the absorbance of the liberated p-nitrophenol at 400 nm. One unit of BGL activity is defined as the micromoles of p-nitrophenol released per milliliter of enzyme per minute under standard assay conditions.

5.6.2

b-Glucosidase Assay Using Cellobiose as Substrate

The cellobiase assay is done using 15.0 mM cellobiose as substrate in 0.05 M citrate buffer (pH 4.8). Fresh cellobiose solution should be prepared daily. For the assay, 1.0 mL of the crude enzyme sample, diluted in citrate buffer, is added to a small test tube. At least two dilutions must be made of each enzyme sample. One dilution should release slightly more and one slightly less than 1.0 mg (absolute amount) of glucose in the reaction conditions. After heating it to 50 C, 1.0 mL substrate solution is added and incubated at 50 C for 30 min. After the reaction is terminated by immersing the tube in boiling water for 5 min, the tube is placed in a cold water bath and the glucose produced is determined using standard procedures (e.g., using a kit based on the glucose oxidase reaction). Substrate or cellobiose blank (without enzyme) and enzyme blank (without substrate) absorbance should be subtracted from the sample values. One unit of cellobiase activity is defined as the micromoles of glucose released per milliliter of enzyme per minute under standard assay conditions. More details can be found in the work of Ghosh [52].

5.7 Strain Improvement Strain improvement for higher BGL titers as well as improved desired traits has been attempted by either mutation or genetic manipulation. Random mutations are a popular method but site-specific mutation is more promising as it causes changes at specific sequences. It involves prior knowledge about the primary structure of the protein so that a change can be made in a particular amino acid sequence.

5.7.1

Mutation

Random mutagenesis and selection is also called classical strain improvement or nonrecombinant strain improvement. In this method, there is a permanent alteration of one or more nucleotides at specific sites along the DNA strand. Mutation may be associated with a change in a single nucleotide (point mutation), through substitution or deletion, or rearrangement of one or more base pairs in a chromosome. In many cases, mutations are harmful but sometimes they create a strain that is more adaptable to its environment and improve its biocatalytic performance. The efficiency of random selection also depends on several other factors such as type of culture used (spores or conidia), mutagen dose and time of exposure, type of damage to DNA (deletion, addition, transversion, substitution of bases, or breakage of DNA), frequency of mutagen treatment, etc. [53].


A number of mutant strains overproducing BGL were isolated from the cellulolytic fungus Penicillium pinophilum 87160iii by UV mutagenesis [54]. Selection was carried out using either an agar plate or an enrichment technique. Improvements in BGL production were on the order of eight- to ninefold. The morphology of the mycelia of the mutants was quite different from that of the wild type. The mutants, for example, produced mycelia that were highly branched and thicker in cross section. In several of the mutants, synthesis of xylanase and BGL was completely derepressed in the presence of glycerol, which is a known repressor of the synthesis of these enzymes. Several mutants produced BGL enzyme that showed altered kinetics of hydrolysis in the presence of inhibitors [55]. A mutant of P. pinophilum produced BGL on the order of 35 units/mL. The yield of BGL by P. pinophilum was much higher than that of T. reesei [56]. It has been shown that the 184th residue of a novel BGL family 1 member could be responsible for its glucose tolerance, which was proved based on site-directed mutagenesis studies in which the mutant strain exhibited a better glucose tolerance level, with Ki 76.9 compared to 14.9 in the wild type [57].

5.7.2

Genetic Manipulation

The BGL genes from a large number of bacterial, mold, yeast, plant, and animal systems have been cloned and expressed in both Escherichia coli and eukaryotic hosts, such as Saccharomyces cerevisiae and filamentous fungi. Cloning has been performed by two methods, either by (1) the formation of a genomic DNA library, followed by the selection of recombinant clones by screening for BGL production, or (2) starting with a cDNA library (or a genomic library) and screening of recombinant clones with specific nucleotide probes designed from a priori knowledge of the polypeptide sequence [4]. Though fungi are known to be good producers of BGL, reports on the cloning of BGL from fungi are relatively low. This is mostly due to the complexities associated with the presence of introns in their genes and the complexities associated with glycosylation. Along with the ability to secrete the proteins, filamentous fungi can perform posttranslational modifications such as glycosylation and disulfidation [56]. Most of these are transformed with the plasmids, which integrate into their genome, and thereby provide stability to the fungal transformants. Thus, filamentous fungi have tremendous potential to be employed as hosts for recombinant DNA. The Aspergillus genus and T. reesei species have been used as hosts for expressing several genes of fungal as well as nonfungal origin. BGL genes have been cloned from fungi such as Talaromyces emersonii [58] and several Aspergillus spp. [59,60]. The majority of reports also mention the existence of multiple genes and gene products that are differentially expressed. Fungal genes have been cloned and expressed mostly in eukaryotic expression systems such as T. reesei [58,61], Aspergillus sp. [62], S. cerevisiae [59], and Pichia pastoris [59]. Pichia pastoris, because of its exceptionally high extracellular protein production capacity, has been an ideal gene expression system for industrial enzyme production and exploited for heterologous protein expression.

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Several GH-3 family BGL genes have been cloned and expressed successfully as active proteins but the amount of BGL produced always remained low [63e65]. Yeasts have been also well exploited both as a source of BGL genes and as host organisms to express the fungal BGL. Saccharomyces cerevisiae is the first choice to clone the gene for BGL as well as the genes for other components of the cellulases to bring simultaneous saccharification to reality [65]. However, here also comes the limitation of the expression of cellulolytic enzymes and conversion of biomass to glucose; thus the ethanol yield remains low even with potent ethanol producers from glucose. A trifunctional minicellulosome complex was generated that consisted of T. reesei endoglucanase 2 and cellobiohydrolase 2 and Aspergillus aculeatus BGL1 enzymes and was expressed in S. cerevisiae, which showed 8.8-fold enhanced activity because of both enzymeeenzyme and enzyme-proximity synergy [66]. This was a significant achievement to show consolidated bioprocessing for bioethanol production. The production of cellulosic ethanol has been demonstrated by expressing the heterologous endoglucanase and BGL genes from Clostridium thermocellum and Saccharomycopsis fibuligera, respectively, in S. cerevisiae [67]. A P. pastoris strain expressing a cDNA encoding BGL isolated from the buffalo rumen fungus Neocallimastix patriciarum was engineered for enhanced saccharification efficiency and was better than the commercial BGL, Novozym 188 [68]. Lee et al. [69] engineered an S. cerevisiae strain expressing a cellodextrin transporter and an intracellular BGL from Neurospora crassa. The genes responsible for expression of efficient and versatile BGL can be transferred to more industrially tractable and robust organisms to enhance the saccharification of the complex biomass polysaccharide. An industrial ethanol-producing strain of S. cerevisiae was constructed that could produce ethanol directly from microcrystalline cellulose by simultaneous saccharification with 88% of the theoretical yield of ethanol. The BGL2 gene from T. reesei along with an a-galactosidase gene was cloned into S. cerevisiae, which expressed both enzymes extracellularly [70]. A thermostable native BGL from Aspergillus fumigatus was cloned and expressed in P. pastoris X33 and was stable at 70 C and pH 4.0e7.0, which proved its suitability for lignocellulose hydrolysis [71]. Because T. reesei produces the other components in sufficient quantity, it would be of high value to engineer T. reesei with a BGL gene having desirable properties such as thermostability and acidic pH tolerance, which could produce sufficient amounts of BGL, leading to a better biomass hydrolysis rate. Engineering T. reesei in such a way that it produces the whole suite of saccharifying enzymes, including BGL, could be one of the approaches to reduce the processing cost of the hydrolysis of lignocellulosic biomass. While constructing a T. reesei strain with high BGL expression, it is necessary to employ a cellulose-inducible strong promoter. The cbh1 promoter has been widely used in eukaryotic systems for the expression of various proteins and has been extensively used for heterologous gene expression constructs in T. reesei [72]. However, the cbh1 and cbh2 (cel6a) promoters, which are known to be very strong, could reduce overall cellulase activity [73]. However, the xyn3 promoter to drive the expression of T. reesei BGL1


(Cel3a) could provide higher BGL activity than the parent strain without a significant decrease in total cellulase activity [74]. A recombinant T. reesei was constructed with the Aspergillus aculeateus BGL1 gene expressed under the control of the xyn1 promoter, which was capable of saccharification of pretreated cellulosic biomass. The recombinant strain expressed 25-fold higher BGL than the endogenous T. reesei BGL and 63-fold higher than that of the parent strain, A. aculeateus [57]. It would be interesting to have a hyper-BGL-producing gene with the desired properties cloned into a filamentous fungus that is already producing the other components of cellulase, which in turn will help to make the process economic. Table 5.2 shows the sources and hosts of BGL genes and also properties of the recombinant BGL.

Table 5.2 Source Organisms From Which b-Glucosidase Genes Have Been Cloned and Expressed into Host Organisms and Their Properties Source

Host Organism

Saccharomycopsis fibuligera Talaromyces emersonii Periconia sp.

Saccharomyces cerevisiae Trichoderma reesei Pichia pastoris

Penicillium decumbens Aspergillus niger Caldicellulosiruptor saccharolyticus Paecilomyces thermophila Periconia sp.

T. reesei

Neocallimastix patriciarum A. niger Chaetomium thermophilum Fervidobacterium islandicum Aspergillus aculeateus Aspergillus fumigatus Z5

T. reesei Escherichia coli P. pastoris

Properties of Recombinant BGL

References

1.02 IU/mg

[65]

GH family 3, thermostable active at 71.5 C, Vmax ¼ 512 IU/mg, Ki ¼ 0.254 mM against glucose Thermotolerant BGL, optimal activity at 70 C and at pH 5e7 Six- to eightfold increased BGL activity compared to native strain 5.3 IU/mL (106) times higher than native BGL Thermostable with 13 U/mg, having optimum activity at 70 C and pH 5.5 GH family 3, 274.4 U/mL, optimal at pH 6 and 60 C

[58] [46] [75] [72] [76] [63]

10.5-fold BGL activity increased from 2.2 to 23.9 IU/mg, thermotolerant and active in acidic pH GH family 3, 34.5 U/mg against cellobiose, optimally active at 40 C and pH 5.0 Specific activity of BGL increased by 22%

[77]

Optimally active at pH 5.0 and 60 C

[64]

E. coli

GH family 1, thermostable

[79]

T. reesei

10 U/mg against cellobiose

[57]

P. pastoris X33

Active at pH 6.0 and 60 C with specific activity of about 100 IU/mg

[71]

T. reesei QM9414 P. pastoris Penicillium verruculosum P. pastoris

BGL, b-glucosidase; GH, glycosyl hydrolase.

[68] [78]

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5.8 Applications of b-Glucosidases BGLs have broad substrate specificity and are used in a range of biotechnological processes. They play important roles in nature, including the degradation of cellulosic biomass by fungi and bacteria, breakdown of glycolipids in mammalian lysosomes, and cleavage of glycosylated flavonoids in plants.

5.8.1

Applications Based on Hydrolysis

BGLs as part of the cellulase enzyme complex hydrolyze cellobiose and cellooligosaccharides to yield glucose, which is fermented by yeasts into bioethanol. The conversion of cellulosic biomass into fermentable sugars is a very attractive method of bioethanol production, as discussed in more detail in Section 5.8.3. Using BGLs as additives in cellulose-based feeds is beneficial for single-stomach animals, such as pigs and chickens, by enhancing the digestibility of the feed. In winemaking, BGLs play a key role in the enzymatic release of aromatic compounds from glycosidic precursors present in fruit juices, nuts, and wines. The natural process by endogenous plant BGLs is very time consuming. Supplementation with external enzymes may enhance aroma release. In tea beverages treated with immobilized BGL, the essential oil content increases by 6.79e20.69% [80]. In citrus fruit juices, the hydrolysis of naringenin to prunin reduces the bitterness of the juices [81]. Isoflavones found in soybean have phytoestrogenic properties, so they can relieve menopausal symptoms and can help prevent several chronic diseases and certain cancers. However, in soy-based foods, the isoflavones are mainly in the inactive form of glycosides. Production of aglycones by the hydrolysis via BGLs is highly desired [82]. Enrichment of genistein, the most potent inhibitor of cancer cell growth, in soy protein concentrate has been reported [83]. In soy milk, the aglycone content is increased significantly either by treatment with BGL or by fermentation with a BGL-producing Lactobacillus strain [84]. Detoxification of cassava (a staple food in tropical countries) can be achieved by the addition of linamarase (BGL) of Mucor circinelloides. After 24 h of fermentation, all cyanogenic glycosides are hydrolyzed to remove the toxic eCN moiety [85]. SSF of cranberry and pineapple pomace with good BGL-producing fungal strains (Lentinus edodes and Rhizopus oligosporus) enhances the amount of extractable free phenolics, showing antioxidant activity [86,87].

5.8.2

Applications Based on Transglycosylation

The synthetic activity of BGLs is used for the biosynthesis of oligosaccharides and alkylglycosides. Oligosaccharides can be used as therapeutic agents, diagnostic tools, and growth-promoting agents for probiotic bacteria. They have important functions in biological systems including fertilization, embryogenesis, and cell proliferation. Galactooligosaccharides, the transgalactosylation products from lactose, are good


growth factors for intestinal Bifidobacteria. Alkylglycosides are nonionic surfactants with high biodegradability, and have good antimicrobial properties. Hence, they have potential application in pharmaceutical, chemical, cosmetic, food, and detergent industries [88]. Their enzymatic synthesis using the transglycosylation activity of glycoside hydrolases can be performed in one step, instead of the several protectionedeprotection steps required in chemical synthesis. There are some examples of the use of the synthetic activity of BGLs in the literature. Galactooligosaccharides and isobutylgalactosides were synthesized from lactose in watereorganic medium via A. oryzae BGL [88]. Arbutin-b-glycosides were synthesized via the transglycosylation reaction of Thermotoga neapolitana BGL to develop a new skin-whitening agent, and the products were evaluated for their melanogenesisinhibitory activities [89]. Naringin and naringenin have diverse biological activities, including antibacterial, antioxidant, antiinflammatory, antifibrotic, and antiulcer activities. Naringin is a flavanone responsible for bitter taste. In the citrus juice industry, to improve the quality and the commercial value of juice, naringin should be converted to naringenin (Fig. 5.1). The flavanone aglycones hesperetin and naringenin are produced by hydrolyzing the sugar moieties in the flavanone glycosides hesperidin, naringin, neohesperidin, and narirutin. These flavanone aglycones have been obtained by the hydrolysis reactions of microorganisms, including A. niger and other fungal and yeast cells.

5.8.3

Role of BGL in Biomass Conversion

With the advent of the need for alternative energy, there is a resurgence in bioethanol production from lignocellulosic biomass through the enzymatic route. BGL has received immense attention, being the key enzyme in complete cellulosic hydrolysis. The cellulolytic enzyme system secreted by the filamentous fungus T. reesei is the one most used in industrial applications. The hydrolysis step converting cellulose to glucose is recognized as the major limiting operation in the development of processes for the production of bioethanol from lignocellulosic raw materials because of the low efficiency of cellulases and their high cost. This issue has been addressed in various ways, like supplementation of exogenous BGL to the T. reesei cellulase to remove the cellobiose, or engineering BGL genes into the host organism so as to overproduce it, which will enable improvement in the efficiency of biomass hydrolysis and cost reduction of biomass-to-bioethanol conversion by reducing feedback inhibition and cellobiose-mediated repression of cellulases. Enzymatic hydrolysis of cellulose is a multistep complex process, the last step being a homogeneous catalysis reaction involving the action of BGL on cellobiose [90]. Cellobiose is a strong inhibitor of both cellobiohydrolases and endocellulases, and the BGL action can reduce its effect. In addition, the glucose produced also inhibits cellulose hydrolysis and exerts feedback inhibition. Glucose at high concentrations can either block the active site for the substrate or prevent the hydrolyzed substrate from leaving [14]. The amount of BGL1

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FIGURE 5.1 (A) Conversion of hesperidin plus naringin and neohesperidin plus narirutin to hesperetin plus naringenin, respectively, by b-glucosidase from Pyrococcus furiosus. (B) Hydrolytic pathways from hesperidin to hesperetin by two enzyme reactions with a-rhamnosidase plus b-glucosidase and one enzyme reaction with P. furiosus b-glucosidase. Two hydrolytic pathways are shown: hesperidin / hesperetin-7-O-glucoside / hesperetin and hesperidin / hesperetin.


generated by T. reesei hyperproducing strains represents a very low percentage of the total secreted proteins [90,91]. Although 12 BGLs have been identified in the T. reesei genome, most are intracellular. Most of the cellulase-producer filamentous fungi are characterized by low secretion of BGL [90,92], which indicates the activity will be insufficient to convert cellobiose (an intermediate product in cellulose hydrolysis) to glucose. The low abundance of BGL, even under conditions of cellulase induction, and the product inhibition to which it is susceptible limit the use of native cellulase preparations in lignocellulosic biomass hydrolysis for alcohol production. Alternative strategies such as cocultivation of fungal strains producing cellulase and BGL, like T. reesei and an Aspergillus strain such as A. phoenicis or A. niger, were used for better cellulase with increased activity of BGL [93]. This limitation can be alleviated either by overexpressing BGL in T. reesei or by adding extra BGL from other sources [94]. Supplementing the native T. reesei enzymatic cocktail with BGL from other fungi is also often performed to avoid inhibition of cellobiose [95]. An enzyme cocktail prepared from cellulase-producing T. reesei and an extracellular BGL-producing mutant strain of Trichoderma atroviride was better than a commercial preparation for enzymatic hydrolysis as well as for the simultaneous saccharification and fermentation of pretreated spruce [96]. BGLs with high hydrolyzing activity, heat and glucose tolerance, acid resistance, and possible transglycosylase activity are in demand. There are other reports also that show enzyme preparations from different fungal strains perform even better than commercial preparations such as Accellerase 1000, Novozym 188, and Celluclast 1.5L [96]. Commercially, the enzyme majors Genencor and Novozymes have launched a series of cocktails of cellulolytic enzymes for biomass hydrolysis, such as the Accellerase series of enzymes [97] and the Cellic series of enzymes [98]. The advanced enzyme preparations from both companies probably are from genetically modified T. reesei with high BGL activity. Although detailed information on the composition of T. reesei cellulase is not disclosed, it is assumed to contain heterologously expressed BGL from Aspergillus sp.

5.9 Conclusions and Perspectives BGL is a key enzyme involved in the sugareenzyme platform for bioethanol production from lignocellulosic biomass. In the present scenario, the commercialization of this platform is limited by various technological challenges, which include a slow enzymatic degradation rate and feedback inhibition of the enzyme, mainly BGL. Development of glucose-tolerant BGL and supplementing it with cellulases from other sources, e.g., T. reesei, could offer potential benefits in overcoming feedback inhibition, leading to high reaction rates, and also help in high solids processing. Heterologously expressed BGL from genetically modified host organisms could also be an effective tool in developing enzymes with desired properties.

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Industrial Enzymes: Xylanases

6

L. Thomas1, *, A. Joseph2, Reeta R. Singhania3, A.K. Patel3, A. Pandey4 1

CSIR-NATIONAL INSTITUTE F OR INTERDISCIPLINARY SCIENCE AND TECHNOLOGY (NIIST), TRIVANDRUM, INDIA; 2 KANNUR UNIVERSITY, KANNUR, INDIA; 3 DBT- IOC C ENTRE FOR AD V ANC E D B IOENER GY R ESE ARC H, I NDIANOIL CORPORATION LIMITED; 4 CENTER OF INNOVATIVE AND APPLIED BIOPROCESSING, (A NATIONAL INSTITUTE UNDER DEPT OF BIOTECHNOLOGY, MINISTRY OF S&T, GOVT OF INDIA), MOHALI, PUNJAB , IND IA

6.1 Introduction Xylanases are hydrolytic enzymes comprising endo-1,4-b-xylanase (EC 3.2.1.8) and b-xylosidase (EC 3.2.1.37). They hydrolyze xylan (present in hemicelluloses of plants) to monomeric sugars along with assisting hydrolytic enzymes such as a-L-arabinofuranosidase, a-glucuronidase, acetylxylan esterase, and phenolic acid (ferulic and p-coumaric acid) esterase [1]. Of these, endoxylanase (EC 3.2.1.8, often referred as xylanase), which catalyzes the hydrolysis of b-1,4 linkages in xylan, is particularly important, as it breaks the main backbone of xylan for hydrolysis. The role of assisting enzymes comes into play depending upon the nature of the side chain with which the xylan is associated. These side chains help in the complete breakdown of xylan to its monomer, xylose. Xylanases were first reported in 1955 and were originally termed as pentosanases. They were recognized by the International Union of Biochemistry and Molecular Biology in 1961 and were assigned the enzyme code EC 3.2.1.8. They have been referred to by various names. Their commonly used synonymous terms include xylanase, endoxylanase, endo-1,4-b-D-xylanase, b-1,4-xylanase, and b-xylanase, but the official name is endo-1,4-b-xylanase. Based on amino acid sequence similarities and hydrophobic cluster analysis [2], xylanases fall mainly into two glycosyl hydrolase families, family 10 (>30 kDa with low pI values) and family 11 (